Canola and pulse disease management; maintaining the vigilance
Author: Kurt Lindbeck, Stephen Marcroft, Angela Van de Wouw, Vicki Elliott and Barb Howlett | Date: 11 Feb 2014
Kurt Lindbeck1 , Stephen Marcroft2, Angela Van de Wouw2, 3, Vicki Elliott2 and Barb Howlett3
1NSW - Department of Primary Industries, Wagga Wagga 2Marcroft Grains Pathology P/L, Horsham 3The University of Melbourne.
Take home messages
- Elevated levels of internal blackleg infection were detected across southern NSW at blackleg monitoring sites in all varieties evaluated in 2013.
- Early flowering canola crops in combination with wet weather conditions were conducive to sclerotinia stem rot development in 2013.
- Consider the past frequency of sclerotinia stem rot outbreaks and yield potential when deciding to apply a foliar fungicide in 2014.
- Make informed decisions about blackleg and sclerotinia stem rot management. Monitor crops during the growing season to understand the impact of these diseases on production.
- Consult the Sclerotinia Stem Rot in Canola factsheet and Blackleg Management Guide for further information. These publications are available from the GRDC website.
- Early sown pulse manure crops are more prone to developing disease.
Blackleg of Canola
Blackleg – the challenge continues
The blackleg fungus, Leptosphaeria maculans is sexually reproducing, resulting in enormously diverse populations, and therefore, a high propensity to overcome resistance in Brassica napus (canola) cultivars. Therefore, the fungal population evolves very rapidly and responds quickly to selection pressures such as wide-scale sowing of cultivars with specific resistance genes. This will lead to resistance being overcome when cultivars of the same resistance gene are sown for three or more years. Cultivar resistance has been overcome in many regions around Australia, the most recent being Hyola®50 which went from a rating of resistant to susceptible on the Eyre Peninsula in 2012.
There is a very strong relationship between the intensity of canola production within a region and the level of blackleg development within commercial crops. The blackleg pathogen survives and reproduces on the previous season’s canola stubble, therefore the 500,000 ha canola crop in NSW in 2013 will result in 500,000 ha of blackleg infested stubble in 2014 releasing windblown spores every time it rains.
Blackleg – the warning signs for southern NSW
Cultivars representing each of the blackleg resistance groups were sown at 32 National Variety Trial sites across Australia (ten sites were located in NSW) and monitored for levels of blackleg development in 2013. Each site contained a representative cultivar of each of the six blackleg resistance groups, Groups A, B, C, D, E and G. There was no fungicide applied to seed, fertiliser or the growing plot (foliar) at these blackleg monitoring sites. These data indicate which resistance groups have higher levels of disease compared to the national average at each of the regionally based NVT canola yield sites and serve as a monitoring tool of local blackleg pathogen populations.
Overall blackleg severity increased across all cultivars at blackleg monitoring sites in 2013 in southern NSW (see Table 1). Blackleg severity increased from 26% average internal infection level in 2012 to 38% in 2013. The blackleg severity in NSW in 2013 was twice as high compared to infections levels in Victoria and South Australia. This increase in disease severity is likely due to the increasing area sown to canola in NSW since 2010.
In addition to overall increased blackleg severity, the Group D monitoring cultivar had a marked increase in blackleg severity. When similar increases in blackleg severity in Group D were detected on the Eyre Peninsula in South Australia in 2011, the Group D cultivars showed increased susceptibility to blackleg in the following season (2012). This situation could potentially occur in some regions of NSW in 2014.
Use the appropriate management strategy to minimise yield loss
Spores of the blackleg fungus are released from the previous year’s canola stubble, so an increased area of canola results in increased disease pressure. The most effective blackleg management tool is to keep a 500 m distance from this season’s crop and last year’s canola stubble. However, as more canola is grown this control measure is becoming harder to achieve, particularly in tight wheat/canola rotations.
Blackleg can be minimised by a number of factors including the sowing of cultivars with high blackleg resistance, avoiding last year’s stubble and applying the appropriate fungicides (see 2014 Blackleg Management Guide for details - www.grdc.com.au). An additional method for minimising disease is rotating cultivars with different resistance genes. All canola cultivars are now classified into different resistance groups. Refer to the current Blackleg Management guide (www.grdc.com.au) for individual cultivar groups.
Remember to monitor the level of blackleg development in canola crops during the growing season as a basis for selecting appropriate management strategies in the future.
Table 1. Summary data of all Australian blackleg monitoring sites for levels of internal infection
Cultivars representing each of the resistance groups were sown adjacent to canola National Variety Trial sites across Australia and monitored for levels of blackleg. These data indicate which resistance groups have high levels of disease compared to the national average at each site. For more detail consult the individual site summaries and recommendations on the NVT online website.
|
Site |
Resistance Group |
Comments |
|||||
|---|---|---|---|---|---|---|---|
|
NSW |
A |
B |
C |
D |
E |
G |
|
|
BECKOM |
H |
H |
M |
M |
L |
L |
High blackleg severity in groups A, B. Moderate in C, D. |
|
BELLATA |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
COOTAMUNDRA |
H |
H |
L |
L |
L |
L |
High blackleg severity in groups A, B. |
|
CUDAL |
H |
H |
H |
H |
L |
L |
High blackleg severity in groups A, B, C and D. |
|
GEROGERY |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
GRENFELL |
H |
M |
L |
L |
L |
L |
High blackleg severity in group A. Moderate in group B. |
|
LOCKHART |
H |
H |
L |
M |
L |
L |
High blackleg severity in groups A and B. Moderate in group D. |
|
MULLALEY |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
PARKES |
H |
H |
M |
L |
L |
L |
High blackleg severity in groups A and B. Moderate in group C. |
|
WAGGA WAGGA |
H |
H |
H |
H |
L |
L |
High blackleg severity in groups A, B, C and D. |
|
SA |
A |
B |
C |
D |
E |
G |
|
|
ARTHURTON |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
BORDERTOWN |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
MT HOPE |
L |
L |
L |
H |
L |
L |
High blackleg severity in Group D. |
|
RIVERTON |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
SPALDING |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
TURRETFIELD |
H |
M |
L |
L |
L |
L |
High blackleg severity in group A. Moderate in Group B. |
|
VIC |
A |
B |
C |
D |
E |
G |
|
|
CