Downy mildew on oilseed Brassicas – understanding the drivers of disease epidemics and potential of novel
host resistances
Akeel Emad Mohammed
Bachelor of Agricultural Science, Master of Plant Protection
This thesis is presented for the degree of Doctor of Philosophy of
The University of Western Australia
School of Agriculture and Environment
Plant Pathology
2018
THESIS DECLARATION
I, Akeel Emad Mohammed, certify that:
This thesis has been substantially accomplished during enrolment in the degree.
This thesis does not contain material which has been accepted for the award of any other degree or diploma in my name, in any university or other tertiary institution.
No part of this work will, in the future, be used in a submission in my name, for any other degree or diploma in any university or other tertiary institution without the prior approval of The University of Western Australia and where applicable, any partner institution responsible for the joint-award of this degree.
This thesis does not contain any material previously published or written by another person, except where due reference has been made in the text.
The work(s) are not in any way a violation or infringement of any copyright, trademark, patent, or other rights whatsoever of any person.
This thesis contains published work and/or work prepared for publication, some of which has been co-authored.
Signature:
Date: 10/04/2018
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ABSTRACT
Downy mildew disease caused by (Hyaloperonospora brassicae syn. H. parasitica) is a major disease limitation to oilseed Brassica production
(particularly rapeseed, canola, mustard) worldwide and also causes significant damage in vegetable Brassica crops. Infection can be high as 100% on some
Brassica crops especially in the early seedling stage. Up to date, no Brassica varieties have been specifically developed with effective resistance to this important disease.
Screening for the host resistance to H. brassicae of 131 Brassicaceae varieties at the cotyledon stage, including 109 Australian canola varieties
(Brassica napus and B. juncea) and 22 diverse Brassicaceae (including B. napus,
B. carinata, B. juncea, B. nigra, B. rapa, Crambe abyssinica and Raphanus sativus) highlighted new excellent resistance to downy mildew. Using a mixture of 10 H. brassicae isolates collected from southern Australia areas severely affected by downy mildew disease in 2015, new high level resistances were identified across R. sativus, B. carinata, B. napus, B. juncea and C. abyssinica.
Cluster analysis revealed six distinct clusters (highly resistant, resistant, moderately resistant, moderately susceptible, susceptible, and very susceptible) based on disease index (%DI) values, and this opens the way for breeders having to only select a single genotype from within each of the clusters determined as highly resistant or resistant in developing new resistant commercial varieties. This is the first study to demonstrate the existence of these very high levels of pathotype-independent resistance to H. brassicae, particularly in Australian canola varieties.
The above studies were followed up with additional search for further new sources of resistance to H. brassicae, but this time across more diverse iii
Brassicaceae including 78 B. napus, 38 B. carinata, 25 B. juncea and 13 miscellaneous Brassicaceae including (three of Raphanus sativus, two of each of Rapistrum rugosum, B. incana and one each of C. abyssinica, B. fruticulosa,
Hirschfeldia incana, B. insularis, B. oleracea and Sinapis arvensis. Further new sources of effective resistance were identified to H. brassicae among these, particularly in R. sativus; B. carinata, B. juncea B. carinata, B. incana and C. abyssinica.
To examine the role of environmental factors on the development of downy mildew epidemics, the effects of temperature (14/10⁰C and 22/17⁰C day/night) and plant age (15, 23, 31 and 40 day-old-plants) on the severity of downy mildew on B. juncea and B. napus varieties were determined. There were significant effects of temperature, plant age and their interaction, with more severe disease under warmer conditions and on very young seedlings. Findings explain the recent increase in severe disease epidemics in canola as seasonal temperatures increase and why most severe epidemics are on youngest plants.
H. brassicae isolates collected 2006-2008, and more recently, were inoculated onto cotyledons of 28 diverse Brassicaceae genotypes to identify and select suitable Brassica spp. differentials to enable characterisation of H. brassicae pathotypes and to define phylogenetic relationships among isolates across Australia. Using octal classification, the six Brassicaceae host genotypes most suitable as host differentials to characterise pathotypes of H. brassicae were identified and then used to define eight distinct pathotypes occurring in Australia.
Phylogenetic relationships, determined across 20 H. brassicae isolates collected in 2006-2008 and 88 isolates collected in 2015-2016, highlighted seven distinct clades. These are the first studies to define the phylogenetic relationships and pathotype structure among H. brassicae isolates in Australia and set a
iv benchmark for understanding current and future genetic and phenotypic pathotype shifts within pathogen populations.
In summary, these studies are the first to determine the expression of pathotype-independent host resistances across an extensive set of canola varieties and diverse Brassicaceae to H. brassicae isolates. Highly resistant varieties identified have resistance(s) that can, in some instances, be directly deployed into downy mildew prone regions and, in all instances, provide resistances for use in Brassicaceae breeding programs. Studies of the effect of temperature and plant age explain, for the first time, why the incidence and severity of downy mildew is most severe on seedlings and why both have increased over the past decade and a half when season temperatures have been increasing as a consequence of climate change. The new set of Brassicaceae host differentials developed provides the first practical means for monitoring changes in pathotype structure within H. brassicae populations. Further, these host differentials not only allow early warning of new pathotypes able to overcome current and future host resistances, but provide breeders with the relevant pathotype information needed to develop and deploy appropriate and effective host resistances to counter the inevitable changes in pathotypes of H. brassicae over coming decades.
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TABLE OF CONTENTS
Thesis declaration……………………………………………………………………...ii
Abstract……………………………………………………………………….………..iii
Table of contents…...... vi
Acknowledgements ………………….…….…………………………………….….viii
Authorship declaration: Co-authored publications…………..……….………..…..ix
Chapter 1: Literature review …………………………………………………………1
1.1 Introduction……………………………………………………..……………….…2
1.2 Brassicas (particularly oilseed brassicas rapeseed, canola, mustard)...……3
1.3 Oilseed brassica downy mildew disease……………………………………….4
1.3. 1 Classification of Hyaloperonospora brassicae……………………….…4
1.3. 2 Life cycle of Hyaloperonospora brassicae…………………...………….5
1.3. 3 Symptoms of downy mildew on brassicas……………………...……….6
1.3. 4 Host range of Hyaloperonospora brassicae on cruciferous species....7
1.3. 5 Downy mildew on canola and mustard………………………….………8
1.3. 6 Severity and impact of downy mildews on canola and mustard………8
1.3. 7 Physiological specialisation within Hyaloperonospora brassicae
populations………………………………………………………………………...9
1.3. 8 Impact of environmental factors on development of downy
mildew…………………………………………………………………………….11
1.3. 9 The effect of plant age on downy mildew epidemics …………….…..12
1.3. 10 Management of downy mildew disease…………………………..….13
1.4 Sources of resistance in oilseed brassicas and other brassicas……………14
1.5 Concept and aims of work………………………………………………………16
Bibliography for literature review……………………………………….…………..19
vi
Chapter 2: New resistances offer opportunity for effective management of the downy mildew (Hyaloperonospora parasitica) threat to canola………..…..……35
Chapter 3: Resistances to downy mildew (Hyaloperonospora brassicae) in diverse Brassicaceae offer new disease management opportunities for oilseed and vegetable crucifer industries………………………………………………..….45
Chapter 4: Temperature and plant age drive downy mildew disease epidemics on oilseed Brassica napus and B. juncea…………………………………………80
Chapter 5: Pathotypes and phylogenetic variation determine downy mildew epidemics in Brassica spp. in Australia……………………………………………90
Chapter 6: General Discussion…………………………………………....………105
Bibliography for general discussion………………………………...…….………113
Appendix- Presentations-Proceedings………………………………...…………118
vii
ACKNOWLEDGEMENTS
First of all, I would like to acknowledge my supervisors Professor Martin Barbetti and Dr Ming Pei You for their efforts, guidance and continuous support during my
PhD journey. It is a wonderful opportunity to complete my study under the supervision of a great pathologist and respectful scientist, who treats his students respectfully to achieve their goals.
A huge thank to my family, my mother, father, my lovely wife and my dear sons
(Emad, Alaa and Karar) for their support and their patience in the difficult circumstances of my study.
I am grateful to my colleagues, Hebba, Margaret, Dolar and Solomon for their assistance and encouragement. My friends in Iraq, who support me by their sincere wishes are deeply appreciated.
I would also like to thank University of Kufa in Iraq for awarding me a scholarship to complete my studies in Australia.
I am grateful to the financial support of the Grains Research and Development
Corporation (GRDC) as this work constitutes part of the GRDC UWA 170 project
“Emerging foliar diseases of canola”. Operational Funds from the School of
Agriculture and Environment, The University of Western Australia are acknowledged.
Finally, I would like to acknowledge the technical support from Robert Creasy and
Bill Piasini in the University of Western Australia Plant Growth Facilities. viii
AUTHORSHIP DECLARATION: CO-AUTHORED PUBLICATIONS
This thesis contains work that has been published.
Details of the work:
Mohammed AE, You MP, Barbetti MJ, 2017. New resistances offer
opportunity for effective management of the downy mildew
(Hyaloperonospora parasitica) threat to canola. Crop and Pasture Science
68, 234–242.
Location in thesis: Chapter 2
Student contribution to work:
I maintained H. brassicae isolates on susceptible B. napus cultivars,
planted test genotypes, prepared the inoculum, did the pathogenicity
testing, collected and recorded the data, analysed the data and drafted this
paper.
Co-author signatures and dates:
Mingpei You Martin Barbetti
10/ 04/2018 10 /04/2018
Details of the work:
Mohammed AE, You MP, Banga SS, Barbetti MJ, 2018. Resistances to
downy mildew (Hyaloperonospora brassicae) in diverse Brassicaceae offer
new disease management opportunities for oilseed and vegetable crucifer
ix
I prepared and planted test varieties, inoculated cultivars within the mixture of ten H. brassicae isolates in different temperatures and plant ages, collected the data and drafted this paper.
Co-author signatures and dates:
Mingpei You Martin Barbetti
10/04/2018 10/04/2018
Details of the work:
Mohammed AE, You MP, Barbetti MJ, 2018. Pathotypes and phylogenetic variation determine downy mildew epidemics in Brassica spp. in Australia.
Plant Pathology. 67, 1514–1527. (Online at: Doi:10.1111/ppa.12861)
Location in thesis: Chapter 5
Student contribution to work:
I prepared and planted the studied cultivars, extracted the DNA from downy mildew isolates, inoculated cultivars within each H. brassicae isolate separately, determined genotype x isolate interactions, collected the data, and drafted this paper.
xi
Co-author signatures and dates:
Mingpei You Hebba Al-lami
10/04/2018 10/04/2018
Martin Barbetti 10/04/2018
Student signature:
Date: 10/04/2018
I, Professor Martin Barbetti certify that the student statements regarding their contribution to each of the works listed above are correct.
Coordinating supervisor signature:
Date: 10/04/2018
xii
Chapter 1
Literature review
Page 1 of 119 Chapter 1: Literature review
1.1 Introduction
Brassicas are a diverse group of plants that include oilseed crops that are economically important worldwide (Warwick et al., 2009) such as Brassica napus
(rapeseed), B. juncea (Indian mustard), B. carinata (Abyssinian/Ethiopian mustard) and B. rapa (turnip rape; syn. Brassica campestris) (Rich 1991; Dixon
2007; Snowdon et al., 2007). They also include a range of different leaf and root vegetables that together have a wide range of different nutritional and industrial uses (Tsunoda et al., 1980). The value of the canola crop in Australia ranks third after wheat and barley and the canola area grown has nearly doubled from 1.4 million ha in 2009 to 2.4 million ha in 2013 (Elliott et al., 2015) and more recently has increased further. There is also interest in replacing diesel fuel with biofuel produced from oilseeds and interest in developing such alternative crops has increased recently in Western Australia (Rustandi & Wu 2010). However, many diverse and significant diseases infect oilseed Brassicas (particularly rapeseed, canola, mustard) worldwide, but particularly in Australia (Delourme et al., 2011;
Salisbury & Barbetti 2011; Barbetti et al., 2012). These include blackleg caused by (Leptosphaeria maculans and L. biglobosa), sclerotinia rot (Sclerotinia sclerotiorum) that are particularly devastating (Delourme et al., 2011) but there are also several other important diseases including white leaf spot
(Pseudocercosporella capsellae), Alternaria blight (Alternaria spp.), powdery mildew (Erysiphe cruciferarum (Uloth et al., 2016) and downy mildew
[Hyaloperonospora brassicae (syn. H. parasitica)].
H. brassicae has become an increasingly widespread and severe pathogen over recent decades across canola growing areas in Australia (Barbetti
Page 2 of 119 2000). Its importance on canola crops has been demonstrated clearly in Western
Australia after the destructive outbreaks of this disease in recent years, especially when it occurs at the early seedling stage (Ge et al., 2008). While downy mildew disease most adversely impacts on young Brassica seedlings, it also can reduce the productivity and quality even when plants are infected at a later stage of growth (Silué et al., 1996; Coelho et al., 2012). A survey in 2004 (Oilseeds
Industry Association of Western Australia, unpublished), estimated the economic losses from downy mildew on canola to be as much as AU$13 million annually in
Western Australia alone.
1.2 Oilseed Brassicas (particularly rapeseed, canola, mustard)
Brassicaceae contains 338 genera and 3709 species (Warwick et al.,
2006). Some oilseed species such as Brassica napus, B. juncea, B. carinata and
B. rapa have enormous commercial value worldwide (Rich 1991). Canola is a rich source of oil for human consumption and the meal, containing a high concentration of protein, and is important in a range of animal feeds (Si et al.,
2003). Canola represents a significant source of vegetable oil internationally and ranks third among oilseed crops (Ashraf & McNeilly 2004). There has been strong interest and expansion of canola in recent years in Western Europe, North
America (Zanetti et al., 2013) and in Australia (Salisbury & Barbetti 2011). There are industrial and commercial needs for producing canola and related crops for use as biofuels, food and cosmetics (Carlsson 2009).
In Australia, while Juncea canola (B. juncea) was grown widely for the first time in 2007 as a drought-tolerant B. juncea (Elliott et al., 2015), overridingly, the main canola grown is B. napus. Canola is grown as a winter crop over a wide
Page 3 of 119 range of areas from southern Queensland to Tasmania, and across southern
Australia to the Western Australian coast (Robertson et al., 2002). In Western
Australia, the area that is grown by canola has expanded from 35,000 ha in 1993 to 510,000 ha in 1998 (Barbetti & Khangura 1999) and subsequently to 1,100,000 ha in 2014 (AOF 2014). A major focus on canola in Australian breeding programs has resulted in high levels of oil and high protein concentration in cultivars
(Salisbury & Wratten 1999). However, this ability is severely and adversely impacted by a range of diseases. Due to their diversity and great value economically, the oilseed Brassicas, and especially canola, have become a host group where reducing the impact of different pathogens has been and remains a major focus (Barbetti et al., 2012).
1.3 Oilseed brassica downy mildew disease
1.3.1 Classification of Hyaloperonospora brassicae
Downy mildews are a group of pathogens belonging to Oomycetes under two orders Peronosporales and Sclerosporales which both belong to the class
Peronosporomycetes (Dick 2002a, b). Peronosporacea is the largest family in oomycetes and 800 species of downy mildews belong to this family (Göker et al.,
2007). Downy mildews (Peronosporaceae) pose challenges in terms of defining their taxonomy. Gäumann (1918, 1923) established a taxonomic classification for
Peronospora using the conidial measurements and that indicated specialization on host species rather than on host families. The taxonomy of downy mildew members, however, has been constantly reviewed and altered by researchers.
For instance, Peronospora parasitica that causes downy mildew diseases on brassicas was recently renamed as Hyaloperonospora parasitica
Page 4 of 119 (Constantinescu & Fatehi 2002) and they suggested division of Peronospora into
3 genera (Peronospora s. str., Hyaloperonospora and Perofascia). This suggestion was later supported by multi molecular studies that found some genetic differences even within H. parasitica and proposed this genus should be further divided (Choi et al., 2003; Göker et al., 2003; Voglmayer 2003; Thines
2007). More recently, the downy mildew pathogen of brassicas has been reclassified by Göker et al., (2009) as H. brassicae (syn. Hyaloperonospora parasitica).
While the morphology, physiology and ecology of downy mildews are close to fungi, molecular and biochemical studies have confirmed that downy mildews are not true fungi, but belong to the kingdom Chromista (Dick 2001). All downy mildews are considered obligate parasites and examples of endophytic biotrophs that typically form an intercellular mycelium with haustoria, and obtain their nutrition from within plant host cells (Spencer-Phillips & Jeger 2004). Downy mildews are obligate parasites worldwide, particularly in tropical and sub-tropical regions (Shivas et al., 2012).
1.3.2 Life cycle of Hyaloperonospora brassicae
The fungus is both soil and seed-borne and the disease cycle starts when resting oospores germinate and infect susceptible plants (Delourme et al., 2011), leading to development of pathogen colonies on host plants (Populer 1981). The infection process commences when an oospore germinates via development of a germ-tube that subsequently formats into a haustoria which is the key infection organ for the pathogen to penetrate the host. Within a few hours, the haustoria penetrates the epidermal cells of the host and the hyphae continues to progress
Page 5 of 119 between anticlinal walls of epidermal cells (Li et al., 2001). Subsequent secondary cycles of zoosporangial production in colony pustules on foliage are spread by wind and/or splashing water to infect other plants with new infections evident in as little as 3 to 4 days. Disease epidemics and the frequency of the infection can be increased rapidly when the period of moisture on host leaves increases (Mehta et al., 1995). Towards the end of the growing season, downy mildew colonies start to produce resting oospores within infested plant tissues which then remain in plant debris and/or soil in the field and can survive for the next season and, in most cases, for many years (Delourme et al., 2011).
1.3.3 Symptoms of downy mildew on brassicas
Hyaloperonospora brassicae (syn. H. parasitica) causes distinct foliar disease symptoms on oilseed and other Brassica species (Coelho et al., 2012).
First symptoms include chlorotic or yellow spots on the upper surface of leaves and with hyphal growth on the underside of the leaf surface, particularly in young plants. Infection at later stages of plant growth also reduces the productivity and quality of host plant (Silué et al., 1996). In some Brassica species, infection may extend to the curds, such as those of cauliflower, cabbage heads and broccoli
(Channon 1981). Symptoms on adult plants can vary, but most often appear as brown, black or greyish on the surface of leaves of oilseed and vegetable
Brassicas and on the curd heads of vegetable Brassicas. Systemic infection of internal plant tissues can lead to black discolouration (Koike et al., 2007;
Johansen-Hladilová 2010). H. brassicae causes downy mildew disease on canola and other oilseed brassicas to an extent that it is considered a major concern in Australia, particularly after the widespread serious outbreaks of this disease in the past decade (Ge et al., 2008).
Page 6 of 119
1.3.4 Host range of Hyaloperonospora brassicae on cruciferous species
The disease has been recorded frequently across a wide range of horticultural and agricultural species of the genus Brassica (Silué et al., 1996), affecting almost all crops belonging to Brassicaceae worldwide, including canola
(Khangura 2003). For example, H. brassicae (syn. H. parasitica) has been reported from cabbage (B. oleracea var. capitata) and turnip (B. campestris)
(Sherriff & Lucas 1990) and in Western Australia it was first recorded on B. napus
(oilseed rape) in 1973 (Shivas 1989). IIn the United Kingdom, it occurs frequently on winter oilseed rape (B. napus.) (Evans et al., 1984; Gladders 1987). Likewise, the disease it is also very widespread on Brassica juncea (mustard) in India
(Nashaat & Awasthi 1995). It also attacks vegetable brassicas at later growth stages such as curd heads of broccoli and cabbage (Natti 1958; Kontaxis &
Guerrero 1978) and cauliflowers (Lund & Wyatt 1978), and also on turnip
(Summer et al., 1978). H. brassicae can also infect weedy cruciferous species
(Chang et al., 1964; McMeekin 1969; Dickinson & Greenhalgh 1977) and in
Western Australia, H. brassicae is widespread on R. raphanistrum and isolates from this species are highly virulent (Maxwell & Scott 2008). H. brassicae also reported as causal agent of downy mildew on arugula (Eruca sativa) a leafy crucifer (Koike 1998; Romero & Zapata 2005) and wild rocket (Diplotaxis tenuifolia) (Garibaldi et al., 2004).
1.3.5 Downy mildew on canola and mustard
Downy mildew disease is endemic across the canola-growing regions in
Australia (Barbetti & Carter 1986; Howlett et al., 1999; Barbetti & Khangura 2000),
Page 7 of 119 Europe (Paul et al., 1998), China and Japan (Sato & Fukumoto 1996), and, in particular, on mustard on the Indian subcontinent (Nashaat et al., 2004).
Devastating outbreaks of downy mildew disease, particularly at the early seedling stage in the past few growing seasons, have demonstrated the severe impact of this disease on canola production in southern Australia. Further, the recent release of canola-quality B. juncea mustards for southern Australia is cause for major concern as this species is extremely susceptible to H. brassicae (Ge et al.,
2008). The infection of downy mildew on oilseed Brassica crops, can be high as
100%, especially in the early seedling stage (Smith & Price 1996).
1.3.6 Severity and impact of downy mildews on canola and mustard
Infection by H. brassicae (syn. H. parasitica) can be high as 100% on some
Brassica crops especially in the early seedling stage (Smith & Price 1996). The worldwide impact of downy mildew on oilseed brassicas is particularly significant because the productivity of vegetable oils from Brassica spp. represents approximately one-third of total vegetable oil production worldwide (Kaur et al.,
2011). The disease is readily transmitted via seed infestation (Smith & Price
1997), fostering long distance spread and establishment in new areas via this means. As mentioned earlier, losses from downy mildew on canola are estimated to be up to AU$13 million annually for Western Australia alone back in 2004.
Subsequently, monitoring of up to 33 sites across southern and eastern Australia across 2013-2015 by van de Wouw et al., (2016) found “downy mildew present at most sites assessed, with the site incidence ranging from 100% in 2013 to 52% in 2015.”
Page 8 of 119 1.3.7 Physiological specialisation within Hyaloperonospora brassicae populations
Pathotypes and pathogen races are very common across many plant pathogens, where isolates of similar morphology can have different potentials to infect different species within a particular genus of plant host (Lebeda & Cohen
2011). The host specificity can be utilized as indicator to recognize these different pathogen pathotypes (Sherriff & Lucas 1990). Thomas et al. (1987) used a particular method to recognize Pseudoperonospora cubensis pathotypes causing downy mildew on cucurbits. Defining pathotypes is a critical requirement if control of plant diseases is to be successfully achieved by using resistance cultivars
(Limpert et al., 1994). In particular, an understanding of host and non-host resistance to downy mildew (H. brassicae) could lead to the identification of physiological forms and pathotype markers for use in identifying and utilising the resistance(s) of the host against pathogen races (Li et al., 2011).
Isolates of H. brassicae exhibit specialization within the host family and generally are compatible with host genotypes of the species from which the isolate was derived (Channon 1981; Lucas et al., 1988; Satou 2000). While isolates of H. brassicae obtained from different Brassica spp. were found to be most virulent on their species of origin, they were still able to infect related species
(Chang et al., 1964; McMeekin 1969; Dickinson & Greenhalgh 1977; Kluczewski
& Lucas 1983; Sherriff & Lucas 1990; Nashaat & Awasthi 1995; Silué et al.,
1996). Genes for pathotype (sensn Lebeda) specific resistance to H. brassicae have been reported in B. oleracea (Natti et al., 1967), in B. napus (Lucas et al.,
1988; Nashaat et al., 1997), and in B. juncea (Nashaat & Awasthi 1995; Nashaat et al., 2004). Among these reports, Natti et al. (1967) was the only one that reported existence of separate distinct physiologic races (1 and 2) in broccoli (B.
Page 9 of 119 oleracea var. italica). Sherriff & Lucas (1990) used 33 isolates from four different
Brassica hosts (B. napus, B. juncea, B. oleracea and B. campestris) and locations to infect a set of Brassica accessions (including B. napus, B. juncea, B. oleracea,
B. carinata, B. nigra, B. campestris and Raphanobrassica). They analysed the variation in host range within the H. brassicae population on agriculturally and horticulturally important Brassica crops, but failed to delineate pathotypes of H. brassicae. Unfortunately, none of these studies above allows any comparison between pathotype structure in different crops, different countries or continents.
To date, no effort has been made to compile a listing of known pathotypes due to lack of access to host varieties/lines and/or pathogen isolates that have previously been used. Therefore, knowledge of pathotype structure of H. brassicae still remains unclear and for Australia is totally unknown and addressing this deficiency is an important aspect of the research reported in this thesis.
1.3.8 Impact of environmental factors on development of downy mildew
Oilseed Brassicas are affected by climate variations including fluctuations in temperature, different levels of rainfall, emissions of CO2, drought and even consequent higher levels of salinity in some circumstances. For example, in the southwest of Western Australia, rainfall variation annually affects growth of
800,000 ha or more of canola (Barbetti et al., 2012), affecting yield and oil content
(Salisbury & Barbetti 2011). Hocking & Stapper (1993) showed that an increase in temperature by 1°C can cause a 1.2-1.5%, reduction in canola oil content,
Page 10 of 119 especially if this occurs at the seed-fill stage. Downy mildews are also similarly affected, with the economic losses from downy mildew on pea, grapevine, or rose dependent upon temperature, humidity, infected part of plant and/or growth stage
(Biddle, 2001; Williams, 2005; Aegerter et al., 2003). H. brassicae (syn. H. parasitica) is no exception, as it is known to be favoured by warm days (20-24°C), cool nights (8-16°C), and a relative humidity >80% (Channon 1981). Temperature affects its conidial germination, formation of the appressoria and the rate of hyphal penetration into the host (Chu 1935). Felton & Walker (1946) reported that on leaves of cabbage (B. oleracea), H. brassicae germinates and penetrates at
6-24⁰C, colonizes at 8-16⁰C, and sporulates at 4-24⁰C (optimum 12-16°C), the latter optimum confirmed by Hartmann et al., (1983) at 13-18⁰C. Achar (1998), however, showed H. brassicae germination on cabbage was greatest at 20⁰C with 100% relative humidity; while Kofoet & Fink (2007) noted that on radish
(Raphanus sativus) downy mildew was observed from 8.3-26.7⁰C. Sangeetha &
Siddaramaiah (2007) reported that downy mildew occurs across temperatures
14-29⁰C. However, there have been no studies specifically defining the role of temperature on the development of downy mildew epidemics on either canola (B. napus) or mustard (B. juncea) and, as such this is also an important research focus of this thesis.
1.3.9 The effect of plant age on downy mildew epidemics
While H. brassicae (syn. H. parasitica) particularly affects seedlings of brassicas, it also can infect and damage adult plants (Silué et al., 1996; Coelho et al., 2012). For example, at later stages of growth in some B. oleracea plants, infection by H. brassicae results in sporulating lesions that led to significant
Page 11 of 119 economic losses as well as poor quality of broccoli and cauliflower heads (Niu et al., 1983; Jensen et al., 1999). Plant growth stage or plant age can affect host susceptibility to H. brassicae (Coelho et al., 2009). For example, broccoli (B. oleracea var. italica) leaves on plants with more than eight leaves were less susceptible than cotyledons (Coelho & Monteiro 2003). Also, Agnola et al., (2003) used leaf discs from different ages of Brassica plants to compare the resistance to H. Brassicae and found that discs collected from the first to sixth leaf were susceptible in comparison with discs collected from the seventh upwards leaves that showed moderate susceptibility to resistant. In addition, association between expression of resistance to H. brassicae at cotyledon stage with that at the adult stage has been reported by Jensen et al., (1999) on cauliflower (Brassica oleracea var. botrytis), by Wang et al., (2000) on inbred broccoli, and by Zhang et al., (2012) in B. rapa (Chinese cabbage). Coelho et al., (1998) reported that resistance in B. oleracea cotyledons had no association with that in adult plants.
However, there are no studies defining the role of plant age on the development of downy mildew epidemics specifically on oilseed B. napus or B. Juncea, and for this reason this was an important research focus of this thesis
1.3.10 Management of downy mildew disease
One of the challenges in managing downy mildew pathogen is that it has a very short period of latency following infection, leading to rapid disease increase
(Spencer-Phillips & Jeger 2004). This can make appearance and development of severe epidemics fast and unforeseen. Fungicides or chemical controls are widely applied by brassica growers to protect the crops from downy mildew disease (Monteiro et al., 2005) and offer a degree of control of H. brassicae
(Brophy & Laing 1992). Fungicides may be used in different ways, such as seed
Page 12 of 119 treatment or spraying on foliar parts of infected plants (Gisi 2002). For instance,
Jensen et al., (1998) reported that treating B. oleracea seeds with “CGA245704, an activator of systemic acquired resistance, effectively reduced H. brassicae sporulation. However, excessive use of fungicides can lead to some negative consequences. For example, downy mildews have the potential to evolve and overcome the control by fungicides (Tyler 2007). Crute et al., (1985) reported that
P. brassicae developed resistance to metalaxyl. Likewise, Molina et al., (1998) mentioned that P. brassicae became resistant to other fungicides. The use of fungicides or cultural strategies has provided only limited or partial control with high cost (Barbetti et al., 2011; Neik et al., 2017). While controlling downy mildew disease by fungicides can be inefficient or can offer limited protection, using resistant genotypes is considered the most effective method to manage this important pathogen (Thomas & Jourdain 1992).
1.4 Sources of resistance in oilseed brassicas and other brassicas
The limited and often inadequate control from fungicides has fostered the search to find new strategies to reduce yield losses in canola and other Brassica crops from H. brassicae (Eshraghi et al., 2007). Plants have a range of different mechanisms to protect against or at least reduce the impact of pathogens like H. brassicae (Li et al., 2011). Sources of resistance across different oilseed
Brassicas have been identified and at different stages of plant growth (Dias et al.,
1993). Systemic acquired resistance (SAR; natural defence of the plants toward pathogens) also offers opportunities (Ryals et al., 1994). For example, Jensen et al., (1998) found that the sporulation intensity of H. brassicae on seedlings of
Page 13 of 119 Brassica was controlled by using CGA 245704, a chemical activator of systemic acquired resistance. Further, Greenhalgh & Mitchell (1976) conducted a study on commercial and wild populations of B. oleracea cultivars to determine the resistance of these taxa to H. brassicae. Whilst one commercial cultivar was resistant to the pathogen, all wild populations of B. oleracea showed high resistance to H. brassicae due to its high concentrations of toxic volatiles to this pathogen. Genetic mapping of a doubled haploid population from a cross two cultivars of B. napus highlighted a single major gene QTL that controls seedling resistance of Chinese cabbage (B. rapa var. pekinensis) to H. brassicae (Yu et al., 2009).
Specific resistances to H. brassicae (syn. H. parasitica) have been reported in B. oleracea (Natti et al., 1967; Farnham et al., 2002; Monteiro et al., 2005), and various resistances reported in B. napus (Lucas et al., 1988; Nashaat et al., 1997) and in B. juncea (Nashaat & Awasthi 1995; Nashaat et al., 2004; Chattopadhyay
& Séguin-Swartz 2005). Expressions of field susceptibilities/resistances to downy mildew have been reported for the seedling stage of B. napus (Ge et al., 2008) and on more mature leaves of B. oleracea (Dickson & Petzoldt 1993; Coelho et al., 1998; Jensen et al., 1999a,b; Carlsson et al., 2004; Wang et al., 2001; Coelho et al., 2009). In B. oleracea, Dickson & Petzoldt, (1996) reported presence of quantitative resistance, Hoser-Krauze et al., (1991) reported a single recessive gene, Hoser-Krauze et al., (1995), Moss et al., (1988), Caravalho and Monteiro
(1996) and Vicente et al., (2012) all reported one or more dominant resistance genes. In B. juncea, Nashaat et al. (2004) reported a single dominant gene while, in B. napus, Lucas et al. (1988) and Nashaat & Rawlinson (1994) both reported a single dominant gene.
Page 14 of 119 The first attempt to screen genotypes to locate new sources of resistance in Australian genotypes of canola and mustard H. brassicae was conducted in
Western Australia (Ge et al., 2008). That study found two genotypes of Australian oilseed rape were highly resistant to H. brassicae. In other studies, light and electron microscopy technology (TEM) have also been used to study the interactions between Arabidopsis leaves and H. brassicae during the development of the disease. For example, TEM was used to study the penetration of H. brassicae hyphae, haustorium development and the structure and contents of host cells (Soylu & Soylu 2003). Further, Li et al. (2011) investigated the responses and resistances of host and non-host resistances to H. brassicae to identify the physiological forms and pathotype markers for developing host resistance against prevailing pathogen races. However, as there has not been subsequent investigations into resistances of more current Australian canola varieties, nor has a wider search been made across B. juncea and diverse
Brassica species for alternative host resistances, these were a focus of the PhD studies reported in this thesis.
1.5 Concept and Aims of Work
As detailed above, downy mildew is a major debilitating disease adversely affecting the production of oilseed brassicas like canola and mustard that together represent about one-third of the total vegetable oil production worldwide (Kaur et al., 2011). Production of canola in Australia ranks third after wheat and barley, and the canola area has nearly doubled from 1.4 million ha in 2009 to 2.4 million ha in 2013 (Elliott et al., 2015). Further, Brassicas have been recognised as providing significant scientific educational tools as this group makes a ‘near- perfect’ crop for research studies (Dixon 2007). In particular, H. brassicae on
Page 15 of 119 canola (and mustard) has become a major concern across Australia particularly after the devastating outbreaks of this disease in the past decade (Ge et al.,
2008).
This project highlights how the incidence, severity and impact of downy mildew on oilseed Brassicas varies with different stages of plant growth. It also highlights new sources of host resistance in different Brassica species as identified using controlled environment studies and defines the impact of environmental factors like temperature on the development of downy mildew disease. Finally, it provides the first understanding and definition of the pathogen phenotypic (i.e., pathotypes) and genetic variation in Australia and a system that can be universally applied for characterisation of pathogen pathotypes.
The broad aim of this work is to develop new understanding of the role of pathogen phenotypic and phylogenetic variation, environmental factors and plant growth stage in development of downy mildew epidemics on oilseed Brassicas
(particularly rapeseed, canola, mustard) and the potential for locating and utilizing novel host resistances to allow development of effective control strategies for downy mildew.
The particular objectives of the project as follows:
1. Define relative host resistances in Australian oilseed and other diverse
Brassica germplasm.
Page 16 of 119 2. Define the role of environmental factors like temperature and plant growth
stage in determining the development of downy mildew epidemics.
3. Define pathogen phenotypic and phylogenetic variation across Australia
(i.e., phylogenetic relationships between isolates and whether there are
pathogen races or pathotypes).
To achieve the objectives above, the following experimental works have been completed:
1. Testing 131 Brassicaceae varieties at the cotyledon stage, including 109
Australian canola varieties (Brassica napus and B. juncea) and 22 different
Brassicaceae to examine their level of resistance to mixture of ten isolates
of H. brassicae under controlled environment conditions. This was
followed by screening of a further 154 diverse Brassicaceae for additional
new sources of resistance to H. brassicae.
2. Investigation of the effect of two different temperatures (14/10⁰C and
22/17⁰C day/night) on the development of downy mildew disease on B.
juncea and B. napus varieties.
3. Investigation of the effect of different plant ages (15, 23, 31 and 40 day-
old-plants) on the downy mildew severity of H. brassicae on selected B.
juncea and B. napus varieties.
Page 17 of 119 4. Inoculation of 28 diverse Brassicaceae genotypes at the cotyledon stage
with 11 different isolates of H. brassicae to identify and select suitable
Brassica spp. host differentials to enable characterisation of H. brassicae
pathotypes; and then definition of the phylogenetic relationships among
isolates across Australia.
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Page 34 of 119 Chapter 2
New resistances offer opportunity for effective management of the downy mildew (Hyaloperonospora parasitica) threat to canola
This chapter focuses on the level of resistance among Australian canola genotypes to downy mildew at the cotyledon stage.
Page 35 of 119 CSIRO PUBLISHING Crop & Pasture Science, 2017, 68, 234 242 http://dx.doi.org/10.1071/CP16363
New resistances offer opportunity for effective management of the downy mildew (Hyaloperonospora parasitica) threat to canola
A. E. Mohammed A,B, M. P. You A, and M. J. Barbetti A,C
AUWA School of Agriculture and Environment and the UWA Institute of Agriculture, Faculty of Science, The University of Western Australia, Crawley, WA 6009, Australia. BDepartment of Plant Protection, Faculty of Agriculture, University of Kufa, Najaf, Iraq. CCorresponding author. Email: [email protected]
Abstract. Downy mildew (Hyaloperonospora parasitica) is a problem for canola production worldwide, including in Australia where it has remained a persistent threat since 1998. Testing of 131 Brassicaceae varieties, including 109 Australian canola varieties (Brassica napus and B. juncea) and 22 diverse Brassicaceae (including B. napus, B. carinata, B. juncea, B. nigra, B. rapa, Crambe abyssinica and Raphanus sativus) highlighted excellent resistance to downy mildew. Using a mixture of 10 H. parasitica isolates, R. sativus Colonel and Boss showed highest resistance to H. parasitica, with per cent disease index (%DI) values of 3.7% and 10.2%, respectively. These were followed by (%DI values): B. carinata ATC 94011 (11.1%), B. napus CB Tanami (14.1%) and Komet 741 A (14.3%), B. juncea 397.23.2.3.3 (14.8%), B. napus ATR Banjo (16.9%), Hyola 575 CL (16.9%), Komet 744 A (18.1%), Cresor 770 B (18.5%), Wamus (18.5%), Surpass 400 (19.2%), Hyola 432 (19.4%) and Hyola 76 (19.4%), and C. abyssinica (19.9%). These varieties were also considered highly resistant. Another five B. juncea genotypes and B. nigra P.23845 were considered highly resistant with %DI of 22.2%. Those considered resistant (but not highly resistant) included hybrid B. napus Hyola 444 TT, Hyola 500 RR, Hyola 504 RR, Pioneer 46Y78, Pioneer 45Y77 and Hyola 650 TT, and the non hybrid variety ATR Eyre, all with %DI values 23.1 28.2%. By contrast, B. napus Thunder TT, Hyola 450 TT and ATR Grace were highly susceptible with %DI values of 90.3%, 88.2% and 81.7%, respectively. Cluster analysis revealed six distinct clusters (highly resistant, resistant, moderately resistant, moderately susceptible, susceptible, very susceptible) for the tested Brassicaceae genotypes that, on average, showed similar responses within eachcluster againstH. parasitica based on their%DI values. From 2000 onwards (with theexception of Surpass 400), 10 B. napus varieties and one B. juncea released were classified as highly resistant; however, there was no overall correlation between year of variety release and level of resistance expressed against H. parasitica. This is the first study to demonstrate the existence of very high levels of pathotype independent resistance in Australian canola varieties to H. parasitica. The most resistant varieties identified can be used in canola breeding programs and also directly deployed into regions where downy mildew is prevalent, providing the canola industry with an immediate and effective option for management of this important disease.
Additional keywords: disease screening, disease management, host resistance, oilseed rape.
Received 4 October 2016, accepted 2 March 2017, published online 29 March 2017
Introduction and Kummer 2013). Downy mildew has most adverse effects Downy mildew in Brassicaceae, caused by the obligate oomycete on young Brassicaceae seedlings, but can still significantly Hyaloperonospora parasitica, is widespread worldwide (Koch reduce the productivity and quality even when plants are and Slusarenko 1990), including in Europe (Paul et al. 1998) infected at a later stage of growth (Silué et al. 1996). In and China and Japan (Satou 1996). It is also an endemic Western Australia, downy mildew was first recorded on disease across canola growing regions in Australia (Howlett B. napus (oilseed rape) in 1973 (Shivas 1989). There and et al. 1999; Barbetti and Khangura 2000), and has become elsewhere across southern Australia, devastating outbreaks of much more prevalent since 1998 (Barbetti and Khangura downy mildew demonstrated the capacity of this disease to 2000), particularly in Western Australia where there have been impact canola production severely. A survey in 2004 (Oilseeds severe epidemics in the past two seasons (M. J. Barbetti, unpubl. Industry Association of Western Australia, unpubl. data), data). Although this pathogen generally attacks leaves and estimated the economic losses from downy mildew on canola cotyledons of oilseed and other Brassica species (Coelho et al. to be up to AU$13 million annually in Western Australia alone. 2012), it can also infect flowering structures and cause systemic Subsequently, monitoring of up to 33 sites across southern and symptoms on flowers or non floral parts of their hosts (Thines eastern Australia during 2013 15 by van de Wouw et al.(2016)
Journal compilation CSIRO 2017 www.publish.csiro.au/journals/cp Page 36 of 119 Downy mildew resistance in canola Crop & Pasture Science 235 found ‘downy mildew present at most sites assessed, with the Origins of H. parasitica isolates ’ site incidence ranging from 100% in 2013 to 52% in 2015 . One A mixture of 10 isolates of H. parasitica was used in this study. of the challenges in managing this pathogen is that it has only Nine isolates were collected from canola across geographically a very short period of latency following infection, leading to diverse sites in Western Australia: isolates WA 5 (E116.38017, rapid disease increase (Spencer Phillips and Jeger 2004). This can S31.14390), WA7 E116.50517, S31.05416), WA11 (E116. make appearance and development of severe epidemics fast and 78858, S31.08365), WA12 (E116.80164, S31.124770), WA13 unforeseen. (E116.81889, S31.18898), WA22 (E117.11864, S31.64116), Limited and often inadequate control from fungicides, in WA29 (E116.97926, S31.94476), WA30 (E116.86858, S32. fi addition to their cost, has provoked a search to nd new 40118) and WA49 (117.14737, S33.79678). Isolate SA78 strategies to reduce yield losses in canola and other Brassica (E138.20108,S32.83453) was collected from South Australia. crops from H. parasitica (Eshraghi et al. 2007). Among the These isolates were chosen because they were collected from Brassicaceae, there are genotypes within B. napus, B. oleracea, across a wide geographical area severely affected by downy and B. juncea that show at least some host resistance to one or mildew in a southern Australia 2015 canola disease survey. more H. parasitica isolates. For example, Gröuntoft (1993) All H. parasitica isolates used were either single spored showed that some varieties of B. napus and B. campestris have (conidium) or hyphal tipped from the original field samples resistance at the cotyledon stage. Resistance has also been collected. Identity of all isolates as H. parasitica was fi identi ed in broccoli (B. oleracea var. cephalata) (Thomas confirmed through comparison with available sequence data and Jourdain 1990; Dickson and Petzoldt 1993; Silué et al. information for H. parasitica in GenBank by using the primer 1996; Giovannelli et al. 2002) and in B. juncea (mustard) ITS4 H (50 TCC TCC GCT TAT TAA TAT GC), a modification (Nashaat and Awasthi 1995; Nashaat et al. 2004; Delourme of ITS4, following DNA extraction. fi et al. 2011). The rst study to highlight the existence of DNA was extracted by using the procedure of Cenis (1992) resistance in canola against H. parasitica in Australia was that whereby mycelium from leaf samples was harvested directly, of Ge et al.(2008), who screened 63 canola varieties and blotted dry with filter paper, washed with Tris EDTA (TE) fi identi ed two varieties, Pioneer 45Y77 and Pioneer 46Y78, buffer, and pelleted by centrifugation (5 min at 13 000 rpm), fi that were highly resistant to one or more speci c H. parasitica and the supernatant decanted. After adding 300 mL of extraction isolates. Despite these earlier studies, the increased and ongoing buffer [200 mM Tris HCl (pH 8.5), 250 mM NaCl, 25 mM EDTA, incidence of severe downy mildew epidemics in oilseed brassicas and 0.5% (w/v) sodium dodecyl sulfate], the mycelia were in Australia since 1998 (Barbetti and Khangura 2000) highlights crushed manually, then 150 mLof3M sodium acetate (pH 5.2) an urgent need to locate and deploy Australian cultivars with was added, and tubes maintained at 208C for 10 min. The high level effective resistance to downy mildew. This would supernatant was then transferred to a fresh tube before adding not only provide an immediate management tool for growers an equal volume of isopropanol and centrifuged for 5 min at affected by severe downy mildew, but would enable breeding of 13 000 rpm. After at least 5 min at ambient temperature (~258C), new, more resistant cultivars. the precipitated DNA was collected by centrifugation for 15 min The study we report in this paper highlights new and very at 13 000 rpm. The pellet was vacuum dried for several minutes high level resistances within 131 Brassicaceae including 109 after washing with 70% (wt/vol) ethanol and resuspended in Australian canola varieties and 22 diverse Brassicaceae varieties. 50 mL of TE buffer. The concentration and quality of the extracted It discusses the implications and opportunities for improved DNA was determined by a NanoDrop 1000 Spectrophotometer management of this important disease in Australia from (Thermo Scientific, Waltham, MA, USA). The DNA was stored deployment of effective resistance across disease prone regions. at 48C until required. Each H. parasitica isolate was maintained separately in Materials and methods isolated containers on cotyledons of 7 day old seedlings of susceptible B. napus Thunder TT and Tranby (Ge et al. 2008) Varieties in a controlled environment room with a 12 h photoperiod, light The 131 Brassicaceae genotypes included 109 Australian canola intensity 150 mEm–2 s–1 and temperature regime 138C (night) varieties (Brassica napus and B. juncea) and 22 diverse and188C (day). Equal numbers of cotyledons supporting Brassicaceae (B. napus, B. carinata, B. juncea, B. nigra, abundant sporulation by each of the 10 different pathogen B. rapa, Crambe abyssinica and Raphanus sativus). Brassica isolates were collected and placed in 50 mL distilled water in a napus Pioneer 46Y78 and 45Y77 were included as controls 100 mL flask. The flask was shaken to dislodge the zoosporangia. fi because they have previously been identi ed as resistant The resulting suspension was filtered through one layer of genotypes (Ge et al. 2008). Other Brassicaceae included cheesecloth to remove any cotyledon contamination and the B. napus Q2 [a Canadian variety highly susceptible to concentration was adjusted to 105 sporangia mL–1 by using a blackleg disease (Leptosphaeria maculans)], used universally haemocytometer counting chamber. A range of representative as a control check for L. maculans studies (Sosnowski et al.2005), isolates of H. parasitica was chosen and mixed to enable fi along with ve winter types of B. napus (Komet 741 A, Komet identification of pathotype independent host resistances. 744 A, Cresor 770 B, Cresor 771 B and Capricorn C) (Table 1). Information on the approximate year in which each Australian variety was released (or if not commercially released, the year Inoculation the selection was available for testing) was gathered from Six seeds of each Brassicaceae test genotype were sown in 6 cm commercial sources. by 6 cm seedling tray cells and thinned to four plants per cell
Page 37 of 119 236 Crop & Pasture Science A. E. Mohammed et al.
Table 1. Per cent disease index (%DI) values on cotyledons, and origin and year of release for 131 Brassicaceae genotypes including 109 Australian canola varieties (Brassica napus and B. juncea, indicated A) and 22 diverse Brassicaceae (including B. napus, B. carinata, B. juncea, B. nigra, B. rapa, Crambe abyssinica and Raphanus sativus) inoculated with mixture of 10 different isolates of Hyaloperonospora parasitica Significance of genotypes for %DI, P < 0.001; l.s.d. at P 0.05, 7.56. Cluster number relates to cluster distributions shown in Fig. 1
Brassicaceae sp. Genotype Seed origin %DI Release date Cluster number Raphanus sativus Colonel Europe 3.7 n.a. I R. sativus Boss Europe 10.2 n.a. I Brassica carinata ATC 94011 Australia 11.1 n.a. I B. napus (A) CB Tanami Australia 14.1 2007 I B. napus Komet-741 A Europe 14.3 n.a. I B. juncea 397.23.2.3.3 Australia 14.8 n.a. I B. napus (A) ATR-Banjo Australia 16.9 2006 I B. napus (A) Hyola 575 CL Australia 16.9 2010 I B. napus Komet-744 A Europe 18.1 n.a. I B. napus Cresor-770 B Europe 18.5 n.a. I B. napus (A) Wamus Australia 18.5 n.a. I B. napus (A) Surpass 400 Australia 19.2 1999 I B. napus (A) Hyola 432 Australia 19.4 2000 I B. napus (A) Hyola 76 Australia 19.4 2008 I Crambe abyssinica CMB 94054 19.9 n.a. I B. juncea Montara TT China 21.3 n.a. I B. juncea (A) Dune Australia 22.2 2007 I B. nigra P.23845 22.2 n.a. I B. juncea JM06018 Australia 22.2 n.a. I B. juncea Montara China 22.2 n.a. I B. juncea Muscan 963 Australia 22.2 n.a. I B. napus (A) Hyola 444 TT Australia 23.1 2013 II B. napus (A) Hyola 500 RR Australia 24.3 2013 II B. napus (A) Hyola 504 RR Australia 25 2016 II B. napus (A) Pioneer 46 Y78 Australia 25.9 2007 II B. napus (A) Pioneer 45Y77 Australia 26.4 2007 II B. napus (A) Hyola 650 TT Australia 26.6 2013 II B. napus (A) ATR-Eyre Australia 28.2 2002 II B. napus (A) ATR-Summit Australia 30.8 2006 II B. napus (A) ATR-Hyden Australia 32.2 2001 II B. napus (A) Trooper Australia 36.3 2002 II B. napus (A) AV-Jade Australia 38.7 2006 II B. napus (A) Jackpot TT Australia 41.4 2011 II B. napus (A) Hyola 559 TT Australia 42.4 2012 II B. napus (A) AV-Sapphire Australia 43.1 2003 II B. napus (A) Hyola 474 CL Australia 43.3 2011 II B. napus (A) Hurricane Australia 46.3 2008 II B. napus (A) Crusher TT Australia 46.5 2010 II B. napus Q2 Canada 46.8 n.a. II B. napus (A) Rottnest Australia 47.2 2007 II B. napus RT108 Australia 47.7 n.a. II B. napus (A) Oscar Australia 48.1 1992 II B. napus (A) Atomic Australia 48.8 2004 II B. napus (A) Hyola 577C Australia 48.8 2013 II B. napus (A) Hyola 400 RR Australia 49.1 2013 II B. napus (A) Warrior Australia 49.1 2006 II B. napus (A) Hyola 61 Australia 49.3 2004 II B. napus (A) Hyola 505 RR Australia 50 2010 III B. napus (A) CB Telfer TT Australia 50.2 2009 III B. napus (A) Bravo TT Australia 50.9 2005 III B. napus (A) Cresor-771 B Europe 50.9 n.a. III B. napus (A) ATR-Barra Australia 51.2 2007 III B. napus (A) Maluka Australia 51.2 1988 III B. napus (A) Surpass 501 TT Australia 51.2 2001 III B. napus (A) Rocket CL Australia 51.8 2005 III B. napus (A) CB Trilogy Australia 51.8 2004 III (continued next page)
Page 38 of 119 Downy mildew resistance in canola Crop & Pasture Science 237
Table 1. (continued )
Brassicaceae sp. Genotype Seed origin %DI Release date Cluster number B. rapa Vaslitti 51.8 n.a. III B. napus (A) CB Trigold Australia 52.1 2004 III B. napus (A) ATR-Wahoo Australia 52.3 2013 III B. napus (A) Beacon Australia 52.8 2002 III B. napus (A) Granite TT Australia 53 2015 III B. napus (A) Scoop Australia 53 1996 III B. napus (A) AG-Muster Australia 53.2 2007 III B. napus Westar Canada 53.5 1989 III B. napus (A) Hyola 404 RR Australia 53.9 2010 III B. napus (A) Hyola 635 CC Australia 54.4 2014 III B. napus (A) Surpass 300 Australia 55.1 2001 III B. napus ZY006 China 55.1 n.a. III B. napus (A) Fighter TT Australia 55.3 2007 III B. napus (A) Hyola 50 Australia 55.3 2007 III B. napus (A) Hyola 43 Australia 55.6 2003 III B. napus (A) AG-Emblem Australia 55.8 2000 III B. napus (A) AG-Outback Australia 56 2001 IV B. napus (A) Surpass 402 Australia 56 2001 IV B. napus (A) ATR-Stubby Australia 56.5 2004 IV B. napus (A) Georgie Australia 56.5 2000 IV B. napus (A) Signal 6.421 Australia 56.5 n.a. IV B. napus (A) Surpass 603 CL Australia 56.5 2001 IV B. napus (A) ATR-Cobbler Australia 56.7 2007 IV B. napus (A) Hyclean Australia 56.7 n.a. IV B. napus (A) 47CO2 Australia 57.2 1999 IV B. napus (A) Hyola 401 Australia 57.6 1991 IV B. napus (A) Mystic Australia 57.6 1998 IV B. napus (A) Siren Australia 57.6 1993 IV B. napus (A) Hyola 433 Australia 58.1 2010 IV B. napus (A) Purler Australia 58.3 2000 IV B. napus (A) Lantern Australia 58.6 2002 IV B. napus (A) 46CO1 Australia 59.7 1999 IV B. napus (A) CB Boomer Australia 59.7 2006 IV B. napus (A) 44C76 Australia 59.9 2004 IV B. napus (A) Hyola 60 Australia 60.4 2001 V B. napus (A) Warrior CL Australia 60.9 2006 V B. napus (A) AG Comet Australia 61.3 2005 V B. napus (A) Pioneer 47 Co2 Australia 61.3 1999 V B. napus (A) AV-Ruby Australia 62 2006 V B. napus (A) Surpass 404 CL Australia 62.7 2003 V B. napus (A) Wesbarker Australia 63 1987 V B. napus Capricon C Europe 63.2 n.a. V B. napus (A) Surpass 600 Australia 63.9 1999 V B. napus (A) Archer Australia 64.1 2012 V B. napus (A) Insignia Australia 64.1 2000 V B. napus 03-P74-11 China 64.3 n.a. V B. napus (A) Flinders Australia 64.3 2008 V B. napus (A) Hylite 200 TT Australia 64.6 1999 V B. napus (A) AG-Castle Australia 65.5 2002 V B. napus (A) Marnoo Australia 65.5 1980 V B. napus (A) Hyola 51 Australia 65.7 2007 V B. napus (A) AG-Spectrum Australia 66.4 2004 V B. napus (A) Tornado TT Australia 67.6 2004 V B. napus (A) CB Tribune Australia 67.6 2004 V B. napus (A) Bugle Australia 67.8 2000 V B. napus (A) Mystic TT Australia 68.1 n.a. V B. napus (A) Rivette Australia 68.3 2002 V B. napus (A) Monty Australia 68.5 1996 V B. napus (A) Karoo Australia 69.2 1996 V B. napus (A) Skipton Australia 69.2 2005 V
(continued next page)
Page 39 of 119 238 Crop & Pasture Science A. E. Mohammed et al.
Table 1. (continued )
Brassicaceae sp. Genotype Seed origin %DI Release date Cluster number B. napus (A) 46C72 Australia 69.4 2000 V B. napus (A) Sturt TT Australia 69.4 2012 V B. napus (A) Pinnacle Australia 72.9 1996 VI B. napus (A) Tranby Australia 74.8 2004 VI B. napus (A) 45C75 Australia 75.2 2001 VI B. napus (A) ATR-Signal Australia 75.7 n.a. VI B. napus 06-P71-2 China 76.6 n.a. VI B. napus (A) Hyola 656 TT Australia 77.3 2012 VI B. napus (A) 44C73 Australia 77.8 2001 VI B. napus (A) Grouse Australia 78 1996 VI B. napus (A) Dunkeld Australia 78.5 1993 VI B. napus (A) Hyola 42 Australia 79.4 1991 VI B. napus (A) ATR-Grace Australia 81.7 2001 VI B. napus (A) Hyola 450 TT Australia 88.2 2013 VI B. napus (A) Thunder TT Australia 90.3 2005 VI following seedling emergence. Genotypes were grown in a findings, the experiment was repeated once, but using only 30 of controlled environment room (188C day 138C night) with the original genotypes from the first experiment. In each a daylength of 12 h. Plants were watered daily with deionised experiment, each treatment replicate consisted four plants with water and allowed to drain to field capacity. At growth stage the disease index data for each of the four plants in each replicate. 1.00 (Sylvester Bradley and Makepeace 1984), 7 days after Common genotypes across both experiments behaved similarly, seedling emergence, seedling trays were placed in clear plastic with data across common varieties from the original and the repeat boxes (790 mm long, 390 mm wide, 155 mm high) with lids. experiments not significantly different (P > 0.05) using a t test. Seedlings were inoculated with a mixture of 105 sporangia mL–1 Hence, only data from the first experiment are presented. Disease suspension containing each of the 10 H. parasitica isolates, severity (%DI) data were analysed by one way ANOVA with by using a micropipette to place a 10 mL droplet of the mixed GENSTAT Release 14.2 (14th Edition; Lawes Agricultural Trust, sporangial suspension onto each of two lobes of both cotyledons Rothamsted Research, UK). Fisher’s least significant differences of each seedling. High humidity was maintained for 24 h (l.s.d.) were used to separate significant differences between post inoculation by hand misting the walls and lids of the each genotypes. Subsequently, cluster distributions (Ward’sminimum plastic box with deionised water and then keeping the clear variance showing Mahalnobis squared Euclidean distance, where plastic lid on. Subsequently, lids were removed and plants the initial cluster distances in Ward’s minimum variance method maintained by daily hand watering for 6 days. Then the plastic are therefore defined to be the squared Euclidean distance between boxes were covered again for a further 24 h to maintain a second points) were used to group test genotypes of similar level of period of high humidity. resistance to H. parasitica, based on %DI as expressed against this pathogen. We chose this method because Ward’sminimum Disease assessment variance criterion minimises the total within cluster variance, and The procedure of Williams (1985) was used to assess disease as it has been utilised to determine cluster distributions for other severity on plants at 7 days post inoculation (dpi) by using a 0 9 canola pathogens such as Sclerotinia sclerotiorum (Barbetti et al. scale: 0, no symptoms or sign of downy mildew disease; 1, minute 2014).Correlation coefficients were computedfor the relationship scattered necrotic flecks under the inoculum drop, no sporulation; between year of Australian variety release and the level of 2, larger necrotic flecks under the inoculum drop, no sporulation; resistance against H. parasitica. 3, very sparse sporulation, one to a few conidiophores, necrotic flecking but often with tissue necrosis evident; 5, sparse Results sporulation, tissue necrosis; 7, abundant sporulation, tissue There were significant (P < 0.001) differences across the test necrosis and chlorosis may be present; 9, heavy sporulation, genotypes in both the original and the duplicate experiments cotyledon collapse. Subsequently, 0 9 disease severity scores of this study, but overall, common genotypes across the two for each seedling tray cell were converted into a disease index experiments behaved similarly. The two R. sativus genotypes, (%DI) based on the methods described by McKinney (1923): Colonel and Boss, showed highest resistance to H. parasitica, %DI ¼ ððða 0Þþðb 1Þþðc 2Þþðd 3Þþðe 4Þ with %DI values of 3.7% and 10.2%, respectively. These were þ ...ðj 9ÞÞ 100Þ=ðða þ b þ c þ d þ ...jÞ 10ÞÞ followed by (%DI values): B. carinata ATC 94011 (11.1%), B. napus CB Tanami (14.1%) and Komet 741 A (14.3%), where a, b, c, d, e, ...j are the number of plants with disease B. juncea 397.23.2.3.3 (14.8%), B. napus Banjo (16.9%), Hyola severity scores of 0, 1, 2, 3, 4, ...9, respectively. 575 CL (16.9%), Komet 744 A (18.1%), Cresor 770 B (18.5%), Wamus (18.5%), Surpass 400 (19.2%), Hyola 432 (19.4%) and Experimental design and statistical analyses Hyola 76 (19.4%), and C. abyssinica (19.9%). These genotypes The experiment was arranged in a randomised complete block were also classified as highly resistant (Table 1). Another five design with three replicates of each treatment. To confirm the B. juncea genotypes and B. nigra P.23845 were also considered
Page 40 of 119
Downy mildew resistance in canola Crop & Pasture Science 241 demonstrated that the overall resistance level across B. napus varieties identified in this study as ‘highly resistant’ will most varieties has changed little across the period from 1989 until significantly curtail the incidence, severity and impact of now. However, with the exception of Surpass 400 released in downy mildew in those regions and situations. This is 1999, the highly resistant canola varieties have all been released especially so because the highly resistant cultivars identified since 2000. These are agronomically well adapted varieties that in this study almost certainly have pathotype independent could easily be deployed strategically both to reduce the adverse resistance(s) that should be effective across different regions impact of downy mildew and to reduce the overall amount of where canola is grown. Existence of pathotypes is also inoculum pressure, particularly on other, less resistant varieties suggested by comparison with the findings of van de Wouw grown in proximity. Moreover, these highly resistant varieties et al.(2016), who reported Telfer as having significantly higher may moderate the development of, and the threat from, new races resistance than Hyola 650, whereas in the present study, Hyola or pathotypes of H. parasitica in future, primarily from reduced 650 has a %DI score of 26.6% v. Telfer with a %DI of 50.2%. inoculum loads. However, any such deployment should be This difference in expression of host resistance between studies undertaken only where they have adequate and relevant is unlikely to be due to differences in resistance expression resistance to blackleg disease. between seedlings and adult plants, because seedling and adult None of the varieties tested in the present study were immune plant resistances are correlated (e.g. Zhang et al. 2012). to downy mildew nor did they show any indication of a hypersensitive response. This suggests that the resistance Acknowledgements found in the study is also quantitative and/or continuously The first author gratefully acknowledges a scholarship from the University variable resistance to H. parasitica as shown by others (e.g. of Kufa in Iraq. The authors gratefully acknowledge the financial support of Johnston 1975; Bradshaw et al. 1989; Kumar and Singh 2002). the Grains Research and Development Corporation (GRDC) as this work However, immunity has been reported elsewhere in a few constitutes part of the GRDC UWA 170 project ‘Emerging foliar diseases Brassicas including B. carinata and B. alba (Shivpuri et al. of canola’. We are very grateful to Dr Nash Nashaat, formally of Rothamsted 1997; Saharan and Krishna 1999). Lucas et al.(1988) found Research, UK, for provision of seed of B. napus lines Komet-741 A, Komet- resistance in B. napus (oilseed rape) that was controlled by a 744 A, Cresor-770 B, Cresor-771 B and Capricorn C; and to commercial seed single gene. Yu et al.(2009) found the gene that controlled companies for provision of Australian varieties of B. napus and B. juncea. the resistance to downy mildew in cotyledons in Chinese Exceptional technical support from Robert Creasy and Bill Piasini in the cabbage (B. rapa ssp. pekinensis) by using a phenotypic assay UWA Plant Growth Facilities is also gratefully acknowledged. in conjunction with mapping a major QTL on linkage group A8. Studies to map the QTL associated with the strongest resistance References in the current study may be enlightening. Barbetti MJ, Khangura R (2000) Fungal diseases of canola in Western Although H. parasitica has been present in Australian Australia. Bulletin No. 4406. Agriculture Western Australia, South canola and mustard crops for decades (Barbetti and Khangura Perth, W. 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www.publish.csiro.au/journals/cp Page 44 of 119 Chapter 3
Resistances to downy Mildew (Hyaloperonospora brassicae) in diverse Brassicaceae offer new disease management opportunities for oilseed and vegetable crucifer industries
European Journal of Plant Pathology (submitted 27th May 2018, under review)
This chapter focuses on new sources of resistance to Hyaloperonospora brassicae across diverse Brassicaceae genotypes.
Page 45 of 119 Resistances to downy mildew (Hyaloperonospora brassicae) in diverse Brassicaceae offer new disease management opportunities for oilseed and vegetable crucifer industries
Akeel E. Mohammed1,2 • Ming Pei You1 • Surinder. S. Banga3 • Martin J.
Barbetti1* ()
1 UWA School of Agriculture and Environment and the UWA Institute of
Agriculture, Faculty of Science, The University of Western Australia, Crawley,
WA, 6009, Australia
2 Department of Plant Protection, Faculty of Agriculture, University of Kufa, Najaf,
Iraq
3 Department of Plant Breeding and Genetics, Punjab Agricultural University,
Ludhiana 141004, Punjab, India
*Corresponding author: M. J. Barbetti; E-mail: [email protected]
E-mail: [email protected] (M.J. Barbetti)
Abstract Some 154 Brassicaceae genotypes (78 Brassica napus, 38 B. carinata, 25 B. juncea, three Raphanus sativus, two each of Rapistrum rugosum and B. incana and one each of Crambe abyssinica, B. fruticulosa, Hirschfeldia incana, B. insularis, B. oleracea and Sinapis arvensis), were inoculated with a mixture of seven isolates of Hyaloperonospora brassicae to identify effective host resistances. Many highly resistant genotypes were identified, particularly R.
Page 46 of 119 sativus Krasnodar Market B (%Disease index 6.6) and Pegletta (%DI 9.0); B. carinata Tamn Rex-sel Green (%DI 7.6), BRA926/18 (%DI 9.7) and PI360884
(%DI 9.7); and B. juncea, Ringot1 (%DI 9.7). A further 13 B. carinata, seven B. juncea and single R. sativus (Boss) and B. incana (UPM6563) genotypes were also highly resistant (%DI 11.1), as were B. oleracea CPI106844 (%DI 14.6) and
Crambe abyssinica (%DI 17.4). Almost all B. carinata and B. juncea genotypes showed high resistance (%DI 7.6-22.2). In contrast, B. napus genotypes showed wide ranging responses, from high resistance in SN-8 (%DI 22.2%) to extreme susceptibility in Hyola 450TT and Thunder TT (%DI 83.7, 95.5, respectively). R. rugosum, B. fruticulosa, H. incana and B. insularis genotypes ranged from moderately to highly susceptible (%DI 55.2-78.8). This study highlights the ready availability of very high levels of pathotype-independent resistance across diverse Brassicaceae to H. brassicae, particularly R. sativus, B. carinata, B. juncea, B. oleracea and C. abyssinica. Resistances identified can be utilized as sources of resistance in oilseed and vegetable Brassicaceae breeding programs and/or directly deployed as new varieties where downy mildew is prevalent.
Keywords Hyaloperonospora brassicae, downy mildew, weedy crucifers,
Brassicaceae, Brassica carinata, B. napus, B. juncea, host resistance
Introduction
Downy mildew, caused by Hyaloperonospora brassicae, is a serious foliar disease on oilseed and other Brassicaceae species (Silué et al. 1996; Coelho et al. 2012), adversely affecting production of vegetable oils from Brassica spp. that represent about one-third of the total vegetable oil production worldwide (Kaur et al. 2011). The disease is endemic across Brassica-growing regions worldwide,
Page 47 of 119 including Australia (Howlett et al. 1999; Barbetti and Khangura 2000). While plants can be particularly severely affected by H. brassicae at the seedling stage, downy mildew can also cause systemic symptoms on flowers or non-floral parts of their hosts (Thines and Kummer 2013) and can increase severity of the co- infecting white rust pathogen, Albugo candida, in Brassica juncea (Kaur et al.
2011). Across southern Australia there have been severe outbreaks of downy mildew in oilseed rape, and van de Wouw et al. (2016) found “downy mildew present at most sites assessed, with the site incidence ranging from 100% in
2013 to 52% in 2015.” Australia-wide surveys of oilseed rape crops in 2015 and
2016 showed that the worst affected crops had up to 55% of leaves diseased,
15% of leaf area lost to lesions, and 13% of leaf area collapsed (MJ Barbetti and
MP You unpubl.). Estimated losses up to AUS$13 million occur annually in
Western Australia alone (Mohammed et al. 2017).
The challenge in trying to manage this disease is the rapid epidemic increase of downy mildew (Spencer-Phillips and Jeger 2004), making disease development fast and unforeseen, and this is one of the reasons why control with fungicides or cultural strategies have provided only limited or partial control with high cost (Barbetti et al. 2011; Neik et al. 2017). Hence, various studies have been conducted to locate resistance in Brassica spp. to H. brassicae. For example, there are reported resistances by Natti et al. (1967), Farnham et al.
(2002) and Monteiro et al. (2005) in B. oleracea, by Lucas et al. (1988) and
Nashaat et al. (1997) in B. napus, and by Nashaat and Awasthi (1995), Nashaat et al. (2004) and Chattopadhyay and Séguin-Swartz (2005) in B. juncea.
Resistance in B. napus at the cotyledon stage has also been reported (Ge et al.
2008; Mohammed at el. 2017) as has adult plant resistance in B. oleracea var. italica (Dickson and Petzoldt 1993; Coelho et al. 1998; Jensen et al. 1999a,b;
Page 48 of 119 Wang et al. 2001; Carlsson et al. 2004; Coelho et al. 2009). Dickson and Petzoldt
(1996) studied the quantitative resistance in 49 lines of B. oleracea, whereas
Carlier et al. (2012) found that the Pp523 gene enhances resistance to H. brassicae in adult plants. A recessive gene was reported by Hoser-Krauze et al.
(1991) also in B. oleracea, likewise, others reported single or more-dominant resistance genes (e.g., Moss et al. 1988; Hoser-Krauze et al. 1995; Carvalho and
Monteiro 1996; Vicente et al. 2012). Further, Nashaat et al. (2004) reported a single dominant gene in B. juncea and both Lucas et al. (1988) and Nashaat and
Rawlinson (1994) reported the same for B. napus.
While the above resistances to H. brassicae relate to studies on individual oilseed and vegetable brassicas, an exception to these was the study of
Mohammed et al. (2017) that sought resistances across more than 100 Australian oilseed rape varieties (B. napus and B. juncea) and a small number of diverse
Brassicaceae genotypes of B. carinata, B. nigra, B. rapa, Crambe abyssinica and
Raphanus sativus. As that study highlighted the potential of R. sativus and B. carinata in particular and C. abyssinica to a lesser extent, current studies were undertaken to locate additional new sources of resistance to H. brassicae across diverse oilseed (B. napus, B. carinata, B. juncea, C. abyssinica), vegetable (B. oleracea, R. sativus), and wild or weedy (B. fruticulosa, B. incana, B. insularis,
Hirschfeldia incana, Rapistrum rugosum, Sinapis arvensis) Brassicaceae. The high-level resistances identified against H. brassicae in the current studies offer opportunities to develop new highly resistant commercial varieties, and even direct deployment, where agronomically suitable, into regions where downy mildew is severe. A comparison of the levels of resistance to H. brassicae of studied genotypes in the current study was made with the levels of resistance for the same genotypes against other pathogens, including Sclerotinia (Sclerotinia
Page 49 of 119 sclerotiorum) on cotyledons from Uloth et al. (2013) and on stems from Uloth et al. (2014), and white leaf spot (Pseudocercosporella capsellae) from Gunasinghe et al. (2014).
Materials and Methods
Brassicaceae genotypes
One hundred and fifty four Brassicaceae genotypes were used in the current study, including: 78 B. napus, 38 B. carinata, 25 B. juncea and 13 diverse
Brassicaceae including (three of R. sativus, two each of R. rugosum, B. incana and one each of C. abyssinica, B. fruticulosa, H. incana, B. insularis, B. oleracea and S. arvensis) (Table 1). B. napus Tranby, Hyola 450 TT and Thunder TT were specifically chosen as they are known to be highly susceptible to H. brassicae, while B. juncea Montara, Muscan 963 and JM06018 and R. sativus Boss were specifically chosen as they had shown high-level resistance in the earlier study of Mohammed et al. (2017). Details of origin of test genotypes is given in Table
1.
Hyaloperonospora brassicae isolates
A mixture of seven isolates of H. brassicae was used in this study: six isolates collected from oilseed rape across geographically diverse sites in
Western Australia, viz. isolates WA2, WA5, WA7, WA11, WA29, WA30 and isolate VC36 collected from Victoria, Australia. These isolates were deliberately chosen as they were collected from across a wide geographical area severely
Page 50 of 119 affected by downy mildew in southern Australia during 2015-2016 oilseed rape disease surveys (MP You and MJ Barbetti unpubl.) and tested by Mohammed et al. (2018) to characterize the pathotype structure. All H. brassicae isolates had been obtained from the original field samples by removing hyphal tips using fine tweezers and a dissection microscope and then transferring to cotyledons of 7- day-old seedlings of highly susceptible B. napus cultivars Thunder TT and
Tranby. DNA of all isolates was extracted using the procedure of Cenis (1992) where mycelium from the leaf samples was harvested directly then blotted dry with filter paper. Mycelia were homogenized using Precellys Evolution
Homogenizer (Bertin TECHNOLOGIES), washed with Tris-EDTA (TE) buffer, pelleted by centrifuging at (13,000 rpm for 10 min), and then supernatant discarded. After adding 300 μL of extraction buffer [200 mM Tris-HCl (pH 8.5),
250 mM NaCl, 25 mM EDTA], 0.5% (wt/vol) sodium dodecyl sulphate (SDS)] was added to the pellet. Then, 150 μL of 3 M sodium acetate (pH 5.2) was added and maintained at 20°C for 10 min, followed by vortexing briefly and centrifuging at
13,000 rpm for 5 min. Subsequently, supernatant was transferred to a fresh tube prior to adding an equal volume of cold isopropanol (450 µL) and centrifuged for
13,000 rpm for 10 min and then left to stand at least 15 min at room temperature.
Precipitated DNA was collected by centrifugation for 13,000 rpm for 15 min. Pellet was air dried, then washed with 70% (vol/vol ethanol:water) and resuspended in
50 μL of TE buffer. Quantity and the quality of extracted DNA was determined with a NanoDrop 1000 Spectrophotometer (Thermo Scientific). DNA was stored at 4°C. The DNA was subjected to PCR using a master mix of a total volume of
(50 µL) that contained 0.2 µM of each primer (primers ITS1-O (5’-CGG AAG GAT
CAT TAC CAC-3’; Bachofer 2004) and ITS4-H (5’-TCC TCC GCT TAT TAA TAT
GC-3’; Göker et al. 2004), a modification of ITS4. ITS1-O was chosen as the
Page 51 of 119 specificity of it greatly reduces the problem of additional amplification of host ITS rDNA (Bachofer 2004; Göker et al. 2009). PCR was undertaken as follows: Initial denaturation 94°C for 2 min; followed by 35 cycles at 94°C for 1 min, with annealing gradient temperature set at range 50°C-60°C for 1 min and extension at 72°C for 2 min; followed by a final extension step at 72°C for 7 min and then held at 4°C. Final PCR was run using the selected best annealing temperature of
53°C. PCR products were subjected to agarose gel electrophoresis at 50 V for
120 min on a 1% (w/v) agarose gel containing 0.1% GelRed™ Biotium Inc.
(United States) and then visualised under UV light. Aliquots of PCR products of
830 bp (30 μL of each) were sent to Macrogen Inc. (Korea) for sequencing.
Finally, sequences of all isolates were confirmed as H. brassicae using BLAST in
GenBank (>98% similarity).
Each H. brassicae isolate was maintained separately in isolated containers on cotyledons of 7-day old seedlings of susceptible B. napus Thunder
TT and Tranby (Ge et al. 2008; Mohammed et al. 2017), in a controlled environment room with a 12 h photoperiod, light intensity 420 µmol.m-2s-1 at 18⁰C day and 13⁰C night. Equal numbers of cotyledons that supported abundant sporulation by each of the seven different pathogen isolates were collected and placed in 50 mL distilled water in a 100 mL Erlenmeyer flask. The flask was shaken to dislodge the zoosporangia. The resulting suspension was filtered through one layer of cheesecloth to remove any cotyledon contamination and the concentration adjusted to 105 zoospores mL-1 using a haemocytometer counting chamber. This range of representative isolates of H. brassicae were chosen and mixed as distinct pathotypes of H. brassicae occur across southern Australia
(Mohammed et al. 2018), hence avoiding any possible specific pathotype-variety interactions that could otherwise complicate test outcomes.
Page 52 of 119
Inoculation
Four seeds of each Brassicaceae test genotype were sown in 4 x 4 cm seedling tray cells and thinned to two plants per cell following seedling emergence. Genotypes were grown at 18⁰C day and 13⁰C night in a controlled environment room with a day length of 12 h. Plants were watered daily with deionised water and allowed to drain to field capacity. At growth stage 1.00
(Sylvester-Bradley and Makepeace 1984) 7 d after seedling emergence, seedling trays were placed in 790 mm (L), 390 mm (W), 155 mm (H) clear plastic boxes with lids where seedlings were inoculated with the 105 zoosporangia mL-1 suspension of the mixture of the seven H. brassicae isolates, using a micropipette to place a 10 µL droplet of the sporangial suspension on to each of two lobes of both cotyledons of each seedling. High humidity was maintained for 24 h post- inoculation by hand-misting the walls and lids of each plastic box with deionized water and then keeping the clear plastic lid on for 24 h. Subsequently, lids were removed and plants maintained by daily hand-watering for 6 d; then the plastic boxes were covered again for a further 24 h to maintain a second period of high humidity.
Disease assessment
The procedure of Williams (1985) was used to assess disease severity on plants at 7 d post-inoculation (dpi) using a modified 0-9 scale, where: 0 = no symptoms or sign of downy mildew disease; 1 = minute scattered necrotic flecks under the inoculum drop or inoculated plant surface area, no sporulation; 2 =
Page 53 of 119 larger necrotic flecks under the inoculum drop or inoculated plant surface area, no sporulation; 3 = very sparse sporulation, one to a few conidiophores, necrotic flecking but often with tissue necrosis evident; 5 = sparse sporulation, tissue necrosis; 7 = abundant sporulation, tissue necrosis and chlorosis may be present; and, 9 = heavy sporulation, cotyledon or leaf tissue collapse. Subsequently, 0-9 disease severity scores for each seedling tray cells were converted into a disease index (%DI) based on the methods described by McKinney (1923), where:
%DI = {[(a × 0) + (b × 1) + (c × 2) + (d × 3) + (e × 4) + ……. (j x9)] × 100} / [(a + b + c + d + ……j) × 10)]
And where a, b, c, d, e ……j are the number of plants with disease severity scores of 0, 1, 2, 3, 4, …..9, respectively.
Experimental design and statistical analyses
The experiment was arranged in a randomised complete block design, with four replicates of each treatment. The experiment was repeated once. The relationship between the initial and repeat experiments was assessed using a paired t test using GenStat (Release 14.2; 18th Edition, Lawes Agricultural Trust,
Rothamsted Research, UK) and for homogeneity of variances across the original and repeat experiments using Bartlett’s test (Snedecor and Cochran 1989). In each case, there were no significant differences between the original and repeat experiments (P > 0.05) using the t test, and variances were similar using Bartlett’s test. Therefore, the data sets were pooled, reanalysed, and presented as a single data set. Fisher’s least significant differences (LSD) were used to separate significant differences between genotypes. The test genotypes were listed according to their level of resistance measured by %DI% from lowest to highest
Page 54 of 119 where genotypes with %DI 5.6-22.2 were considered highly resistant, 29.9-39.9 resistant, 41.3-50 moderately resistant, 51-60.1 moderately susceptible, 61-75 susceptible, and >75 highly susceptible (Table 1).
In addition, a set of data was extracted from Uloth et al. (2013) for cotyledon resistance to S. sclerotiorum, from Uloth et al. (2014) for field stem resistance to S. sclerotiorum, and from Gunasinghe et al. (2014) for resistance to
P. capsellae, to compare the expressions of resistance of the studied different
Brassicaceae genotypes to H. brassicae with that against these other pathogens.
Results
There was no disease at any stage on the water-only inoculated controls
(Fig. 1A). Expression of host resistance ranged from typical highly susceptible reaction like for genotypes B. napus Hyola 450 TT and H. incana Hin37 (Fig. 1B,
C) to a typical highly resistant reaction for genotypes such as B. juncea
Hetianyoucai (Fig. 1D), B. carinata Ethiopia B (Fig. 1E) and R. sativus Boss (Fig.
1F). Among the test genotypes there were significant differences (P ≤ 0.001)
(Table 1). The general range in levels of resistance expressed across the three main Brassica species showed that the 25 B. juncea and 38 B. carinata genotypes showing best overall resistance in comparison with the generally more susceptible 78 B. napus genotypes (Fig. 2). Further, there were overall different ranges in comparative resistance responses in accordance to the country of origin (Australia, China, Ethiopia, India) as indicated by %DI values (Fig. 3). In particular, Ethiopian genotypes (all B. carinata) overall had least disease, with no highly susceptible genotypes. In contrast, China genotypes (30% B. juncea and
70% B. napus), Australian genotypes (mainly B. napus and other species in small
Page 55 of 119 portion) and Indian genotypes (40% B. juncea and 60% B. napus) showed a wide range in host response from highly resistant to highly susceptible (Fig. 3).
R. sativus Krasnodar Market B and Pegletta were highly resistant with %DI values 5.6, and 9.0, respectively, followed by B. incana UPM 6563 (%DI 11.1), B. oleracea CPI 106844 (%DI 14.6) and C. abyssinica (%DI 17.4). All genotypes of
B. carinata were highly resistant: Tamn Tex-sel Green (%DI 7.6), BRA 926/81 and PI 360884 (%DI 9.7), 045103, 054099, 054101, 054106, CPI 100551, CPI
99838, Ihanja, Karate, ML-EM-1(Rungwe), PI 195552, ST 27, ST 57,TZ-SMN-
36-5 and USD-14 (%DI 11.1), BRA 927/72 and Field Station (%DI 11.8), Ethiopia
B and ML-EM-8 (GKK 70) (%DI 12.5), Mbeya green, Peela Raya and PI 193459
(%DI 13.2), USD-13 (%DI 13.9), Brown Raya (%DI 14.6), BRA 1028/79, INIA
0572-69, SMP 3-82 and TZ-SMN-61-4 (%DI 15.3), TZ-SMN-46-7 (%DI 16),
Ethiopian mustard, Addis A (%DI 17.4), ML-EM-3 (18.1%), 054104 and Mbeya purple (%DI 18.8), ML-EM-7 (%DI 20.1), and Chembere Dzagwinhanha (%DI
22.2). The exception within B. carinata was the highly susceptible TZ-SMN-1-1
(%DI 61.8). B. juncea genotypes were also generally highly resistant; for example, Ringot 1 (%DI 9.7), Ashirwad, CBJ-004, Geeta, Kanti, Kranti, Maya,
Swarna Jyoti and Vaibhav (%DI 11.1), Hetianyoucai and Varuna (%DI 11.8),
Laxmi (%DI 12.5), Basanti and Jinshahuang (%DI 13.2), Narendra Ageti, Prakash and Pusa Mahak (%DI 13.9), Tunhiuhuangjie (%DI 14.6), Rohini (%DI 15.3), Sej-
2 (%DI 16), Yilihuang (%DI 16.7) and Vasundhra (%DI 18.8). Genotypes of B. napus showed widely differing responses. For example, B. napus SN-8 was highly resistant (%DI 22.2), while there were varying levels of resistance expressed across YM 05 (%DI 29.9), NS-2 (%DI 31.6), RMNL-20 (%DI 31.9), SN-
2 and YM 06 (%DI 33.3), Charlton-NCA-28 (%DI 34), NS-9 (%DI 36.1), Mystic-
NCA-1 (%DI 37.2), Charl1DN-NCA-18 (%DI 37.9), RMNL-14 (%DI 38.2), SN-3
Page 56 of 119 (%DI 38.9), RMNL-11 (%DI 39.6) and SN-6 (%DI 39.9); compared to the 20 highly susceptible B. napus genotypes with %DI values >60 and up to 83.7 and 95.5 for
B. napus Hyola 450 TT and Thunder TT, respectively. S. arvensis SAR 1 also showed resistance to H. brassicae (%DI 31.9). There was a wide range in host responses across the remaining test genotypes from partially resistant, to susceptible and to highly susceptible (Table 1).
Comparisons were also made with resistances expressed for some of these same genotypes in earlier studies for Sclerotinia (S. sclerotiorum) on seedling cotyledons in controlled environment studies, and on stems and leaves for white leaf spot caused by (P. capsellae) in field studies (Table 1). These comparisons showed that some genotypes highly resistant to H. brassicae also showed no disease symptoms (e.g., B. carinata 054099, 054101 and BRA
927/72), high resistance (e.g., B. carinata BRA 926/81 and PI 195552) or moderate resistance (e.g., B. carinata PI 360884) to P. capsellae leaf disease; and/or showed high resistance (e.g., R. sativus var. oleiformis Krasnodar Market
B, B. carinata BRA 927/72 and PI 193459), resistance (e.g., B. carinata 054099) or moderate resistance (e.g., B. carinata PI 195552) to S. sclerotiorum stem disease; and/or showed moderate resistance (e.g., B. carinata BRA926/81 and
PI 193459, and R. sativus Boss) to S. sclerotiorum seedling cotyledon disease
(Table 1).
Discussion
This study highlights very high levels of pathotype-independent resistance that is readily available across diverse Brassicaceae genotypes to H. brassicae, particularly R. sativus, B. carinata B. juncea, B. oleracea and C. abyssinica.
Page 57 of 119 Pathotype-independent resistances are of high valuable against H. brassicae as there are distinct pathotypes of this pathogen present in southern Australia
(Mohammed et al. 2018). These host resistances identified offer new opportunities to improve the management of this important pathogen across the oilseed and vegetable Brassicaceae industries; first, by their utilization as sources of resistance in breeding programs to produce resistant varieties, and, second, by the fact that in many instances they could be directly deployed as new varieties where downy mildew is prevalent. Using a mixture of a range of representative isolates of H. brassicae for the current study meant that host resistances identified were pathotype-independent, critically important in a pathogen like H. brassicae that shows strong sub-specific variation (Barbetti et al. 2012). This approach aligns well with the drive over the past decade to identify pathotype- independent host resistance for diseases of Brassicas, such as Sclerotinia rot
(Barbetti et al. 2014) and white leaf spot disease (Eshraghi et al. 2007;
Gunasinghe et al. 2016a, b). Our approach ensures that many of the pathotype- independent resistances to H. brassicae identified in the current study are of immediate and widespread relevance for commercial deployment in both horticultural and broad-acre agricultural Brassica industries. Importantly, while the present study involved cotyledons tests, expression of resistance on cotyledons is generally similar as on leaves under controlled (Wang et al. 2000) or field conditions (Jensen et al. 1999a).
In the current study, R. sativus Krasnodar Market B and Pegletta were both highly resistant. High level resistance has also been reported by Satou and
Fukumoto (1996), Silué et al. (1996) and Mohammed et al. (2017) for different genotypes within R. sativus. Also, Li et al. (2011) highlighted a hypersensitive reaction when cotyledons of R. sativus were inoculated by different H. brassicae
Page 58 of 119 isolates. As a range of resistance responses, including high level resistance, have been reported within R. sativus against Sclerotinia rot (Uloth et al. 2013; Ge et al.
2015), it is likely that R. sativus offers resistances across these different pathogens. However, it is clear that these disease resistances are not universal across R. sativus genotypes as they can be extremely susceptible to white leaf spot disease (e.g., Gunasinghe et al. 2014).
Also in the current study, genotypes of B. juncea, all but one of the genotypes of B. carinata, along with B. incana UPM 6563, B. oleracea CPI
106844 and C. abyssinica, were also highly resistant to H. brassicae. Resistance to downy mildew in B. carinata has also been reported by Saharan and Krishnia
(2001), Li et al. (2011), Mehta (2014) and Mohammed et al. (2017) and resistance in B. juncea reported by Nashaat and Awasthi (1995), Nashaat et al. (2004) and
Chattopadhyay and Séguin-Swartz (2005). The high level resistance of B. juncea accessions in the current study provides a possible explanation for the field scenario in Australia where mustard crops are free of downy mildew while fields of B. napus growing nearby are severely infected (MP You and MJ. Barbetti unpubl.). This latter conclusion is also supported by the fact that while isolates of
H. brassicae are most virulent on their species of origin, those obtained from different Brassica spp. can infect related species (Channon 1981; Lucas et al.
1988; Satou 2000). That the expression of resistance on cotyledons to H. brassicae generally reflects resistance in the adult stage (Jensen et al. 1999a;
Wang et al. 2000; Zhang et al. 2012) provides confidence in these new sources of resistance, as highlighted in the current study, for field deployment.
This study also highlighted the wide variation in host response for B. napus genotypes, a few with resistance but with the majority being susceptible or highly susceptible. Best was the highly resistant B. napus SN-8 followed by YM 05, NS-
Page 59 of 119 2, RMNL-20, SN-2 and YM 06, Charlton-NCA-28, NS-9, Mystic-NCA-1,
Charl1DN-NCA-18, RMNL-14, SN-3, RMNL-11 and SN-6 that showed resistance but at a lower expression. Resistance in B. napus to H. brassicae has been reported (e.g., Lucas et al. 1988; Nashaat et al. 1997; Ge et al. 2008; Mohammed et al. 2017) and it has been suggested that the B. napus genotypes showing high resistance have one or more major genes (Kimber 1981; Thompson and Hughes,
1986). Some B. napus genotypes highlighted as resistant in the present study are agronomically suitable as commercial oilseed rape varieties that could be deployed in downy mildew prone regions, particularly so as in recent years when downy mildew has become more prevalent, possibly a consequence of climate variability that strongly affects fungal and oomycete disease epidemics in oilseed rape (Barbetti et al. 2012).
The general range in levels of resistance expressed across the three main oilseed Brassica species in the current study showed that B. juncea and B. carinata had better overall resistance in comparison with B. napus. It is noteworthy that all B. juncea and most B. carinata genotypes used in this study were almost universally highly resistant regardless of their origin (Australia, India,
China, Africa or Pakistan). Interestingly, Ethiopian genotypes (all B. carinata) were most resistant in terms of country of origin, with no susceptible genotypes.
In contrast, Chinese genotypes (30% B. juncea and 70% B. napus), Australian genotypes (mainly B. napus with other species in small portion) and Indian genotypes (40% B. juncea and 60% B. napus) showed a wider range in host responses from highly resistant to some that were highly susceptible. As such, the current studies highlight both the already wide availability of high level resistance within B. carinata and B. juncea and also the need to now focus on locating similar high level resistances within B. napus and/or crossing resistances
Page 60 of 119 from B. carinata and B. juncea into B. napus to improve resistance in this latter species. The critical need for improved resistance in B. napus was recently demonstrated by Mohammed et al. (2017) who highlighted the general paucity of effective resistance in current Australian oilseed rape varieties, with very few exceptions such as CBTM Tanami, ATR-Banjo and Hyola 575 CL.
Comparisons made with resistances expressed for some of these same genotypes in earlier studies for Sclerotinia rot caused by (S. sclerotiorum) on cotyledons in controlled environment studies (Uloth et al. 2014) and on stems in field studies (Uloth et al. 2013) and on leaves for white leaf spot (P. capsellae) in field studies (Gunasinghe et al. 2014) highlighted genotypes highly resistant to
H. brassicae that also showed resistance to one or both of these other pathogens.
Particularly outstanding examples included genotypes highly resistant to H. brassicae in the current study that showed no disease symptoms (e.g., B. carinata 054099, 054101 and BRA 927/72) or high resistance (e.g., B. carinata
BRA 926/81 and PI 195552) to P. capsellae leaf disease; and/or showed high resistance (e.g., R. sativus var. oleiformis Krasnodar Market B, B. carinata BRA
927/72 and PI 193459), or resistance (e.g., B. carinata 054099) to S. sclerotiorum stem disease. These genotypes with resistances to multiple pathogens will be of greatest value to breeding programs in developing new Brassica varieties with combined resistances to a range of important but co-occurring diseases.
Managment currently relies upon chemical and/or cultural control options that generally only provide erratic and/or inadequate control (Barbetti et al. 2011), particularly as such options are generally applied to varieties with little resistance to downy mildew. Hence, there is an urgent need to now commercially utilise the new host resistances identified in the current study. Genotypes with high-level resistance will be of greatest value for developing new resistant varieties of
Page 61 of 119 oilseed, forage and vegetable crucifers. However, even some of the more moderate levels of resistance identified, provided they are in genotypes with agronomically suitable backgrounds enabling direct deployment commercially, offer significant prospect for improving current management of downy mildew.
Finally, it is likely that at least some of the resistances identified in the current study across diverse cruciferous species will constitute new sources and/or types of host resistance not previously identified.
Acknowledgements
The first author is grateful to the scholarship from the University of Kufa in Iraq.
The authors are grateful for some additional financial support of the Grains
Research and Development Corporation (GRDC) as this work is aligned with part of the GRDC UWA170 project ‘Emerging foliar diseases of canola’. We are very grateful to commercial seed companies for provision of Australian varieties of B. napus and B. juncea and to Huang Yi, Oil Crops Research Institute, Chinese
Academy of Agricultural Sciences, Wuhan, China, for seed of some Chinese genotypes. The exceptional technical support from Robert Creasy and Bill Piasini in the UWA Plant Growth Facilities is also gratefully acknowledged. No author has any conflict of interest to declare.
Compliance with ethical standards
Ethical statement: This research did not involve any animal and/or human participants.
Conflict of interest: The authors declare that they have no conflict of interests.
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Page 71 of 119 Table 1 Germplasm origin and percent disease index (%DI) values on cotyledons of 154 Brassicaceae genotypes (78 Brassica napus, 38 B. carinata, 25 B. juncea and 13 miscellaneous Brassicaceae) inoculated with mixture of seven different isolates of Hyaloperonospora brassicae. Comparative host resistances to
Sclerotinia (Sclerotinia sclerotiorum) on cotyledons (controlled environment;
Uloth et al. 2014) and stems (field; Uloth et al. 2013) and to white leaf spot (field;
Pseudocercosporella capsellae) on leaves are also shown.
Downy Downy Sclerotinia Sclerotinia White leaf Brassica sp. Genotype Origin mildew mildew cotyledon stem spot DI% resistance resistancea resistanceb resistancec Raphanus HR -d HR S sativus var. Krasnodar Market B Australia 5.6 oleiformis Brassica HR - - - Tamn Tex-sel Green Unknown 7.6 carinata R. sativus Pegletta Italy 9 HR - MS HS B. carinata BRA 926/81 Ethiopia 9.7 HR MR HR R B. carinata PI 360884 Ethiopia 9.7 HR - S MR B. juncea Ringot 1 China 9.7 HR - MS R B. carinata 45103 Ethiopia 11.1 HR - - - B. carinata 54099 Ethiopia 11.1 HR - R Immune B. carinata 54101 Ethiopia 11.1 HR - S Immune B. carinata 54106 Ethiopia 11.1 HR S HS - B. juncea Ashirwad India 11.1 HR - - - R. sativus Boss Italy 11.1 HR MR S S B. juncea CBJ-004 China 11.1 HR - - - B. carinata CPI 100551 Australia 11.1 HR - - - B. carinata CPI 99838 Australia 11.1 HR - - - B. juncea Geeta India 11.1 HR - - - B. carinata Ihanja Tanzania 11.1 HR - - - B. juncea Kanti India 11.1 HR - - - B. carinata Karate Ethiopia 11.1 HR - - - B. juncea Kranti India 11.1 HR - - S B. juncea Maya India 11.1 HR - - - B. carinata ML-EM-1 (Rungwe) Malawi 11.1 HR - - - B. carinata PI 195552 Ethiopia 11.1 HR - MR R B. carinata ST 27 Tanzania 11.1 HR - - - B. carinata ST 57 Tanzania 11.1 HR - - - B. juncea Swarna Jyoti India 11.1 HR - - - B. carinata TZ-SMN-36-5 Tanzania 11.1 HR - - - B. incana UPM 6563 Australia 11.1 HR S - - B. carinata USD-14 Unknown 11.1 HR - - - B. juncea Vaibhav India 11.1 HR - - HS B. carinata BRA 927/72 Ethiopia 11.8 HR - R Immune B. carinata Field station Tanzania 11.8 HR - - - B. juncea Hetianyoucai China 11.8 HR - - - B. juncea Varuna India 11.8 HR - - S B. carinata Ethiopia B Ethiopia 12.5 HR - - - B. carinata ML-EM-8 (GKK 70) Malawi 12.5 HR - - - B. juncea Laxmi India 12.5 HR - - - B. juncea Basanti India 13.2 HR - - - B. juncea Jinshahuang China 13.2 HR - - - B. carinata Mbeya green Tanzania 13.2 HR - - - B. carinata Peela Raya Pakistan 13.2 HR - S - B. carinata PI 193459 Ethiopia 13.2 HR MR HR - B. juncea Narendra Ageti India 13.9 HR - - - B. juncea Prakash India 13.9 HR - - HS B. juncea Pusa Mahak India 13.9 HR - - -
Page 72 of 119 B. carinata USD-13 Unknown 13.9 HR - - - B. carinata Brown Raya Pakistan 14.6 HR - MS - B. oleracea CPI 106844 Australia 14.6 HR - MR S B. juncea Montara China 14.6 HR - MS S B. juncea Muscan 963 Australia 14.6 HR - - - B. juncea Tunhiuhuangjie China 14.6 HR - - - B. carinata BRA 1028/79 Ethiopia 15.3 HR - - - B. carinata INIA 0572-69 Spain 15.3 HR - S HS B. juncea Rohini India 15.3 HR - - HS B. carinata SMP 3-82 Pakistan 15.3 HR - HS HS B. carinata TZ-SMN-61-4 Tanzania 15.3 HR - - - B. juncea JM06018 Australia 16 HR - HS HS B. juncea Sej-2 India 16 HR - - MS B. carinata TZ-SMN-46-7 Tanzania 16 HR - - - B. juncea Yilihuang China 16.7 HR - - - Crambe HR - - HR Crambe Australia 17.4 abyssinicia Ethiopian mustard, HR - - - B. carinata Ethiopia 17.4 Addis A B. carinata ML-EM-3 Malawi 18.1 HR - - - B. carinata 054104 Ethiopia 18.8 HR HS R Immune B. carinata Mbeya purple Tanzania 18.8 HR - - - B. Juncea Vasundhra India 18.8 HR - - - B. carinata ML-EM-7 Malawi 20.1 HR - - - Chembere HR - - - B. carinata Ethiopia 22.2 Dzagwinhanha B. napus SN-8 India 22.2 HR - - - B. napus YM 05 China 29.9 R - S MR B. napus NS-2 India 31.6 R - - - B. napus RMNL-20 India 31.9 R - - - Sinapis R - - - SAR 1 Australia 31.9 arvensis B. napus SN-2 India 33.3 R - - - B. napus YM 06 China 33.3 R - MR MR B. napus Charlton-NCA-28 Australia 34 R - - - B. napus SN-9 India 36.1 R - - - B. napus Mystic-NCA-1 Australia 37.2 R - - - B. napus Charl1DN-NCA-18 Australia 37.9 R - - - B. napus RMNL-14 India 38.2 R - - - B. napus SN-3 India 38. 9 R - - - B. napus RMNL-11 India 39.6 R - - - B. napus SN-6 India 39.9 R - - - B. napus Surpass-400-NCA-5 Australia 41.3 MR - - - B. napus SN-1 India 41.7 MR - - - B. napus RMNL-7 India 42 MR - - - B. napus SN-4 India 42 MR - - - B. napus YM 01 China 42 MR - HS - B. napus NS-8 India 43.4 MR - - - B. napus Rainbow-NCA-8 Australia 43.8 MR - - - B. napus SN-7 India 43.8 MR - - - B. napus NS-1 India 44.4 MR - - - B. napus SN-5 India 44.4 MR - - - B. napus RMNL-2 India 45.8 MR - - - B. napus Charl1DN-NCA-48 Australia 46.2 MR - - - B. napus RMNL-10 India 46.5 MR - - - B. napus RMNL-8 India 46.5 MR - - - B. napus Zy 004 China 46.5 MR R MS MS B. napus Mystic-NCA-13 Australia 47.6 MR - - - B. napus Mystic-NCA-14 Australia 48.3 MR - - - B. napus RMNL-9 India 48.3 MR - - - B. napus Surpass-400-NCA-1 Australia 48.3 MR - - - B. napus NS-5 India 48.6 MR - - - B. napus RMNL-18 India 50 MR - - - B. napus RMNL-4 India 51 MS - - - B. napus SN-10 India 51.7 MS - - - B. napus Charlton Australia 52.1 MS R S S B. napus YM 04 China 52.1 MS R HS MS B. napus RMNL-12 India 52.4 MS - - - B. napus RMNL-1 India 52.8 MS - - - B. napus Surpass-400-NCA-18 Australia 53.1 MS - - -
Page 73 of 119 B. napus Ding 110 China 53.8 MS - - - B. napus NC-1 India 53.8 MS - - MR B. napus NS-4 India 53.8 MS - - - B. napus RMNL-19 India 54.2 MS - - - B. napus Charl1DN-NCA-2 Australia 54.5 MS - - - B. napus NS-9 India 54.9 MS - - - B. napus RMNL-16 India 55.2 MS - - - Rapistrum MS - MS MR RRU 24 Australia 55.2 rugosum B. napus NC-2 India 56.6 MS - - - B. napus Mystic-NCA-2 Australia 56.9 MS - - - B. napus RMNL-13 India 57.3 MS - - - B. napus ZY001 China 57.3 MS - - - B. napus NC-3 India 58.3 MS - - - B. napus RMNL-17 India 58.7 MS - - - B. fruticulosa BFR 6 Australia 59.4 MS - MR - B. napus RMNL-5 India 59.4 MS - - - B. napus YM 03 China 59.7 MS - S MS B. napus Fan 189 China 60.1 S - - MR B. napus Fan 23 China 60.1 S - - MS B. napus RMNL-15 India 60.1 S - - - B. napus NC-4 India 61.1 S - - R B. napus ZY006 China 61.1 S - R R B. napus NS-3 India 61.5 S - - - B. napus Rainbow-NCA-14 Australia 61.5 S - - - B. napus ZY003 China 61.5 S - - - B. carinata TZ-SMN-1-1 Tanzania 61.8 S - - - B. napus Fan 028 China 62.2 S - - R B. napus YM 02 China 62.2 S - HR MR B. napus NS-10 India 62.5 S - - - B. napus Surpass-400.NCA-15 Australia 62.5 S - - - B. napus Zhongshu-ang No.4 China 62.5 S - - MR B. napus 06-P71-2 China 64.9 S R HS MS R. rugosum RRU 7 Australia 64.9 S - S - B. napus Yu 178 China 66.7 S - - - B. napus Mystic Australia 68.4 S S HR S B. napus Tranby Australia 70.5 S - - MR B. insularis Moris Unknown 73.3 S - - - B. napus Ding 474 China 74.7 S - - MR Zhongshu You-za S - - MS B. napus China 75 No.8 B. incana Ten Unknown 78.1 HS - - - Hirschfeldia HS - - - Hin 37 Australia 78.8 incana B. napus Hyola 450 TT Australia 83.7 HS - - - B. napus Thunder TT Australia 95.5 HS - - -
Significance of genotype differences for %DI, P < 0.001; LSD at P < 0.05 = 5.73 a Resistances derived from Uloth et al. (2014) b Resistances derived from Uloth et al. (2013) c Resistances derived from Gunasinghe et al. (2014) d - Represents no comparison data available in relevant reference for that genotype
Page 74 of 119
Fig. 1 Showing A, inoculation method of inoculation droplets containing
Hyaloperonospora brassicae conidia onto both cotyledon lobes of both cotyledons of each seedling; B, typical highly susceptible genotype Brassica napus Hyola 450 TT and C, Hirschfeldia incana Hin37; D, typical highly resistant genotype Brassica juncea Hetianyoucai, E, highly resistant genotype Brassica carinata Ethiopia B and F, highly resistant genotype Raphanus sativus Boss.
Page 75 of 119
704 Eur J Plant Pathol (2018) 151:703 711
Introduction Felton and Walker (1946) reported that on leaves of cabbage (B. oleracea), H. parasitica germinates and Oilseed Brassicas are a rich source of oil for human penetrates at 6-24 °C, colonizes at 8-16 °C, and sporu- consumption and of protein in a range of animal feeds lates at 4-24 °C (optimum 12–16 °C), the latter optimum (Si et al. 2003). In Australia, the area grown to canola confirmed by Hartmann et al. (1983) at 13-18 °C. Achar has nearly doubled from 1.4 million ha in 2009 to 2.4 (1998), however, showed H. parasitica germination on million ha in 2013 (Elliott et al. 2015). However, canola cabbage was greatest at 20 °C with 100% relative hu- in Australia faces challenges of increased incidence and midity; while Kofoet and Fink (2007) noted that on severity of some diseases, for example white leaf spot radish (Raphanus sativus) downy mildew was observed (Pseudocercosporella capsellae) (Gunasinghe et al. from 8.3–26.7 °C. Sangeetha and Siddaramaiah (2007) 2016), Sclerotinia (Sclerotinia sclerotiorum) and downy reported that downy mildew occurs across temperatures mildew (Hyaloperonospora parasitica),andthese 14-29 °C. However, there have been no studies specif- increases have coincided with warmer temperatures ically defining the role of temperature on the develop- across southwestern Western Australia over the past ment of downy mildew epidemics on either canola decade and a half (Barbetti et al. 2012; Bureau of (B. napus) or mustard (B. juncea). Meteorology 2014; Uloth et al. 2015). In addition to temperature, plant growth stage or plant Downy mildew (Hyaloperonospora parasitica)isa age also affects host susceptibility to H. parasitica serious disease on canola and other oilseed brassicas in (Coelho et al. 2009). For example, broccoli (B. oleracea Europe (Paul et al. 1998), China and Japan (Satou and var. italica) leaves on plants with >eight leaves were less Fukumoto 1996), North America (Le Beau 1945; susceptible than cotyledons (Coelho and Monteiro 2003). Laemmlen and Mayberry 1984), India (Nashaat et al. However, there are no studies defining the role of plant 2004) and also in Australia (Howlett et al. 1999; age on the development of downy mildew epidemics Barbetti and Khangura 2000). There have been particu- specifically on oilseed B. napus or B. juncea. Hence, larly destructive outbreaks of downy mildew on canola studies were undertaken to define the effects of tempera- crops in Western Australia, especially when it occurs at ture (14/10 °C and 22/17 °C day/night) and plant age the seedling stage (Barbetti and Khangura 2000;Ge (15, 23, 31 and 40 day-old plants) on severity of et al. 2008). However, it can still significantly reduce downy mildew on oilseed B. napus and B. juncea yield and quality even when plants are infected at a later in order to explain the increased incidence and stage of growth (Silué et al. 1996; Coelho et al. 2012). severity of downy mildew on seedlings in Western Downy mildew develops initially on lower leaves and Australia over the past decade and a half when season subsequently progresses upwards on younger leaves temperatures have been increasing as a consequence of (Natti et al. 1956; Coelho et al. 1998; Jensen et al. climate change. 1999). A survey in 2004 (Oilseeds Industry Association of Western Australia, unpublished), estimated the eco- nomic losses from downy mildew on canola to be as Materials and methods much as AU$13 million annually in Western Australia alone. Its importance in Western Australia was again Brassica varieties and experimental environments recently highlighted during Australia-wide surveys across southern Australia in 2015 and 2016 canola Six Brassica cultivars, including B. juncea Montara, growing seasons; particularly in Western Australia B. napus Atomic, ATR-Hyden, Hyola 432, Hyola 450 where worst affected crops showed up to 55% of leaves TTand Thunder TT, were used in the temperature study; diseased, 15% of leaf area lost to lesions, and 13% of but only three cultivars, B. juncea Dune, B. napus Sur- leaf area collapsed from downy mildew (MP You and pass 402 and Hyola 450 TT, were used in the plant age MJ Barbetti, unpublished). study. These cultivars were selected according to a H. parasitica is known to be favoured by warm days previous study (Mohammed et al. 2017)thatshowed (20–24 °C), cool nights (8–16 °C), and a relative hu- they had varying levels of resistance/susceptibility to midity >80% (Channon 1981). Temperature affects co- H. parasitica as follows: (highly resistant: B. napus nidial germination, formation of the appressoria and the Hyola 432 and B. juncea Montara and Dune; resistant: rate of hyphal penetration into the host (Chu 1935). B. napus ATR-Hyden and Atomic; and highly
Page 80 of 119 Eur J Plant Pathol (2018) 151:703 711 705 susceptible: B. napus Thunder TT, Hyola 450 TT). For pellet was vacuum dried for several minutes after wash- all experiments, seedlings were initially germinated ing with 70% (vol/vol) ethanol and resuspended in 50 μl and grown in a controlled environment room of TE buffer. The concentration and quality of the ex- maintained at 13 °C night and 18 °C day, with a tracted DNA was determined with a NanoDrop 1000 12 h photoperiod, and with light source of LED Spectrophotometer (Thermo Scientific). The DNA was cool white and incandescent light bulbs with a stored at 4 °C until required. Then, primers ITS1-O (5’- combined light intensity of 420 μmol.m 2 s 1. CGG AAG GAT CAT TAC CAC; Bachofer 2004)and For the plant age study, plants were maintained ITS4-H (5’-TCC TCC GCT TAT TAATAT GC; Göker under these conditions throughout. However, for et al. 2004), a modification of ITS4, were used for PCR the temperature study, plants were shifted at and sequencing. ITS1-O was chosen as the specificity of growth stage 1.00 (Sylvester-Bradley and Makepeace it greatly reduces the problem of additional amplifica- 1984; at seven days after seedling emergence) into tion of host ITS rDNA (Göker et al. 2009;Bachofer controlled environment rooms maintained at the test 2004). Finally, sequences of all isolates was confirmed temperatures (22/17 or 14/10 °C day/night, 12 h photo- as H. parasitica using BLAST in GenBank (>98% period). Plants were watered daily with deionized water similarity). Each H. parasitica isolate was maintained and allowed to drain to field capacity. To maintain separately in isolated containers on cotyledons of 7-day adequate nutrition, Polyfeed Vegetative Booster (Haifa old seedlings of susceptible B. napus Thunder TT and Chemicals Ltd) was applied using a watering can at Tranby (Ge et al. 2008;Mohammedetal.2017), in a weekly intervals. controlled environment room with a 12 h photoperiod, light intensity 420 μmol m 2 s 1 at 13 °C night and18 °C H. parasitica isolates day. Equal numbers of cotyledons that supported abundant sporulation by each of the 10 different A mixture of 10 isolates of H. parasitica was used in this pathogen isolates were collected and placed in study: nine isolates collected from canola across geo- 50 ml distilled water in a 100 ml Erlenmeyer flask. graphically diverse sites in Western Australia, viz. iso- The flask was shaken to dislodge the conidia. The lates WA5, WA7, WA11, WA12, WA13, WA22, WA29, resulting suspension was filtered through one layer of WA30 and WA49, and an isolate SA78 collected from cheesecloth to remove any cotyledon contamination and South Australia. These isolates were deliberately chosen the concentration adjusted to 105 zoosporangia ml 1 as they were collected from across a wide geographical using a haemocytometer counting chamber. A range area severely affected by downy mildew in a southern of representative isolates of H. parasitica were Australia 2015 canola disease survey. All H. parasitica chosen and mixed so as to avoid any possible isolates used were either single-spored (conidium) or pathotype-cultivar interactions that could otherwise hyphal-tipped from the original field samples collected. complicate outcomes. DNA of all isolates was extracted using the procedure of Cenis (1992) where mycelium from leaf samples was Temperature study – inoculation harvested directly, blotted dry with filter paper, washed with Tris-EDTA (TE) buffer, and pelleted by centrifu- Six seeds of each Brassica cultivar were sown in 6 × gation (5 min at 13,000 rpm), and the supernatant 6 cm seedling tray cells and thinned to four plants per decanted. After adding 300 μl of extraction buffer cell following seedling emergence. At growth stage 1.00 [200 mM Tris-HCl (pH 8.5), 250 mM NaCl, 25 mM (Sylvester-Bradley and Makepeace 1984) 7 days after EDTA, and 0.5% (wt/vol) sodium dodecyl sulphate], the seedling emergence, seedling trays were placed in mycelia were crushed manually. Then, 150 μlof3M 790 mm (L), 390 mm (W), 155 mm (H) clear plastic sodium acetate (pH 5.2) was added, and tubes main- boxes with lids then transferred into controlled environ- tained at 20 °C for 10 min. Subsequently, supernatant ment rooms maintained at 22/17 °C or 14/10 °C day/ was transferred to a fresh tube prior to adding an equal night). Two days later, seedlings were inoculated with a volume of isopropanol and centrifuged for 5 min at mixture, comprising 5 ml of 105 zoosporangia ml 1 13,000 rpm. After at least 5 min at ambient temperature suspension of a mixture of 10 H. parasitica isolates, (approximately 25 °C), the precipitated DNA was col- using a micropipette to place a 10 μl droplet of the lected by centrifugation for 15 min at 13,000 rpm. The mixed zoosporangial suspension on to each of two lobes
Page 81 of 119 706 Eur J Plant Pathol (2018) 151:703 711 of both cotyledons of each seedling. High humidity was index (%DI) based on the methods described by Mc- maintained for 24 h post-inoculation by hand-misting Kinney (1923), where: the walls and lids of the each plastic box with deionized water and then keeping the clear plastic lid on. Subse- %DI A½ðÞþa  0 ðÞþb  1 ðÞþc  2 ðÞþd  3 ðÞe  4 i quently, lids were removed and plants maintained by þ……:ðÞÂj x9 100Z=½ÞðÞa þ b þ c þ d þ ……j Â10 daily hand-watering for 6 days; then the plastic boxes were covered again for a further 24 h to maintain a and where a, b, c, d, e ……j are the number of plants second period of high humidity. with disease severity scores of 0, 1, 2, 3, 4, …..9, respectively. Plant age study – inoculation
Four seeds of each Brassica genotype were sown Experimental design and statistical analyses in 7 × 7 × 18 cm pots and grown at 18 °C day and 13 °C night in a controlled environment room with All experiments were fully repeated once. Both initial a day length of 12 h. At growth stage 1.00 and repeat experiments for each of the temperature and (Sylvester-Bradley and Makepeace 1984), 7 days plant age studies were arranged in a randomised com- after seedling emergence, pots were placed into plete block design, with four replicates of each treat- 770 mm (L), 570 mm (W), 475 mm (H) clear ment. For the temperature experiment, a third confirma- plastic boxes. Plant cotyledons were initially inoc- tion experiment was also undertaken, but only using ulated when seedlings were 7 days old and as three of the original six cultivars used in the first described above for the temperature study. Subse- experiment. The homogeneity of initial and repeat quently, at 8-day intervals for four repetitions, all experiments was assessed using DI% values for leaf adaxial surfaces were wet-brushed with the Btwo samples t-test for mean^.Asthet-test showed no same concentration of the mixed zoosporangial difference (P > 0.05) between the two original versus suspension (105 zoosporangia ml 1). High humidi- the repeat experiments for the temperature or the plant ty was maintained for 24 h post-inoculation for age studies, data for initial and repeat experiments was each inoculation and all other experimental condi- pooled in each case and reanalysed as a single data set. tions were as described above for the temperature Similarly, where three of the original cultivars used were study. repeated in a third experiment of the temperature study, there was no difference (P > 0.05) with either the orig- Disease assessment inal or first repeat experiments, but this latter data was not required or included in this paper. Disease severity The procedure of Williams (1985) was used to assess (%DI) data were analysed using two-way ANOVA disease severity on plants at 7 days post-inoculation with GenStat Release 14.2 (17th Edition, Lawes (dpi) for the temperature study and additionally at 15, Agricultural Trust, Rothamsted Research, UK). 23, 31 and 40 dpi for the plant age study, using a Fisher’s least significant differences (LSDs) were modified 0–9 scale, where: 0 = no symptoms or used to separate significant differences between sign of downy mildew disease; 1 = minute scattered cultivars. Correlation coefficients were computed necrotic flecks under the inoculum drop or inoculated for the relationship between disease index values plant surface area, no sporulation; 2 = larger necrotic versus plant age for each of the three cultivars used in flecks under the inoculum drop or inoculated plant the plant age study. surface area, no sporulation; 3 = very sparse sporulation, one to a few conidiophores, necrotic flecking but often with tissue necrosis evident; 5 = sparse spor- Results ulation, tissue necrosis; 7 = abundant sporulation, tissue necrosis and chlorosis may be present; and, Temperature study 9 = heavy sporulation, cotyledon or leaf tissue col- lapse. Subsequently, 0–9 disease severity scores for Across the six Brassica cultivars, there were significant each seedling tray cells were converted into a disease effects of genotype (P < 0.001), temperature (P <
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0.001), and genotype x temperature (P < 0.001) (Ta- Plant age study ble 1). On cotyledons of susceptible cultivars (B. napus Hyola 450 TT and Thunder TT), plants became symp- There were significant effects of cultivar (P < 0.001), tomatic at 22/17 °C by 48 h post inoculation (hpi) and plant age (P < 0.001), and a significant (P < 0.001) with abundant sporulation evident by 72 hpi. However, cultivar x plant age interaction in terms of % disease at 14/10 °C, no symptoms were observed until 5 dpi. indices. There were significant differences in mean At 22/17 °C, H. parasitica developed and sporu- % disease indices (%DI) between cultivars (LSD lated clearly on cotyledons by 3 dpi on susceptible at P < 0.05 = 1.68) across means for Dune (%DI cultivars B. napus Hyola 450 TT and Thunder TT. 20.2%), Hyola 450 TT (%DI 48.4%) and Surpass At 22/17 °C, all cotyledons of Thunder TT col- 402 (%DI 35.9%); and significant differences in mean lapsed by 7 dpi. In contrast, no sporulation at 14/ % disease indices between plant ages (LSD at P < 10 °C was observable until 7 dpi. DI%values of 0.05 = 1.94) across means for 15 (%DI 56.2), 23 (%DI cultivars at 22/17 °C were 4.5, 49.0, 51.4, 65.8, 86.3 32.4), 31 (%DI 28.5) and 40 (%DI 22.2) day-old-plants. and 96.0 for B. juncea Montara and B. napus DI% values were significantly greater on 15 day-old ATR-Hyden, Hyola 432, Atomic, Hyola 450 TT plants compared with 40 day-old plants across all culti- and Thunder TT, respectively. In contrast, DI% vars (Table 2). B. juncea Dune showed best resistance values of cultivars at 14/10 °C were 2.8, 30.4, with DI% values of 25.8, 24.6, 22.9 and 7.5, for 15, 23, 27.9, 31.1, 44.4 and 76.4, for B. juncea Montara 31 and 40 day-old-plants, respectively. B. napus Surpass and B. napus ATR-Hyden, Hyola 432, Atomic, 402 showed moderate susceptibility on cotyledons at Hyola 450 TT and Thunder TT, respectively. How- 15 day-old-plants but, in comparison, showed increased ever, the most disease resistant cultivar, B. juncea resistance on 23, 31 and 40 day-old-plants (i.e., %DI Montara, showed similar disease levels at both values of 59.0, 31.2, 27.1 and 26.2 for 15, 23, 31 and temperatures. 40 day-old-plants, respectively). B. napus Hyola 450 TT showed a highly susceptible response at the cotyledon stage on 15 day-old-plants, but moderate resistance on Table 1 The impact of different temperatures (22/17 °C, 14/ 10 °C day/night) on the epidemic of downy mildew of six different oilseed Brassica napus and B. juncea cultivars at 7 days post Table 2 Percent Disease Index (DI%) values on four different inoculation with a mixture of ten different isolates of plant ages (15, 23, 31 and 40 day old plants) of two Brassica Hyaloperonospora parasitica napus and one B. juncea cultivars following inoculation with a mixture of ten different isolates of Hyaloperonospora parasitica Brassica spp. Cultivar Temperature Disease (°C day/night) index (%) Brassica sp. Cultivar Plant age Disease (days old) index (%) B. juncea Montara 22/17 4.51 B. napus ATR Hyden 22/17 48.96 B. juncea Dune 15 25.83 B. napus Hyola 432 22/17 51.39 23 24.58 B. napus Atomic 22/17 65.8 31 22.92 B. napus Hyola 450 TT 22/17 86.28 40 7.50 B. napus Thunder TT 22/17 96.01 B. napus Surpass 402 15 58.96 B. juncea Montara 14/10 2.78 23 31.25 B. napus ATR Hyden 14/10 30.38 31 27.08 B. napus Hyola 432 14/10 27.95 40 26.25 B. napus Atomic 14/10 31.08 B. napus Hyola 450 TT 15 83.96 B. napus Hyola 450 TT 14/10 44.44 23 41.25 B. napus Thunder TT 14/10 76.39 31 35.42 40 32.92 Significant of cultivars P < 0.001; LSD at P<0.05=4.38 Significance of temperature P < 0.001; LSD at P < 0.05 =2.53 Significance of cultivars P<0.001;LSD at P < 0.05 =1.68 Significance of cultivars x temperature P < 0.001; LSD at Significance of plant age P < 0.001; LSD at P < 0.05 =1.94 P < 0.05 =6.20 Significance of cultivars x plant age P < 0.001; LSD at P<0.05=3.35
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Eur J Plant Pathol (2018) 151:703 711 709 with 40 dop across all cultivars. B. juncea Dune showed An important outcome of the current study was that greatest resistance, particularly on 40 dop; B. napus expression of resistance at the true leaf stages was often Surpass 402 showed moderate susceptibility on cotyle- greater than at the cotyledon stage. However, the current dons at 15 dop but greater resistance on older plants; study suggests that more rapid and less costly cotyledon while B. napus Hyola 450 TT showed very high screening should be a reliable means for preliminary susceptibility at cotyledon stage on 15 dop, but characterization of host genotype resistances in disease some resistance on 23 dop and increasing resistance on screening and plant breeding programs aiming to devel- 31 and 40 dop. The relative patterns of cultivar op more resistant cultivars to H. parasitica. Adult plant responses to H. parasitica at different plant ages were resistance can then be confirmed in the field on just a generally similar, despite the clear differences between smaller number of highly valuable genotypes. cotyledon and leaf responses in absolute terms with Cultivars need resistance that is effective under both more severe disease on younger plants. Association current and future climate conditions. Studies testing between expression of resistance to H. parasitica at effects of climate variables using cotyledons of oilseed cotyledon stage with that at the adult stage has been brassicas under controlled conditions offer avenues to reported by Jensen et al. (1999) on cauliflower (Brassica investigate and highlight impacts of future climate sce- oleracea var. botrytis), by Wang et al. (2000) on inbred narios on downy mildew epidemics and how best to broccoli, and by Zhang et al. (2012)inB. rapa (Chinese manage such. H. parasitica has been present in Austra- cabbage). Similarly, for the white leaf spot pathogen lian canola and mustard crops for decades (Barbetti and Pseudocercosporella capsellae, seedling cotyledons Khangura 2000). However, the reasons for its increasing across B. napus cultivars in general also provide a reliable severity and threat to commercial canola crops in some estimation of expression of field susceptibilities/resistance regions of Australia, particularly since 1999, have not on both seedlings and adult plants (Gunasinghe et al. been evident until this current study. It appears that 2014). In contrast, some other studies suggest resistance increasing incidence, severity and threat from downy to H. parasitica in cotyledons can be under a different mildew could be linked to increasing temperatures, par- genetic control than resistance in true leaves (Monteiro ticularly at the seedling stage when plants are most et al. 2005), but perhaps this relates to where a single susceptible. Further, it is clear from crop surveys in specific resistance gene has been identified (Satou and 2015 and 2016 that most severe downy mildew epi- Fukumoto 1996; Silué et al. 1996) that may express demics on canola occur in regions such as southwest differentially at different plant growth stages. Likewise, Western Australia when crops are subject to warmer Dickson and Petzoldt (1993) found that resistance to temperatures and more variable rainfall in the autumn- H. parasitica at the cotyledon stage of broccoli plants early winter period (Barbetti et al. 2012). In addition, seemed independent of adult-plant resistance when the development of downy mildew epidemics on Brassica plant has more than eight leaves. Further, Coelho et al. ceae is not only influenced by temperature and plant (1998) reported that resistance in B. oleracea cotyledons age, but also by humidity fluctuations that occur in had no association with that in adult plants. Likewise, for conjunction with temperature fluctuations, as highlight- some other Brassica diseases, cotyledon resistance may or ed on B. oleracea by Chu (1935), Le Beau (1945), may not accurately reflect adult plant resistance. For Felton and Walker (1946), Hartmann et al. (1983)and example, cotyledon resistance reflects adult plant resis- Achar (1998). A similar effect of temperature and hu- tance for specific major gene resistance against the black- midity on H. parasitica epidemics also has been leg stem canker pathogen Leptosphaeria maculans,but highlighted on radish (Bonnet and Blancard 1987; not necessarily for quantitative resistance against Kofoet and Fink 2007). L. maculans (Sivasithamparam et al. 2005;Neiketal. In conclusion, the findings of the current study ex- 2017). Likewise, for S. sclerotiorum, while there may be plain the development of the most severe downy mildew (Garg et al. 2008)ormaynotbe(Geetal.2008; Uloth epidemics in Australia on seedlings of susceptible cul- et al. 2014) a consistent expression of resistance on tivars early in the growing season when warmer tem- cotyledons versus more mature plants, resistance peratures coincide with presence of the more susceptible has been largely genotype-dependent, especially in seedlings rather than with the combination of less sus- relation to pathotype-dependent rather than pathotype- ceptible older plants and cooler temperatures as the independent resistances (Barbetti et al. 2014). growing season progresses during the winter months.
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Likewise, results of the current study indicate that the Cenis, J. L. (1992). Rapid extraction of fungal DNA for PCR ongoing increasing temperatures associated with cli- amplification. Nucleic Acids Research, 20, 2380. Channon, A. G. (1981). Downy mildew of Brassicas.InD.M. mate change mean that downy mildew epidemics in Spencer (Ed.), The downy mildews (pp. 321 339). London: Australia are likely to become even more widespread Academic Press. and severe in the years to come. Chu, H. T. (1935). Notes on the penetration phenomena and haustorium formation in Peronospora brassicae GÄUM. Japanese Journal of Phytopathology, 5,150 157. Acknowledgements The first author gratefully acknowledges a Coelho, P. S., & Monteiro, A. A. (2003). Inheritance of downy Scholarship from the University of Kufa in Iraq. The authors mildew resistance in mature broccoli plants. Euphytica, 131, grateful for the partial funding of both the Grains Research and 65 69. B Development Corporation (GRDC UWA 170 project Emerging Coelho, P., Bahcevandziev, K., Valerio, L., Monteiro, A., Leckie, ^ foliar diseases of canola ) and the School of Agriculture and D., Astley, D., Crute, I. R., & Boukema, I. (1998). The Environment, University of Western Australia. We are also grate relationship between cotyledon and adult plant resistance to ful for the generosity of seed companies in supplying seed of test downy mildew (Peronospora parasitica)inBrassica varieties and the exceptional technical support from Robert Creasy oleracea. Acta Horticulturae, 459,335 342. and Bill Piasini in the UWA Plant Growth Facilities. Coelho, P. S., Valério, L., & Monteiro, A. A. (2009). Leaf position, leaf age and plant age affect the expression of downy mildew Compliance with ethical standards resistance in Brassica oleracea. European Journal of Plant Pathology, 125,179 188. Ethical statement This research did not involve any animal and/ Coelho, P. S , Vicente, J. G., Monteiro, A. A., & Holub, E. B. or human participants. (2012). Pathotypic diversity of Hyaloperonospora brassicae collected from Brassica oleracea. European Journal of Plant Conflict of interest The authors declare that they have no con Pathology, 134,763 771. flict of interests. Dickson, M. H., & Petzoldt, R. (1993). Plant age and isolate source affect expression of downy mildew resistance in broccoli. Hortscience, 28,730 731. Elliott, V. L., Norton, R. M., Khangura, R. K., Salisbury, P. A , & Marcroft, S. J. (2015). Incidence and severity of blackleg References caused by Leptosphaeria spp. in juncea canola (Brassica juncea L.) in Australia. Australas. Australasian Plant Pathology, 44,149 159. Achar, P. (1998). Effects of temperature on germination of Felton, M. W., & Walker, J. C. (1946). Environmental factors Peronospora parasitica conidia and infection of affecting downy mildew of cabbage. Journal of Brassica oleracea. Journal of Phytopathology, 146, Agricultural Research, 72,69 81. 137 141. Garg, H., Sivasithamparam, K., Banga, S. S., & Barbetti, M. J. Bachofer, M. (2004). Molekularbiologische Populationsstudien (2008). Cotyledon assay as a rapid and reliable method of an Plasmopara halstedii, dem Falschen Mehltau der screening for resistance against Sclerotinia sclerotiorum in Sonnenblume (Doctoral dissertation, M. Bachofer). Brassica napus genotypes. Australasian Plant Pathology, 37, Stuttgart: University of Hohenheim. 106 111. Barbetti, M. J., & Khangura, R. (2000). Fungal diseases of canola Ge,X.T.,Li,H.,Han,S.,Sivasithamparam,K.,& in Western Australia. Agriculture Western Australia Bulletin, Barbetti, M. J. (2008). Evaluation of Australian 4406,15. Brassica napus genotypes for resistance to the downy Barbetti, M. J., Banga, S. S., & Salisbury, P. A. (2012). Challenges mildew pathogen, Hyaloperonospora parasitica. Crop for crop production and management from pathogen biodi and Pasture Science, 59,10301034. versity and diseases under current and future climate scenar Göker, M., Riethmüller, A., Voglmayr, H., Weiss, M., & ios case study with oilseed Brassicas. Field Crops Research, Oberwinkler, F. (2004). Phylogeny of Hyaloperonospora 127,225 240. based on nuclear ribosomal internal transcribed spacer se Barbetti, M. J., Banga, S. K., Fu, T. D., Li, Y.C., Singh, D., Liu, S. quences. Mycological Progress, 3,83 94. Y., Ge, X. T., & Banga, S. S. (2014). Comparative genotype Göker, M., Voglmayr, H., Blázquez, G. G., & Oberwinkler, F. reactions to Sclerotinia sclerotiorum within breeding popu (2009). Species delimitation in downy mildews: The case of lations of Brassica napus and B. juncea from India and Hyaloperonospora in the light of nuclear ribosomal ITS and China. Euphytica, 197,47 59. LSU sequences. Mycological Research, 113,308 325. Bonnet, A., & Blancard, D. (1987). Resistance of radish Gunasinghe, N., You, M. P., Banga, S. S., & Barbetti, M. J. (2014). (Raphanus sativus L.) to downy mildew Peronospora High level resistance to Pseudocercosporella capsellae offers parasitica. Cruziferae Newsletter, 12,98 99. new opportunities to deploy host resistance to effectively Bureau of Meteorology. (2014). State of the climate, 2014. Source manage white leaf spot disease across major cruciferous Available at: https://www.climatechangeinaustralia.gov. crops. European Journal of Plant Pathology, 138,873 890. au/en/climate campus/australian climate change/australian Gunasinghe, N., You, M. P., Cawthray, G. R., & Barbetti, M. J. trends/. (2016). Cercosporin from Pseudocercosporella capsellae
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and its critical role in white leaf spot development. Plant Neik, T. X., Barbetti, M. J , & Batley, J. (2017). Current status and Disease, 100, 1521 1531. challenges in identifying disease resistance genes in Brassica Hartmann, H., Sutton, J. C., & Procter, R. (1983). Effects of napus. Frontiers in Plant Science, 8, 1788. https://doi. atmospheric water potentials, free water, and temperature org/10.3389/fpls.2017.01788. on production and germination of sporangia in Paul, V. H , Klodt Bussmann, E., Dapprich, P. D., Capelli, C., Peronospora parasitica. Canadian Journal of Plant Tewari, J. P., Kohr, K., Thomas, J., & Dupprich, P. D. (1998). Pathology, 5,70 74. Results on preservation, epidemiology, and aggressiveness of Howlett, B., Ballinger, D., & Barbetti, M. J. (1999). Diseases. In Peronospora parasitica and results with regard to the disease Salisbury, P. A., Potter, T.D , McDonald, G., Green, A. G. resistance of the pathogen on Brassica napus. Bulletin (Eds.), Canola in Australia:The first thirty years (pp. 47 52). OILB/SROP, 21,49 56. Canberra: Organising committee of 10th International Sangeetha, C. G., & Siddaramaiah, A. L. (2007). Epidemiological Rapeseed Congress. studies of white rust, downy mildew and Alternaria blight of Jensen, B. D., Hockenhull, J., & Munk, L. (1999). Seedling and Indian mustard (Brassica juncea (Linn.) Czern. and Coss.) adult plant resistance to downy mildew (Peronospora African Journal of Agricultural Research, 2,305 308. parasitica) in cauliflower (Brassica oleracea convar. botrytis Satou, M., & Fukumoto, F. (1996). The host range of downy var. botrytis). Plant Pathology, 48,604 612. mildew, Peronospora parasitica, from Brassica oleracea, Jones, R. A. C., & Barbetti, M. J. (2012). Influence of climate cabbage and broccoli crops. Japanese Journal of change on plant disease infections and epidemics caused by Phytopathology, 62,393 396. viruses and bacteria. Plant Sciences Reviews, 22,131 Si, P., Mailer, R. J., Galwey, N., & Turner, D. W. (2003). Influence Online at http://www.cabi.org/cabreviews. of genotype and environment on oil and protein concentra Kofoet, A., & Fink, M. (2007). Development of Peronospora tions of canola (Brassica napus L.) grown across southern parasitica epidemics on radish as modelled by the effects of Australia. Crop and Pasture Science, 54,397 407. water vapour saturation deficit and temperature. European Silué, D., Nashaat, N. I., & Tirilly, Y. (1996). Differential re Journal of Plant Pathology, 117,369 381. sponses of Brassica oleracea and B. rapa accessions to seven Laemmlen, F. F., & Mayberry, K. S. (1984). Broccoli resistance to isolates of Peronospora parasitica at the cotyledon stage. downy mildew. California Agriculture, 38,17. Plant Disease, 80,142 144. Le Beau, F. J. (1945). Systemic invasion of cabbage seedlings by Sivasithamparam, K., Barbetti, M. J., & Li, H. (2005). 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Uloth, M., You, M. P., Finnegan, P. M., Banga, S. S., Yi, H., & Mohammed, A. E., You, M. P., & Barbetti, M. J. (2017). Barbetti, M. J. (2014). Seedling resistance to Sclerotinia New resistances offer opportunnity for effective man sclerotiorum as expressed across diverse cruciferous species. agement of the downy mildew (Hyloperonospora Plant Disease, 98,184 190. parasitica) threat to canola. Crop and Pasture Uloth, M. B., You, M. P., Cawthray, G., & Barbetti, M. J. (2015). Science, 68,234242. Temperature adaptation in isolates of Sclerotinia Monteiro, A. A., Coelho, P. S , Bahcevandziev, K., & Valério, L. sclerotiorum affects their ability to infect Brassica carinata. (2005). Inheritance of downy mildew resistance at cotyledon Plant Pathology, 64, 1140 1148. and adult plant stages in ‘Couve Algarvia’ (Brassica Wang, M , Farnham, M. W., & Thomas, C. E. (2000). Phenotypic oleracea var. tronchuda). Euphytica, 141,85 92. variation for downy mildew resistance among inbred broc Nashaat, N. I., Heran, A., Awasthi, R. P., & Kolte, S. J. (2004). coli. Hortscience, 35,925 929. Differential response and genes for resistance to Peronospora Williams,P.H.(1985).‘Downy mildew.’ Crucifer genetics coop parasitica (downy mildew) in Brassica juncea (mustard). erative (CRGC) resource book. Madison: Department of Plant Breeding, 123,512 515. Plant Pathology, University of Wisconsin. Natti, J. J., Hervey, G. E. R., & Sayre, C. B. (1956). Zhang, S., Yu, S., Zhang, F., Si, L., Yu, Y., Zhao, X., Zhang, D., & Factors contributing to the increase of downy mildew Wang, W. (2012). Inheritance of downy mildew resistance at of broccoli in New York state and its control with different developmental stages in Chinese cabbage via the fungicides and Agrimycin. Plant Disease Reporter., leaf disk test. Horticulture, Environment, and Biotechnology, 40,118124. 53,397 403.
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Pathotypes and phylogenetic variation determine downy mildew epidemics in Brassica spp. in Australia
This chapter focuses on the selection of suitable Brassica hosts to characterise downy mildew isolates into pathotypes and molecular studies to determine the genetic variation among the downy mildew population in Australia.
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Downy mildew pathotypes and variation 1515
Expressions of field susceptibilities/resistances to downy characterization of pathotypes of H. brassicae worldwide mildew have been reported for the seedling stage of and for monitoring and understanding changes in H. bras B. napus (Ge et al., 2008; Mohammed et al., 2017) and sicae populations over time and between locations. on more mature leaves of B. oleracea (Coelho et al., 1998, 2009; Jensen et al., 1999; Wang et al., 2001; Materials and methods Carlsson et al., 2004). In B. oleracea, Dickson & Petzoldt (1996) reported the presence of quantitative Brassicaceae genotypes resistance, Hoser Krauze et al. (1991) reported a single recessive gene, and Moss et al. (1988), Hoser Krauze A set of 28 Brassicaceae genotypes was used in this study, com et al. (1995), Carvalho & Monteiro (1996) and Vicente prising 13 B. napus, two B. juncea, five B. oleracea, two et al. (2012) all reported one or more dominant resis E. vesicaria and one each of B. nigra, B. carinata, B. rapa, tance genes. In B. juncea, Nashaat et al. (2004) reported Crambe abyssinica, R. sativus and R. raphanistrum (Table 1). a single dominant resistance gene and Lucas et al. (1988) and Nashaat et al. (1997) also reported a single domi Hyaloperonospora brassicae isolates nant resistance gene in B. napus. Natti et al. (1967) were the first to report separate dis Nineteen isolates of H. brassicae were collected in 2006 2008 from B. napus, B. oleracea and R. raphanistrum from geograph tinct physiological races (1 and 2) in broccoli (B. oler ical locations representative of key oilseed and vegetable bras acea var. italica) and later, Coelho et al. (2012) sica growing regions in Western Australia (Table 2). In addition, identified six pathotypes of H. brassicae from B. oler a further 11 isolates of H. brassicae were collected in 2015 acea. Sherriff & Lucas (1990) used 33 H. brassicae iso 2016 from canola leaves as part of a southern Australia 2015 lates from four different Brassica hosts (B. napus, 2016 foliar disease survey. These consisted of nine isolates from B. juncea, B. oleracea and B. campestris) and locations Western Australia (WA5c, WA7c, WA11c, WA12c, WA13c, to infect a set of Brassica genotypes (including B. napus, B. juncea, B. oleracea, B. carinata, B. nigra, B. cam Table 1 Brassica and other cruciferous taxa used for current study pestris and Raphanobrassica) and found variation in host and their source. range across agriculturally and horticulturally important Brassica crops. However, they failed to delineate patho Seed Species Cultivar/line Crop source types. Nashaat & Rawlinson (1994) classified 101 B. na pus subsp. oleifera genotypes into three groups, ‘A’, ‘B’ Brassica carinata ATC 94011 Osiebia mustard DAFWAa and ‘C’, using two H. brassicae isolates, R1 and P003. B. juncea Muscan 963 Oilseed rape DAFWA Group A was resistant to both isolates, group B was B. juncea Dune Oilseed rape DAFWA B. napus Cobbler Oilseed rape, spring DAFWA resistant to R1 but susceptible to P003 whereas group C B. napus AG Comet Oilseed rape, spring DAFWA was susceptible to both isolates. Subsequently, Nashaat B. napus Tranby Oilseed rape, spring DAFWA et al. (1997) identified a fourth differential response to B. napus ATR Banju Oilseed rape, spring DAFWA H. brassicae in B. napus that was resistant to P003 but B. napus ATR Stubby Oilseed rape, spring DAFWA partially resistant to R1, but they did not nominate a B. napus CB Trilogy Oilseed rape, spring DAFWA designation for this last grouping. Knowledge of patho B. napus Thunder TT Oilseed rape, spring DAFWA type structure of H. brassicae remains, at best, sparse. B. napus Pioneer 45Y77 Oilseed rape, spring DAFWA Studies to date of the phenotypic expression of host B. napus Capricorn C Oilseed rape, winter RRUKb resistances and genetics of resistance in brassicas to B. napus Cresor 770 B Oilseed rape, spring RRUK H. brassicae have been compromised by a lack of a com B. napus Cresor 771 B Oilseed rape, spring RRUK B. napus Komet 741 A Oilseed rape, spring RRUK mon set of host differentials to characterize isolates. B. napus Komet 744 A Oilseed rape, spring RRUK Successful and efficient breeding for resistance to B. nigra Introduce Black mustard DAFWA H. brassicae in brassicas is dependent upon a reliable P.23845 and comparable system to delineate pathotypes. B. rapa Xi Shui Bai Chinese cabbage DAFWA The present study was conducted using the differential B. oleracea Tighthead Brussels sprout Yates set of Nashaat et al. (2004) and a range of other impor B. oleracea Summer Green Broccoli Yates tant broad acre and/or vegetable and/or weedy Brassi B. oleracea Sweet Eureka Cabbage Yates caceae (including B. carinata, B. juncea, B. napus, B. oleracea Kale Chinese broccoli Yates B. nigra, B. oleracea and B. rapa, Crambe abyssinica, B. oleracea Phenomenal Cauliflower Yates Raphanus sativus, R. raphanistrum and Eruca vesicaria). Early Crambe CMB 94054 Oilseed DAFWA One objective was to identify and select suitable Brassica abyssinica spp. differentials to enable characterization of H. brassi Eruca vesicaria Yellow seed Leaf vegetable DAFWA cae pathotypes, and to use the octal code system of Good E. vesicaria Brown seed Leaf vegetable DAFWA win et al. (1990) to define these pathotypes. In addition, Raphanus sativus Colonel Radish Yates analyses were undertaken to define phylogenetic relation R. raphanistrum Wild radish Weed DAFWA ships across a more extensive set of 2006 2008 and recent isolates of H. brassicae from Australia. This study pro aDepartment of Agriculture and Food Western Australia. vides both a basis for standardizing phenotypic bRothamsted Research, UK.
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Table 2 Isolates of Hyaloperonospora brassicae used in this study, their original host species, agrogeographical origin and year of collection.
Isolatea Australian state Original host Latitude Longitude Year of collection
A UWA2h Western Australia Brassica napus canola 31.2661 116.4692 2006 UWA8h Western Australia B. napus canola 31.0578 116.8583 2006 UWA11h Western Australia B. napus canola 31.7175 116.5306 2006 UWA19h Western Australia B. napus canola 31.7233 117.9325 2006 UWA25h Western Australia B. napus canola 33.7992 117.5417 2006 UWA33h Western Australia B. napus canola 31.2256 116.6417 2007 UWA40h Western Australia B. napus canola 31.0211 116.8644 2007 UWA51h Western Australia B. napus canola 33.5722 121.8592 2007 UWA58h Western Australia B. napus canola 33.5403 121.5958 2007 UWA59h Western Australia B. napus canola 33.7581 121.5961 2007 UWA60h Western Australia B. napus canola 33.5667 121.5958 2007 UWA61h Western Australia B. napus canola 33.6589 121.5886 2007 UWA9h Western Australia Raphanus raphanistrum 31.0578 116.8583 2006 UWA34h Western Australia R. raphanistrum 31.0661 116.5883 2007 UWA65h Western Australia B. oleracea cabbage 31.6972 115.8250 2008 UWA67h Western Australia B. oleracea broccoli 31.6972 115.8250 2008 UWA73h Western Australia B. oleracea cauliflower 34.2828 116.2717 2008 UWA79h Western Australia B. oleracea broccoli 34.4400 116.3781 2008 UWA80h Western Australia B. oleracea broccoli 34.4400 116.3781 2008 WA5c Western Australia B. napus canola 31.3417 116.6381 2015 WA7c Western Australia B. napus canola 31.1989 116.9769 2015 WA11c Western Australia B. napus canola 31.2347 117.5383 2015 WA12c Western Australia B. napus canola 31.3325 117.3789 2015 WA13c Western Australia B. napus canola 31.5494 117.5969 2015 WA22c Western Australia B. napus canola 32.0989 117.4233 2015 WA29c Western Australia B. napus canola 32.6989 117.8739 2015 WA30c Western Australia B. napus canola 32.6994 117.6717 2015 WA49c Western Australia B. napus canola 34.5050 117.4381 2015 SA78c South Australia B. napus canola 33.5092 138.3633 2015 VC36c Victoria B. napus canola 37.3406 141.8775 2016 B UWA4h Western Australia B. napus canola 31.3072 116.4872 2006 UWA24h Western Australia B. napus canola 33.7453 117.7083 2006 UWA31h Western Australia B. napus canola 31.2672 116.4756 2007 UWA44h Western Australia B. napus canola 33.4592 117.1572 2007 UWA53h Western Australia B. napus canola 33.6156 121.7525 2007 UWA56h Western Australia B. napus canola 33.5222 121.5950 2007 UWA57h Western Australia B. napus canola 33.6758 121.5956 2007 UWA68h Western Australia B. oleracea cabbage 34.4178 116.4006 2008 UWA74h Western Australia B. oleracea broccoli 34.4494 116.4958 2008 UWA77h Western Australia B. oleracea cauliflower 34.6222 116.3167 2008 UWA78h Western Australia B. oleracea cabbage 34.5086 116.2614 2008 UWA83h Western Australia B. oleracea cauliflower 32.1297 116.1939 2008 UWA84h Western Australia B. oleracea cabbage 32.4331 116.5964 2008 WA1c Western Australia B. napus canola 31.4486 116.3344 2016 WA2c Western Australia B. napus canola 31.3981 116.3306 2016 WA4c Western Australia B. napus canol 31.3078 116.3789 2016 WA6c Western Australia R. raphanistrum 31.3161 116.6092 2016 WA8c Western Australia B. napus canola 31.1461 116.6583 2016 WA9c Western Australia B. napus canola 31.1297 116.7944 2016 WA17c Western Australia B. napus canola 31.6800 117.3244 2016 WA23c Western Australia B. napus canola 32.4531 117.7978 2016 WA24c Western Australia B. napus canola 32.4439 117.8331 2016 WA25c Western Australia B. napus canola 32.6194 117.8942 2015 WA28c Western Australia B. napus canola 32.1639 117.3683 2015 aA, isolates used for pathogenicity test; B, isolates used for molecular markers; UWA + number + h, 2006 2008 isolates; WA, SA, VC + number + c, 2015 2016 isolates.
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WA22c, WA29c, WA30c, WA49c) and a single isolate each 1 min and extension at 72 °C for 2 min; and a final extension from South Australia (SA78c) and Victoria (VC36c). All step at 72 °C for 7 min and then held at 4 °C. A final PCR was H. brassicae isolates had been obtained from the original field performed using the selected best annealing temperature of samples by removing hyphal tips using fine tweezers and a dis 53 °C. PCR products were subjected to agarose gel elec section microscope and then transferring them to cotyledons of trophoresis at 50 V for 120 min on a 1% (w/v) agarose gel con 7 day old seedlings of highly susceptible B. napus cultivars taining 0.1% GelRed (Biotium Inc.) and then visualized under Thunder TT and Tranby. UV light. Aliquots of PCR products of 830 bp (30 lL of each) Each H. brassicae isolate was maintained in isolation on were sent to Macrogen Inc. (Korea) for sequencing. cotyledons of 7 day old seedlings of susceptible B. napus culti vars Thunder TT and Tranby (Ge et al., 2008) in separate sealed plastic 250 mL containers. These were kept in a con DNA sequence manipulation and analysis trolled environment room with a 12 h photoperiod, light inten sity 180 lmol m 2 s 1 at 13 °C night and 18 °C day. Isolates DNA sequence manipulation and analysis for both directions were maintained by weekly inoculating fresh cotyledons of seed were conducted using GENEIOUS v. 9.1.7 (Biomatters Ltd). lings of susceptible cultivars by taking previous infected seedling Sequences of all isolates were confirmed as H. brassicae using > cotyledons, shaking in sterile distilled water to dislodge zoospor the BLAST search in GenBank ( 99% similarity). GenBank angia, and then adding a 20 lL droplet of the zoospore suspen H. brassicae accessions AY531407 (Germany), DQ447119 (Aus sion to each cotyledon. Equal numbers of cotyledons that tralia), JF9756614 (China), KX231682 (USA) and LC050224 supported abundant sporulation by each of the different patho (Japan) were retrieved from the database and compared with 20 gen isolates were collected separately for each H. brassicae iso sequences of the 2006 2008 isolates and 88 sequences from the late and placed in 50 mL distilled water in a 100 mL flask. 2015 2016 isolates. All sequences were aligned using GENEIOUS Flasks were shaken to dislodge the zoosporangia, the resulting alignment with cost matrix of 70% similarity (IUB) (5.0/ 4.5), suspension was filtered through a single layer of cheesecloth to gap open penalty: 13, gap extension penalty: 3 and global align remove any cotyledon contamination, and the concentration ment with free end gaps as the alignment type. A neighbour was adjusted to 105 zoospores mL 1 using a haemocytometer joining tree was constructed using the function GENEIOUS TREE counting chamber (Superior). BUILDER and adopting the Tamura Nei genetic distance model and the bootstrap resampling method (random seed 900 470, number of replications 1500, and support threshold >70%). The sequence of the ribosomal DNA (rDNA) gene of Leptosphaeria DNA extraction and PCR conditions maculans AY422213 from GenBank was used as the out group. DNA was extracted using the procedure of Cenis (1992) for the Thirty five sequences representing the seven clades of the seven H. brassicae isolates collected in 2006 2008 and the 11 constructed tree (some isolates in the tree are identical) were further isolates (detailed above). In addition, to enable phyloge submitted to the GenBank database (accession numbers netic studies, DNA was similarly extracted from an additional MG757758 MG757792; Table S1) using the submission portal 13 isolates collected in 2006 2008 and 77 recent isolates. In (https//submit.ncbi.nlm.nih.gov/subs/genbank/). brief, mycelia from leaf samples were harvested directly from infested leaves using tweezers and a dissecting microscope, then blotted dry with filter paper. Mycelia were homogenized using Inoculation Precellys Evolution Homogenizer (bertin TECHNOLOGIES), Six seeds of each plant genotype were sown in 6 9 6 cm seed washed with Tris EDTA (TE) buffer, then pelleted by centrifug ling tray cells and thinned to four plants per cell following seed ing at 39 443 g for 10 min and discarding the supernatant. ling emergence. Genotypes were grown at 18 °C day and 13 °C l After adding 300 L of extraction buffer (200 mM Tris HCl night in a controlled environment room with a day length of (pH 8.5), 250 mM NaCl, 25 mM EDTA), 0.5% (w/v) sodium 12 h. Plants were watered daily with deionized water and l dodecyl sulphate (SDS) was added to the pellet. Then, 150 L allowed to drain to field capacity. At growth stage 1.00 of 3 M sodium acetate (pH 5.2) was added and the mixture was (Sylvester Bradley & Makepeace, 1984), 7 days after seedling ° maintained at 20 C for 10 min, followed by vortexing briefly emergence, seedling trays were placed in clear plastic boxes with and centrifuging at 39 443 g for 5 min. Subsequently, the super lids, measuring 790 mm (L), 390 mm (W), 155 mm (H). Subse natant was transferred to a fresh tube prior to adding an equal quently, seedlings were inoculated with a suspension of concen l volume of cold isopropanol (450 L), centrifuged at 39 443 g tration 105 sporangia mL 1 for each H. brassicae isolate using for 10 min and then left to stand at least 15 min at room tem the methodologies described earlier by Mohammed et al. perature. Precipitated DNA was collected by centrifugation at (2017). Briefly, a micropipette was used to place a 10 lL dro 39 443 g for 15 min. The pellet was air dried, then washed with plet of the sporangial suspension on to each of two lobes of l 70% ethanol (v/v ethanol:water) and resuspended in 50 LTE both cotyledons of each seedling. High humidity was maintained buffer. The quantity and quality of extracted DNA was deter for 24 h post inoculation by misting the walls and lids of each mined with a NanoDrop 1000 spectrophotometer (Thermo Sci plastic box with deionized water using a hand held spray and ° entific) and DNA was stored at 4 C. The DNA was subjected then keeping the clear plastic lid on. Subsequently, lids were to PCR using a master mix of a total volume of 50 lL that con 0 removed and plants maintained by daily hand watering; after tained 0.2 lM of each primer (primers ITS1 O (5 CGGAAGGA 0 0 6 days the plastic boxes were covered again for a further 24 h TCATTACCAC 3 ; Bachofer, 2004) and ITS4 H (5 TCCTCC to maintain a second period of high humidity. GCTTATTAATATGC 30;Goker et al., 2004), a modification of ITS4. ITS1 O was chosen due to its specificity that replaces the need for additional amplification of host ITS rDNA (Bachofer, Disease assessment 2004; Goker et al., 2009). PCR was undertaken as follows: ini tial denaturation 94 °C for 2 min; 35 cycles at 94 °C for 1 min, The procedure of Williams (1985) was used to assess disease with annealing gradient temperature set at range 50 60 °C for severity on plants at 7 days post inoculation (dpi) using a 0 9
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scale: 0 no symptoms or sign of downy mildew disease; undertaken to confirm the robustness of these H. brassicae iso 1 minute scattered necrotic flecks under the inoculum drop, late allocations into the different pathotype groupings (Clarke, no sporulation; 2 larger necrotic flecks under the inoculum 1993; implemented in PRIMER v. 6). Heat maps were produced drop, no sporulation; 3 very sparse sporulation, one to a few using CLUSTVIS analysis (Metsalu & Vilo, 2015) to confirm fur conidiophores, necrotic flecking but often with tissue necrosis ther that these were the six most suitable Brassica differentials evident; 5 sparse sporulation, tissue necrosis; 7 abundant (from B. oleracea, B. nigra and B. napus) following inoculation sporulation, tissue necrosis and chlorosis may be present; and with 30 different isolates of H. brassicae. The heat maps also 9 heavy sporulation, cotyledon collapse. provided visualization of the host pathotype interactions in terms of severity of disease expression. Pathotype characterization and nomenclature Octal nomenclature, as developed by Goodwin et al. (1990) to Results characterize Rhynchosporium secalis pathotypes and subse quently used by You et al. (2005) and Ge et al. (2012) to char Isolates collected in 2006 2008 acterize Phytophthora clandestina and Sclerotinia sclerotiorum pathotypes, respectively, was used to code H. brassicae isolates There was a significant effect of isolates (P ≤ 0.001) and according to their virulence on six differential Brassica geno host genotypes (P ≤ 0.001), as well as a significant inter types selected from the 28 host genotypes. The basis of classify action between isolates and genotypes (P ≤ 0.001) in ing Brassica genotype disease reactions across different relation to cotyledon infection (Table 3). H. brassicae isolates tested (expressed as different levels of dis While isolates of H. brassicae were most virulent on ease severity) was bimodal with an assumption of two possible modes, one ‘susceptible’ and the other ‘resistant’, providing that their species of origin, they did cause disease on Brassica there was clear demarcation between these two modes in accor hosts from which the isolates were not originally dance with Goodwin et al. (1990). The six Brassica genotypes obtained. For example, isolates from B. napus, except used as differentials were grouped into two sets according to UWA8h and UWA59h, caused severe disease on B. na their genetic inheritance and continuity with a previous study pus CB Trilogy, Thunder TT and Capricorn C, and iso (Nashaat & Rawlinson, 1994); thus, cultivars that shared resis lates from B. oleracea, except UWA79h, caused severe tance genes or had related resistance genes were grouped in the disease on B. oleracea broccoli, Chinese broccoli, cauli same triplet. Brassica oleracea Brussels sprout, B. nigra Intro flower, cabbage and Brussels sprout. In contrast, isolates duce P.23845 and B. napus Cresor 770 B were grouped as one UWA9h and UWA34h, isolated from R. raphanistrum at set or triplet; B. napus Tranby, CB Trilogy and Thunder TT different locations and in different years, were avirulent were grouped as a second set or triplet. Pathotype codes were based on arranging the differentials from right to left, those with on this species but virulent on most B. napus genotypes. the fewest resistance genes on the right and those with the most Furthermore, two isolates (UWA8h from B. napus, and on the left, and then scoring the virulence of the isolates and UWA79h from B. oleracea) were avirulent on all test their interaction with each differential in the set, where 0 indi genotypes except on B. napus Capricorn C. Overall, iso cates a resistant reaction and 1 indicates a susceptible reaction. lates UWA11h and UWA25h produced greatest sporula Octal digits were assigned to each triplet by converting from the tion across most B. napus and B. oleracea genotypes and binary code as follows: 000 0; 001 1; 010 2; 011 3; caused most severe disease on B. napus and B. oleracea 100 4; 101 5; 110 6; 111 7. (Table 3). Across the 28 test genotypes, symptoms ranged from Experimental design and statistical analysis no visible symptom, hypersensitive reaction in highly resistant hosts to systemic growth and abundant sporula Studies using 2006 2008 isolates were repeated once, and for tion of the pathogen in fully compatible host pathogen each isolate there were six replicates in a completely randomized interactions. Raphanus sativus, R. raphanistrum, C. design with four plants within each replicate. The mean disease abyssinica and E. vesicaria as well as B. carinata, score for the four plants (i.e. 16 separately inoculated cotyledon B. juncea and B. napus Pioneer 45Y77 were highly resis lobes) in each replicate was used as the score for each replicate. Studies using 2015 2016 isolates were also repeated once and tant to all 2006 2008 isolates tested. Raphanus sativus, used a similar completely randomized design, but with three B. napus (Cresor 771, Komet 741 and Komet 744 and replicates for each isolate. All other experimental details were Cresor 770), B. rapa and B. nigra were immune to all similar as for studies described above involving isolates collected isolates from B. oleracea and R. raphanistrum, but were in 2006 2008. The relationship between the initial and repeat susceptible to one or more B. napus isolates. Other experiments was assessed using a paired t test using GENSTAT B. napus genotypes, ATR Cobbler, AG Comet, Tranby, and the homogeneity of variances across the original and repeat ATR Banjo, ATR Stubby, CB Trilogy and Thunder TT, experiments was assessed using Bartlett’s test (Snedecor & while resistant to all B. oleracea isolates, were suscepti Cochran, 1989). In each case there were no significant differ ble to most isolates from B. napus and R. raphanistrum. ences between each pair of experiments (P > 0.05) using t test, B. napus Capricorn C was resistant to most B. oleracea and variances were similar using Bartlett’s test; therefore, the original data sets were combined and presented. Disease severity isolates but susceptible to both isolates from data were analysed using two way ANOVA with GENSTAT R. raphanistrum, 11 out of 12 from B. napus as well as release 18.1 (Lawes Agricultural Trust, Rothamsted Research). B. oleracea isolate UWA79h. Brassica oleracea cabbage, Fisher’s least significant differences (LSD) were used to separate Brussels sprout, broccoli, Chinese broccoli and cauli significant differences between genotypes. Cluster analyses) were flower were susceptible to all but one of the isolates from
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(2018) Table 3 Sever ty of downy m dew d sease on coty edons of 28 Brass caceae genotypes nocu ated w th 19 so ates of Hyaloperonospora brassicae co ected from Western Austra a n 2006 2007 and 2008 67
541527 1514 , UWA2ha UWA8h UWA9h UWA11h UWA19h UWA25h UWA33h UWA34h UWA40h UWA51h UWA58h UWA59h UWA60h UWA61h UWA65h UWA67h UWA73h UWA79h UWA80h Species Genotype BN BN RR BN BN BN BN RR BN BN BN BN BN BN Br Br Br Br Br Means
Raphanus sativus Colonel 0 00 0 00 0 00 1 20 0 40 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 13 0 00 0 00 0 00 0 09 R raphanistrum Wild radishb 0 06 0 06 0 00 0 93 0 53 0 00 0 46 0 00 0 00 0 30 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 00 0 12 Brassica ATC 94011 0 00 0 06 0 06 1 06 1 20 0 00 0 53 0 26 0 60 0 23 0 26 0 00 0 00 0 53 0 00 0 13 0 00 0 00 0 60 0 29 carinata Eruca vesicaria Yellow seedb 1 33 0 80 1 73 1 80 1 80 1 40 1 86 2 00 1 46 1 00 1 00 0 73 1 20 1 66 1 06 0 80 0 46 0 00 1 33 1 23 E vesicaria Brown seedb 0 86 0 80 1 20 1 60 1 13 0 46 1 06 1 33 1 06 0 96 1 00 0 00 0 86 2 00 0 66 0 80 0 33 0 00 1 20 0 91 Crambe CMB 94054 1 06 0 00 0 00 1 60 0 00 0 00 1 13 0 00 1 73 1 10 1 13 0 00 0 80 0 20 0 00 0 13 0 20 0 00 0 00 0 47
abyssinica variation and pathotypes mildew Downy B juncea Muscan 963 2 86 1 60 3 06 3 93 3 53 2 40 2 93 2 40 3 00 2 70 2 33 0 46 2 86 4 06 0 33 1 33 0 20 2 26 2 26 2 34 B juncea Dune 0 06 0 00 0 13 1 06 0 26 0 20 0 00 0 20 1 66 1 60 0 80 0 00 0 60 0 00 0 00 0 00 0 00 0 00 0 00 0 34 B nigra ntroduce 1 33 0 40 1 40 5 33 3 13 1 26 5 40 1 13 2 06 1 60 1 00 0 00 2 06 3 33 0 00 0 00 0 20 0 40 0 40 1 60 P 23845 B rapa Xi Shui Bai 2 66 1 46 2 66 5 40 3 60 2 86 5 86 2 93 3 06 2 83 4 73 2 00 4 93 3 60 0 46 0 46 0 00 0 60 1 66 2 72
Page 94 of119 B napus Cresor-771 B 0 80 0 26 2 20 6 40 3 86 1 86 2 00 0 20 2 00 1 03 2 00 0 00 1 73 1 73 0 00 0 40 0 20 0 20 0 86 1 46 B napus Cresor-770 B 0 46 0 53 4 20 7 53 3 86 2 13 2 20 1 06 2 26 1 10 2 00 0 00 1 80 1 93 0 06 0 86 0 13 0 00 1 00 1 74 B napus Komet-741 A 4 46 4 40 1 26 4 13 4 93 2 20 3 46 3 53 2 13 2 03 2 40 1 13 2 20 3 93 0 20 0 60 0 00 0 00 0 86 2 31 B napus Komet-744 A 6 06 4 80 6 80 7 00 6 06 6 80 7 00 6 60 6 93 6 33 7 06 2 46 7 00 6 86 0 13 0 26 0 00 4 93 0 93 4 95 B napus Capricorn C 2 66 2 33 2 40 4 73 6 60 2 66 3 46 4 20 1 73 2 63 2 46 1 33 2 66 3 60 0 00 0 40 0 06 0 66 1 13 2 40 B napus AG Comet 4 26 2 40 6 20 7 60 5 33 6 13 6 66 4 06 5 73 4 90 7 20 4 13 6 93 6 53 0 46 0 33 0 20 3 26 0 57 4 36 B napus Tranby 5 00 1 13 5 26 8 00 6 66 7 20 5 80 5 93 7 20 6 30 8 13 1 53 5 66 6 86 0 93 1 40 0 33 1 40 1 00 4 51 B napus CB Trilogy 4 33 0 73 7 20 7 93 6 66 7 00 6 73 4 46 5 73 5 96 6 80 4 93 7 20 7 20 0 73 0 66 0 80 3 33 1 33 4 72 B napus ATR-Banjo 6 66 3 86 6 86 8 40 8 13 7 06 7 53 5 33 6 93 7 23 7 86 4 13 6 73 7 53 2 13 2 80 1 46 4 13 4 73 5 76 B napus ATR-Stubby 4 26 2 53 5 73 7 86 6 53 6 00 5 80 4 53 6 53 6 46 6 93 0 73 6 66 6 06 0 80 1 60 0 53 2 46 2 80 4 46 B napus ATR-Cobbler 6 00 1 33 5 86 8 13 6 93 6 66 7 00 4 40 2 73 3 66 5 60 1 80 5 60 5 06 0 33 0 66 0 06 1 93 1 33 3 95 B napus Pioneer 45Y77 2 00 0 26 2 60 2 66 2 78 1 80 4 06 2 60 1 66 2 26 2 73 0 20 3 33 3 33 1 00 0 60 0 46 0 00 0 66 1 84 B napus Thunder TT 7 00 3 73 8 00 8 60 7 93 8 20 8 73 4 93 9 00 7 86 8 80 5 60 8 13 8 06 0 66 1 26 0 46 3 73 1 33 5 89 B oleracea Brussels 5 20 3 06 4 86 6 60 4 86 4 46 6 06 5 00 6 00 6 20 6 00 0 00 6 06 5 66 7 13 6 73 6 66 2 93 6 20 5 24 sproutb B oleracea Cabbageb 4 93 3 46 5 33 3 26 6 00 5 13 1 60 6 13 2 00 4 23 2 40 3 06 1 86 5 60 7 06 6 93 7 06 2 20 7 00 4 48 B oleracea Broccolib 5 33 3 33 6 20 8 13 6 00 5 93 7 06 5 53 7 00 7 23 6 93 5 33 7 13 6 00 7 00 7 13 7 26 3 46 8 00 6 31 B oleracea Chinese 3 33 1 66 5 80 6 53 5 93 4 93 6 13 6 00 6 40 4 80 5 66 3 93 3 73 5 33 7 00 6 86 6 73 2 60 7 40 5 30 broccolib B oleracea Cauliflowerb 4 00 1 40 6 33 6 73 4 93 4 13 6 46 4 26 6 00 5 60 6 20 4 33 5 53 5 60 6 80 6 53 6 86 2 60 7 20 5 34 Means 3 11 1 66 3 69 5 15 4 27 3 53 4 18 3 18 3 66 3 50 3 91 1 71 3 69 4 01 1 60 1 78 1 45 1 54 2 20 3 04
D sease sever ty assessed on a 0–9 sca e where 0 = no d sease 9 = heavy sporu at on coty edon co apsed S gn ficance of cu t vars P < 0 001 LSD at P < 0 05 = 0 188 s gn ficance of so ates P < 0 001 LSD at P < 0 05 = 0 154 s gn ficance of cu t vars 9 so ates P < 0 001 LSD at P < 0 05 = 0 830 aUWA + number + h = 2006–2008 so ates BN Brassica napus Br B oleracea RR Raphanus raphanistrum bCommon name 1519 1520 A. E. Mohammed et al.
B. oleracea or R. raphanistrum, but resistant to just a There was a significant interaction between H. brassicae few isolates from B. napus and one from B. oleracea. isolates and disease severity on six genotypes selected as The five B. napus genotypes previously used as differ differentials. Each of the selected differentials showed entials in the UK (Komet 741, Komet 744, Cresor 770, clear bimodal distribution (Fig. 1). Analyses of histograms Cresor 771 and Capricorn C) behaved differently against based on disease severity scores of 19 2006 2008 isolates the different isolates. For example, Komet 741 expressed of H. brassicae on these differential hosts identified clear resistance to all isolates; Cresor 770 and Cresor 771 resistance versus susceptibility separation points for each were susceptible only to isolate UWA11h; Capricorn C differential as follows: disease severity rating 5 for B. na was susceptible only to isolate UWA19h; but Komet pus accession Cresor 770 B; 4 for Tranby; 4.5 for B. nigra 744 was susceptible to many isolates including UW Introduce P.23845; 6 for B. napus Thunder TT; 3 for CB A2h, UWA9h, UWA11h, UWA19h, UWA25h, UWA33h, Trilogy and 2.5 for B. oleracea Brussels sprout (Fig. 1). UWA34h, UWA40h, UWA52h, UWA58h, UWA60h and Disease severity less than the separation point was consid UWA611h. ered a resistant response, while disease severity greater
B. napus CB Trilogy B. napus Thunder TT 10 10 8 8 6 6 4 4 Frequency
2 Frequency 2 0 0 0 1 1.5 2 3 3.5 4.5 5 6 7 7.5 8 0 0.5 1 1.5 4 5 5.6 6 6.5 7 8 8.5 9 Disease severity Disease severity
B. napus Cresor-770 B B. nigra Introduce P.23845 9 7 8 6 7 6 5 5 4 4 3 3 2 Frequency 2 Frequency 1 1 0 0 3 4 5 6 7 8 0 1 2 0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5 5.5 6 2.5 3.5 4.5 5.5 6.5 7.5 0.5 1.5 Disease severity Disease severity
B. napus Tranby B. oleracea Brussels sprout 12 10 10 8 8 6 6 4 4 2 Frequency Frequency 2 0 0 0 0.5 1 1.5 2 3 4 5 5.5 6 7.5 8 8.2 0 1 1.5 2 3 4 5 5.5 6 6.5 7 7.5 Disease severity Disease severity
Figure 1 Bimodal plots reflecting frequencies of disease severity scores from 30 isolates of Hyaloperonospora brassicae on six Brassica genotypes (B. nigra Introduce P.23845, B. napus Tranby, CB Trilogy, Cresor 770 B and Thunder TT and B. oleracea Brussels sprout). Separation of resistance from susceptibility for each differential is as follows: disease severity rating <5 for B. napus accession Cresor 770 B; <4 for Tranby; <4.5 for B. nigra Introduce P.23845; <6 for B. napus Thunder TT; <3 for CB Trilogy and <2.5 for B. oleracea Brussels sprout. The basis of classifying Brassica genotype disease reactions across different H. brassicae isolates tested (expressed as different levels of disease severity) as bimodal was an assumption of two possible modes, one susceptible and the other resistant, providing that there was clear demarcation between these two modes in accordance with Goodwin et al. (1990). [Colour figure can be viewed at wileyonlinelibrary.com]
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1522 A. E. Mohammed et al.
Table 5 Severity of downy mildew disease on cotyledons of 28 Brassica genotypes inoculated with 11 isolates of Hyaloperonospora brassicae collected from southern Australia in 2015 and 2016.
Species Genotype WA5ca WA7c WA11c WA12c WA13c WA22c WA29c WA30c WA49c SA78c VC36c Means
Raphanus sativus Colonel 0.70 0.87 0.54 0.75 1.00 1.66 0.50 0.45 0.00 0.70 1.00 0.74 R. raphanistrum Wild radishb 0.75 0.83 1.41 0.45 1.00 0.87 0.62 0.37 0.95 1.16 1.00 0.86 Brassica carinata ATC 94011 1.79 1.00 2.00 1.00 1.00 0.29 1.00 1.00 1.04 1.58 1.00 1.15 Eruca vesicaria Yellow seedb 1.83 1.62 2.00 2.00 1.00 2.00 1.00 2.00 2.00 1.37 2.00 1.71 E. vesicaria Brown seedb 6.95 6.00 6.08 5.83 5.16 6.54 5.66 6.41 6.25 4.25 5.25 5.85 Crambe abyssinica CMB 94054 2.00 1.87 2.00 2.00 1.00 1.87 0.91 2.08 2.00 1.91 2.33 1.82 B. juncea Muscan 963 2.12 2.00 2.00 1.20 1.00 1.95 2.04 2.12 2.29 2.04 1.04 1.80 B. juncea Dune 2.00 1.95 2.00 1.83 1.00 1.62 2.00 2.00 1.95 2.04 1.29 1.79 B. nigra Introduce P.23845 2.00 1.08 1.45 2.00 1.00 0.95 1.00 0.66 2.00 1.62 1.87 1.42 B. rapa Xi Shui Bai 2.00 2.00 2.00 2.00 2.00 1.66 1.79 2.00 2.00 2.00 2.00 1.95 B. napus Cresor 771 B 3.41 5.95 2.79 6.83 1.16 6.33 5.04 3.87 2.54 3.16 2.81 3.99 B. napus Cresor 770 B 2.04 3.37 2.00 3.08 1.62 2.00 2.00 6.25 2.83 2.04 2.00 2.66 B. napus Komet 741 A 3.25 4.95 4.66 4.66 1.62 2.37 3.95 3.70 3.12 2.70 2.75 3.43 B. napus Komet 744 A 2.83 4.20 5.20 3.04 1.66 3.16 2.33 3.33 2.70 1.58 2.37 2.95 B. napus Capricorn C 5.54 5.75 3.45 5.25 5.12 3.95 4.91 7.08 5.54 5.25 5.27 5.19 B. napus AG Comet 6.87 6.29 4.66 6.20 5.45 5.62 5.37 6.20 4.66 5.33 5.22 5.62 B. napus Tranby 7.83 5.83 6.87 5.95 6.37 6.83 6.41 7.20 7.75 5.54 5.45 6.55 B. napus CB Trilogy 5.12 5.91 4.79 5.33 4.91 6.29 4.91 5.62 5.08 5.37 4.31 5.24 B. napus ATR Banjo 4.45 4.16 3.45 5.25 4.58 5.00 4.37 3.79 5.16 4.04 4.33 4.42 B. napus ATR Stubby 6.33 5.70 4.62 6.16 5.62 5.95 5.87 5.87 6.00 5.50 4.68 5.66 B. napus ATR Cobbler 5.50 5.54 5.29 5.37 4.12 6.08 5.70 7.00 4.58 4.08 5.25 5.32 B. napus Pioneer 45Y77 3.70 5.87 5.58 2.75 1.95 5.08 3.37 4.50 4.45 2.87 4.45 4.05 B. napus Thunder TT 8.41 8.45 8.12 8.37 7.87 8.75 8.37 8.83 8.70 7.29 8.41 8.32 B. oleracea Brussels sproutb 6.16 6.04 5.87 5.83 4.29 5.25 5.33 5.66 4.37 4.33 4.29 5.22 B. oleracea Cabbageb 3.29 3.62 3.00 3.58 4.12 4.29 3.83 5.41 4.08 4.29 3.72 3.93 B. oleracea Broccolib 4.50 4.79 5.33 4.20 5.04 4.41 2.75 4.04 5.58 3.29 5.37 4.48 B. oleracea Chinese broccolib 3.41 5.50 4.83 5.08 3.70 4.70 4.58 5.83 5.12 5.37 4.50 4.78 B. oleracea Cauliflowerb 6.20 5.25 5.50 4.79 4.87 5.66 4.37 4.75 5.00 3.70 6.41 5.14 Means 3.96 4.16 3.84 3.96 3.19 3.97 3.57 4.22 3.85 3.37 3.58 3.79
Disease severity assessed on a 0 9 scale, where 0 no disease, 9 heavy sporulation, cotyledon collapsed. Significance of cultivars P < 0.001, LSD at P < 0.05 0.158; significance of isolates P < 0.001, LSD at P < 0.05 0.099; significance of culti vars 9 isolates P < 0.001, LSD at P < 0.05 0.52. aWA, SA, VC + number + c 2015 2016 isolates. bCommon name. were overall more virulent than 2006 2008 isolates. R. raphanistrum (WA6.2wrc and WA6.1wrc) grouped Among the agrogeographical locations sampled in 2006 into a separate distinct clade II with bootstrap values of 2008, the predominant and most widely distributed 100%. pathotype was pathotype 47, constituting an overall fre quency 63.3% of total isolates, followed by the patho Discussion type 57 that was only found at a single agrogeographical location and at a frequency of 3.3% (Table 4; Fig. S1). This is the first study to define the phylogenetic relation In 2006 2008 isolates, pathotype 40 was also found ships of H. brassicae isolates in Australia and the first across multiple agrogeographical locations while the anywhere to use octal nomenclature to characterize remaining pathotypes were only found at a single agro pathotypes of H. brassicae. The latter provides a novel geographical location (Fig. S1). basis for standardizing phenotypic characterization of pathotypes of H. brassicae worldwide and monitoring and understanding changes in H. brassicae populations Sequencing and phylogenetic analysis over time and between locations. There were significant The phylogenetic tree highlighted seven distinct groups differences between pathogen isolates, across the diverse of H. brassicae isolates. Seventy percent of 2006 2008 host genotypes and a significant interaction between the isolates were distributed in clade I with bootstrap values two. Host responses ranged from nil, to a hypersensitive of 100% and the remaining 30% of 2006 2008 isolates response, to systemic spread and abundant pathogen were distributed in clade V with bootstrap values of sporulation. While isolates were generally most virulent 83.3% (Fig. 3). All 2015 2016 isolates were distributed on their host of origin, there were exceptions, such as among the seven clades. Two isolates from isolates from R. raphanistrum that were avirulent on
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pathogen diversity and/or distinct isolate types (e.g. The range in symptom variation across the different iso phoma stem canker, L. maculans, Balesdent et al., 2005; late/pathotype 9 host combinations was expected and is sclerotinia stem rot, S. sclerotiorum, Uloth et al., 2015) similar to the findings of Holub et al. (1994) on in southern Australia and the same may be true of Arabidopsis thaliana; those authors investigated seven H. brassicae. While the specific reasons for this have not H. arabidopsidis isolates 9 11 host genotypes and been definitively determined, it probably relates, at least observed a range of interaction phenotypes including in part, to the highly variable Mediterranean type envi localized hypersensitive type necrosis, more extensive cell ronment in southern Australia that maximizes preserva collapse, delayed sporulation, or complete susceptibility. tion of infested crop residues over the long dry and hot Likewise, Nashaat et al. (2004) identified various differen summer periods (Barbetti et al., 2012). tial expressions of resistance when a wide range of Hyaloperonospora brassicae isolates from R. raphani B. juncea genotypes was screened, highlighting 16 differ strum were reported as the most damaging and widespread ential host response groups (groups A P); for example, pathogens of this host species (Maxwell & Scott, 2008). groups A and B expressed the widest range in different However, this contrasts somewhat with the current study, resistance expression (highly resistant to susceptible) to where two H. brassicae isolates from R. raphanistrum did these isolates. However, most importantly, the current not cause any visible disease reaction on R. raphanistrum, study demonstrates that these new resistances can proba whereas these same isolates caused severe disease reaction bly be used to improve oilseed and vegetable brassica resis on most B. napus and B. oleracea genotypes tested. The tances to H. brassicae and can been sourced from a wide lack of visible disease symptoms on R. raphanistrum range of different Brassica species, such as those bearing B under the experimental conditions of the current investiga genomes, including B. juncea (AABB), B. nigra (BB), tion does not, however, exclude them from being hosts B. carinata (BBCC), and in other Brassicaceae hosts such under field conditions from which they were isolated, and as R. sativus, R. raphanistrum, C. abyssinica and E. vesi the possibility of factors for resistance being expressed caria. Nevertheless, it is also possible that some isolates differently under different environmental conditions can commonly associated with these other species may over not be ruled out. Raphanus raphanistrum is a common come the resistances highlighted in the current study, mak weed within and near to oilseed and vegetable brassica ing it important to also seek resistances for each species crop fields (Gunasinghe et al., 2016) and the present against their prevailing H. brassicae populations. results suggest that it may play a role not only as a carry This study reports the development of a set of host dif over host but also as a reservoir for H. brassicae to ferentials for the characterization of pathotypes of develop new strains. However, the extent that this occurs H. brassicae causing downy mildew on B. napus and naturally between R. raphanistrum and crop and/or veg some crucifer species in Australia. Despite the attempt of etable brassica crops under field conditions warrants fur Coelho et al. (2012) to characterize the pathotypes of ther investigation, as R. raphanistrum is deemed to play European isolates of H. brassicae from B. oleracea, the an important role for other important Brassicaceae dis current study is the first attempt to use octal nomenclature eases in both maintaining pathogen diversity and provid (Goodwin et al., 1990) to code H. brassicae isolates ing a reservoir of inoculum to infect neighbouring crops according to their pathogenicity/virulence (expressed as (Barbetti et al., 2012); examples of this include phoma disease severity) on the six newly identified host differen stem canker (Sivasithamparam et al., 2005) and white leaf tials. This same nomenclature system has been used to spot (Gunasinghe et al., 2016). There are genomic linkage delineate races of sunflower downy mildew (Plasmopara groups across the different Brassica species, for example, halstedii) by Tourvieille de Labrouhe et al. (2000). In the there are nine B. oleracea linkage groups conserved in the present study, the six genotypes used as differentials were B. napus map (Carlier et al., 2012; Vicente et al., 2012), different species within the genus Brassica (B. napus, and this may explain why isolates collected from B. nigra and B. oleracea) and these were grouped into two B. oleracea (CC genome) infected some B. napus (AACC sets according to their genetic inheritance and continuity genome). with the previous study of Nashaat & Rawlinson (1994). Raphanus sativus, R. raphanistrum, C. abyssinica, The 30 isolates that were used to define the pathotype E. vesicaria, B. carinata, B. juncea and B. napus Pioneer structure included 12 isolates from B. napus, five from 45Y77 were highly resistant to all H. brassicae isolates. B. oleracea, two from R. raphanistrum and the remainder Importantly, R. sativus was immune to all tested H. bras from B. napus canola crops across a wide geographical sicae isolates. Clearly, these genotypes have pathotype area. This host range used is representative of common independent resistance effective against all identified hosts of H. brassicae in southern Australia. Eight H. bras pathotypes, and such a resistance type is highly valued by sicae pathotypes were delineated using the differential set breeders. In contrast, while B. napus lines Cresor 770 and of Brassicaceae genotypes developed in the current study. Cresor 771, B. rapa and B. nigra were resistant to all iso However, there is scope for a future increase in numbers lates from B. oleracea and R. raphanistrum, they were of differentials, for example, to delineate up to 128 patho susceptible to one B. napus isolate (UWA11h). While types. This is important, as if eight pathotypes could be these latter resistances are clearly pathotype dependent, delineated from a random selection of 30 isolates, then it these are still highly valuable for deployment in regions is highly likely that additional pathotypes will be delin where one or more of these particular pathotypes prevail. eated as additional isolates obtained from different host
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species, different seasons and different locations are char Acknowledgements acterized. The approach used to characterize pathotype delineations is clearly conservative and based on separa The authors are grateful for the financial support of the tion of peaks in the histograms that represent susceptible Grains Research and Development Corporation (GRDC), versus resistant responses of host differentials to these as this work constitutes part of the GRDC UWA 170 pathotypes of H. brassicae. Characterization of patho project ‘Emerging foliar diseases of canola’, and the gen types is crucial for understanding the virulence diversity of erosity of seed companies in supplying seed of test vari this pathogen as well as for coordinating breeding efforts eties. Dr Nash Nashaat, formerly of Rothamsted to develop cultivars with pathotype independent and/or Research, UK, is gratefully acknowledged for the kind pathotype dependent resistances. provision of seed of B. napus lines Komet 741 A, Molecular phylogenetic analysis showed isolates dis Komet 744 A, Cresor 770 B, Cresor 771 B and Capri tributed across seven distinct clades. However, it is corn C. The authors gratefully acknowledge the technical noteworthy that two isolates from R. raphanistrum assistance of Xin Tian Ge and Hua Li with collection of (WA6.2wrc and WA6.1wrc) grouped into a separate historical isolates including DNA extraction from them clade II. Thus, this particular genetic divergence may and the technical support from Robert Creasy and Bill relate to the host of origin from which the isolates were Piasini in the University of Western Australia Plant recovered. Similarly, Maxwell & Scott (2008) found Growth Facilities. All authors declare that they have no that isolates of H. brassicae from R. raphanistrum conflict of interest in relation to this publication. grouped into a distinct clade. In addition, Goker€ et al. (2004) indicated that, in general, isolates from the same References species of the host appeared in the same clade, whereas other isolates recovered from B. napus grouped into a Bachofer M, 2004. Molekularbiologische Populationsstudien an separate clade. This may be similar for other Brassi Plasmopara halstedii, dem Falschen Mehltau der Sonnenblume. caceae pathogens, as Gunasinghe et al. (2016) also sug Stuttgart, Germany: University of Hohenheim, PhD thesis. Balesdent MH, Barbetti MJ, Li H, Sivasithamparam K, Gout L, Rouxel T, gested that while there is a very close genetic 2005. Analysis of Leptosphaeria maculans race structure in a world- relationship between Pseudocercosporella capsellae iso wide collection of isolates. Phytopathology 95,1061 71. lates recovered from B. napus canola crops, isolates Barbetti MJ, Khangura R, 2000. Fungal Diseases of Canola in Western recovered from R. raphanistrum showed some distinct Australia. Bulletin 4406. Perth, Australia: Agriculture Western differences genetically. Australia. Downy mildew remains a significant threat to the oil Barbetti MJ, Banga SS, Salisbury PA, 2012. Challenges for crop production and management from pathogen biodiversity and diseases seed and vegetable brassica industries in Australia and under current and future climate scenarios case study with oilseed elsewhere (Barbetti & Khangura, 2000). Resistance iden brassicas. Field Crops Research 127, 225 40. tified in this study and elsewhere across various Brassi Carlier JD, Alabacßa CA, Coelho PS, Monteiro AA, Leitao~ JM, 2012. The caceae 9 H. brassicae combinations will be critical for downy mildew resistance locus Pp523 is located on chromosome C8 breeding effective disease resistance. 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Tourvieille de Labrouhe DT, Gulya TJ, Masirevic S, Penaud A, Rachid Williams PH, 1985. Downy Mildew. Crucifer Genetics Cooperative KY, Viranyi F, 2000. New nomenclature of races of Plasmopara (CRGC) resource book. Madison, WI, USA: Department of Plant halstedii (sunflower downy mildew). In: Proceedings of the 15th Pathology, University of Wisconsin. International Sunflower Conference, Toulouse, France. [http://isasunf You M, Barbetti MJ, Sivasithamparam K, 2005. Characterization of lower.org/fileadmin/documents/aaProceedings/15thISCToulouse2000/ Phytophthora clandestina races on Trifolium subterraneum in PosterWorkshopI-O/I-F57.pdf]. Accessed 15 March 2018. Western Australia. European Journal of Plant Pathology 113, Uloth M, You MP, Cawthray G, Barbetti MJ, 2015. Temperature 267 74. adaptation in Sclerotinia sclerotiorum affects its ability to infect Brassica carinata. Plant Pathology 64, 1140 8. Supporting Information Vicente JG, Gunn ND, Bailey L, Pink DAC, Holub EB, 2012. Genetics of resistance to downy mildew in Brassica oleracea and breeding Additional Supporting Information may be found in the online version of towards durable disease control for UK vegetable production. Plant this article at the publisher’s web-site. Pathology 61, 600 9. Figure S1. Geographical distribution of eight different pathotypes of Voglmayr H, 2003. Phylogenetic relationships of Peronospora and Hyaloperonospora brassicae in the canola (Brassica napus) cropping related genera based on nuclear ribosomal ITS sequences. Mycological region in southwest Western Australia. Years shown are for date of Research 107, 1132 42. initial isolation for individual and/or groups of pathotypes nearest to the Wang M, Farnham MW, Thomas CE, 2001. Inheritance of true leaf indicative date shown. stage downy mildew resistance in broccoli. Journal of the American Table S1. GenBank accession numbers of some Hyaloperonospora Society for Horticultural Science 126, 727 9. brassicae isolates used in this study and their host of origin.
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Page 102 of 119 Chapter 6. General Discussion
Studies were undertaken on the incidence, importance, resistance, and the role of environmental factors and plant growth stage, pathogen phenotypic (e.g., pathotype) and phylogenetic variation, in the development of downy mildew
(Hyaloperonospora brassicae) epidemics on oilseed Brassicas across southern
Australia. These studies targeted issues that have not previously been investigated comprehensively for either Australian or other diverse Brassicaceae germplasm. Overall, these studies have provided new understanding of the extent and role of pathogen phenotypic and phylogenetic variation in Australian populations, definition of some of some major environmental (temperature) and plant growth stage factors in development of downy mildew epidemics on oilseed rape and mustard Brassicas and have highlighted novel host resistances.
Together, these will foster development and utilization of effective control strategies for downy mildew, particularly in oilseed Brassicas.
New highly resistant genotypes were identified among Australian canola
(Brassica napus and B. juncea) for the first time (Chapter 2). In addition, new sources of host resistances to H. brassicae were identified across diverse
Brassicaceae genotypes, particularly the high level resistances in R. sativus, B. carinata B. juncea, B. oleracea and C. abyssinica (Chapter 3). These important outcomes will enable use of these host resistances as a foundation for protecting oilseed Brassicas against this obligate pathogen in the future. Resistant genotypes identified in this study can, first, be utilized in canola and other
Brassicaceae breeding programs to develop resistant varieties for the canola other Brassicaceae oilseed and horticultural industries to manage downy mildew disease. Second, resistant varieties identified as having agronomically suitable
Page 103 of 119 backgrounds could be directly deployed into regions where downy mildew is prevalent to provide relief for growers. Chapter 4 highlights the significant effects of temperature and plant age in driving the severity of downy mildew epidemics on oilseed B. napus and B. juncea. The findings explain, for the first time, why disease epidemics in oilseed rape fields in Australia have been most severe on the more highly susceptible seedlings rather than adult plants, and why downy mildew is favoured by increasing seasonal temperatures occurring as a consequence of climate change. Studies defining phylogenetic relationships and pathotypes (Chapter 5) provided an evolutionary perspective on this oomycete pathogen and set a benchmark for understanding current and future genetic and phenotypic (pathotype) shifts within the pathogen population in Australia. Further, a comprehensive host differential set developed provides the first opportunities for monitoring changes in H. brassicae populations both in Australia and elsewhere, and will allow early warning of new virulent pathotypes able to overcome any resistances currently deployed commercially.
Of particular importance is the novel, high-level host resistances highlighted and these will allow development and utilization of effective host resistance as a foundation for much improved control strategies for downy mildew, particularly in oilseed Brassicas. The studies in this PhD significantly add to previous historical studies to locate specific resistance to H. brassicae pathogen among Brassica species. For instance, Natti et al. (1967), Farnham et al. (2002) and Monteiro et al. (2005) reported resistance to downy mildew in B. oleracea, Lucas et al. (1988) and Nashaat et al. (1997) reported resistances in B. napus, and Nashaat & Awasthi (1995), Nashaat et al. (2004) and Chattopadhyay
& Séguin-Swartz (2005) noted resistance to H. brassicae in B. juncea. While all these historical studies indicated that the resistance to downy mildew brassicas
Page 104 of 119 in can be either controlled by one specific gene or multiple genes, the nature of the genetic control of resistance was not determined in the current study and future studies to determine such would be beneficial.
Devastating outbreaks of downy mildew caused by H. brassicae since the early 1970s demonstrated the capacity of this disease to impact severely on canola production, particularly in high- and medium-rainfall areas, across southern Australia. This is not surprising as most varieties commercially grown in
Australia have a relatively low level of resistance to this pathogen and some are higly susceptible. Further, the current studies not only provide the first understanding of the diversity of Australian H. brassicae populations, the unique set of host differentials has enabled characterisation of highly virulent pathotypes in the Australian pathogen population. Consequently, the prevalence of serious downy mildew outbreaks, especially in Western Australia, is not surprising. There is an urgent need to develop appropriate management strategies based on host resitance to prevailing pathotypes, strategies that allow effective control of this important disease of oilseed Brassicas. As the use of fungicides or agricultural strategies to control downy mildew pathogen offers only limited or short term control and with high economic cost (Eshraghi et al., 2007; Barbetti et al., 2011;
Neik et al., 2017), this combined new knowledge on new host resistances
(Chapters 2 and 3) and understanding of pathogen populations (Chapter 4) has, for the first time, enabled new opportunities towards more effective management of downy mildew using host resistance.
New resistant genotypes identified in Chapter 2 have enhanced prospects for obtaining more effective control and management of this important disease
(Thomas & Jourdain 1992) through utilisation of these new host resistances.
Despite of the attempt of Ge et al. (2008) to estimate the level of resistance in
Page 105 of 119 some genotypes of B. napus, the current study is the first to demonstrate the existence of these very high levels of pathotype-independent resistance to H. brassicae, particularly in Australian canola varieties. The methods used were appropriate, as screening for host resistance at the cotyledon stage is known to be effective (Lucas et al., 1988; Dickson & Petzoldt 1993; Gröuntoft 1993;
Nashaat & Awasthi 1995; Silué et al., 1996; Nashaat et al., 2004), not just for identifying resistances for Brassica seedlings under controlled environment conditions, but that such resistances identified operate in leaves of plants in the field (e.g., Jensen et al., 1999; Wang et al., 2000; Zhang et al., 2012). The very high levels of pathotype-independent resistance found (Chapter 3) highlight how these new sources of host resistance available across miscellaneous
Brassicaceae species to H. brassicae, particularly oilseed, B. carinata, weedy and the wild crucifer species, can be utilized to improve the management of this important pathogen across the oilseed and vegetable Brassicaceae industries.
As highlighted above, this can be achieved, first, by their utilisation as sources of resistance in breeding programs to produce resistant varieties, and, second, by the fact that in many instances they could be directly deployed as new varieties where downy mildew is prevalent.
While H. parasitica has been present in Australian canola and mustard crops for decades (Barbetti & Khangura 2000), the reasons for its increasing severity and threat to commercial canola crops in some regions of Australia, particularly since 1999, have not been evident until this current study. The current study strongly suggests that increasing incidence, severity and threat from downy mildew is likely linked to increasing temperatures, particularly at the seedling stage when plants are most susceptible. The current study highlighted for the first time how temperature and plant age together drive downy mildew disease
Page 106 of 119 epidemics on oilseed B. napus and B. juncea in Australia (Chapter 4). It largely explains why seedlings are particularly severely attacked, and why both incidence and severity have increased in Australia in conjunction with rising temperatures over recent decades due to climate change (Barbetti et al., 2012; Jones & Barbetti
2012). Findings of these studies confirm a previous study on B. oleracea var. capitata (cabbage) cotyledons by Achar (1998) where 90-100% infection rapidly occurred at 15-25⁰C. Further, it is clear from crop surveys in 2015 and 2016 that most severe downy mildew epidemics on canola occur in regions such as southwest Western Australia when crops are subject to warmer temperatures and more variable rainfall in the autumn-early winter period (Barbetti et al., 2012).
Downy mildew can be extremely severe when rainfall in autumn/early winter rainfall is much lower than average (MP You and MJ Barbetti, unpubl.). The findings of the current study explains the development of the most severe downy mildew epidemics in Australia on seedlings of susceptible varieties early in the growing season when warmer temperatures coincide with presence of the more susceptible seedlings rather than with the combination of less susceptible older plants and cooler temperatures as the growing season progresses during the winter months. Likewise, results of these studies indicate that the ongoing increasing temperatures associated with climate change will likely make downy mildew epidemics in Australia even more widespread and severe in future years.
Studies involving pathogen pathotypes (Chapter 5) applied, for the first time in Australia or elsewhere, the octal nomenclature system to characterize pathotypes of H. brassicae. Evidence of pathogenic and virulence variability among Australian isolates of H. brassicae was evident with 13 distinct disease reactions recorded from 30 isolates (2006–2008 and 2015–2016) inoculated across 28 Brassicaceae genotypes. This was not unexpected, as there was
Page 107 of 119 already molecular evidence confirming existence of genetic differences within H. brassicae (e.g., Choi et al., 2003; Göker et al., 2003; Voglmayr, 2003). With the exception of results for R. raphanistrum, the findings of the current study substantiated earlier reports that isolates of H. brassicae obtained from different
Brassica species are most virulent on their species of origin, but nevertheless able to infect, although generally not as well, other related species (e.g., Chang et al., 1964; McMeekin, 1969; Dickinson & Greenhalgh, 1977; Sherriff & Lucas,
1990; Silué et al., 1996; Vicente et al., 2012). Importantly, the current study
(Chapter 5) highlights existence of sufficient cross-species virulence to enable H. brassicae isolates derived from one Brassica species to be phenotypically characterized across other Brassicaceae species.
Molecular phylogenetic analysis showed isolates distributed across seven distinct clades. However, it is noteworthy that two isolates from R. raphanistrum
(WA6.2wrc and WA6.1wrc) grouped into a separate clade. This particular genetic divergence likely relates to the particular host of origin from which the isolates were recovered. Similarly, Maxwell & Scott (2008) found that isolates of H. brassicae from R. raphanistrum grouped into a distinct clade. In addition, Göker et al. (2004) indicated that, in general, isolates from the same species of the host appeared in the same clade, whereas other isolates recovered from B. napus grouped into a separate clade. The situation demonstrated in the current study regarding R. raphanistrum is particularly important, as this weedy crucifer species is prevalent throughout canola growing regions in southwest and southern
Australia. These same regions also show distinct diversities for other pathogens of canola, including white leaf spot (Pseudocercosporella capsellae)
(Gunasinghe et al., 2016), blackleg crown canker (Leptosphaeria maculans) (Li et al., 2004) and white rust (Albugo candida) (Kaur et al., 2008). The reasons for
Page 108 of 119 the general high pathogen diversity across these geographic regions warrant further exploration.
Future research
As many highly resistant genotypes have now been identified (Chapters 2 and 3), it remains important to utilise these in breeding programs to develop more resistant oilseed and vegetable Brassicaceae genotypes to downy mildew.
Commercial production of genotypes with resistance to prevailing H. brassicae pathotypes in Australia and elsewhere is now feasible as a consequence of these studies. Additional epidemiological studies are warranted to build on the encouraging results (Chapter 4) that explain current and predict future increasingly severe disease from H. brassicae in line with warming temperatures under current and future climate scenarios. Host differentials that can define distinct pathotypes (Chapter 5) pave the way for future studies to expand and utilise the differential set to characterise and compare H. brassicae pathotypes across crucifer taxa worldwide. Further, the reasons for the general high pathogen diversity across these geographic regions warrant further exploration.
Genome sequencing of H. brassicae has opened many new approaches to locating and understanding specific and quantitative resistant genes associated with the new resistances in Brassicaceae identified in the current study (Chapters
2 and 3). In addition, it was observed during these studies that downy mildew was also more severe when autumn/early winter rainfall was significantly below average. This may be due to such an environment directly favouring H. brassicae per se and/or from less competition for infection sites on seedlings due to reduced incidence of other diseases like white leaf spot and blackleg leaf under such conditions and this issue warrants further investigation. Finally, the reasons for
Page 109 of 119 the general high H. brassicae diversity across southern Australia warrants further exploration.
To conclude, the current study has highlighted the first pathotype- independent host resistances in Australian canola varieties to H. brassicae, resistances that should be effective across different regions where canola is grown. Effective resistances to downy mildew were also found across diverse sets of oilseed, weedy and the wild crucifer taxa. Together, these new resistances will play a critical role in the management of this important disease. The current study also showed how temperature and plant age are the key determinants of disease epidemics and explain for the first time why the incidence and severity of downy mildew have increased significantly over the past decade and a half when seasonal temperatures have been increasing as a consequence of climate change, and why downy mildew is particularly severe on seedlings. The outcomes of this research provide the urgently needed opportunity for breeders to incorporate effective resistance against specific and multiple pathotypes of downy mildew into new canola varieties. Moreover, and the current studies set a new benchmark for understanding current and future genetic and phenotypic shifts within pathogen populations. This will enable, for the first time, rapid identification of H. brassicae pathotypes for breeders in Australia and elsewhere, allowing them to target resistances effective against the prevailing H. brassicae pathotypes.
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Page 115 of 119 Appendix and Supporting Information
Oral presentations from this thesis by the author
1. 2016 Rottnest Postgraduate Summer School
The University of Western Australia, School of Agriculture and Environment. 1
Feb 2016.
Oral Presentation: Downy Mildew on oilseed Brassicas – understanding the drivers of disease epidemics and potential of novel host resistances http://www.plants.uwa.edu.au/student/rottnest
2. 2017 Rottnest Postgraduate Summer School
The University of Western Australia, School of Agriculture and Environment. 6
Feb 2017.
Oral Presentation: New resistances offer opportunity for effective management of the downy mildew (Hyaloperonospora parasitica) threat to canola http://www.plants.uwa.edu.au/student/rottnest
Page 116 of 119 Supporting Information for Chapter 5
Table S1. GenBank accession numbers of some Hyaloperonospora brassicae isolates used in this study and their host of origin
DM isolate Host species State GenBank accession no.
WA1.2c Brassica napus Western Australia MG757758 WA1.3c B. napus Western Australia MG757759 WA4.2c B. napus Western Australia MG757760 WA4.5c B. napus Western Australia MG757761 WA5c B. napus Western Australia MG757762 WA6.1wrc Raphanus Western Australia MG757763 raphanistrum WA6.2wrc R. raphanistrum Western Australia MG757764 VC36c B. napus Victoria MG757765 WA23.1c B. napus Western Australia MG757766 WA7.1.3c B. napus Western Australia MG757767 WA6.1.7c B. napus Western Australia MG757768 UWA11h B. napus Western Australia MG757769 UWA24h B. napus Western Australia MG757770 UWA31h B. napus Western Australia MG757771 UWA40h B. napus Western Australia MG757772 UWA51h B. napus Western Australia MG757773 UWA56h B. napus Western Australia MG757774 UWA61h B. napus Western Australia MG757775 UWA65h B. oleracea Western Australia MG757776 UWA67h B. oleracea Western Australia MG757777 UWA73h B. oleracea Western Australia MG757778 UWA74h B. oleracea Western Australia MG757779 UWA83h B. oleracea Western Australia MG757780
Page 117 of 119 WA5.3c B. napus Western Australia MG757781 WA11c B. napus Western Australia MG757782 WA11.4.2c B. napus Western Australia MG757783 WA12c B. napus Western Australia MG757784 WA12.3.5c B. napus Western Australia MG757785 WA13c B. napus Western Australia MG757786 WA30c B. napus Western Australia MG757787 SA78c B. napus South Australia MG757788 WA5.1c B. napus Western Australia MG757789 WA9.2c B. napus Western Australia MG757790 WA25.3.1c B. napus Western Australia MG757791 WA7.1.4c B. napus Western Australia MG757792
Page 118 of 119
Figure S1. Geographical distribution of eight different pathotypes of
Hyaloperonospora brassicae in the canola (Brassica napus) cropping region in southwest Western Australia. Years shown are for date of initial isolation for individual and/or groups of pathotypes nearest to the indicative date shown.
Page 119 of 119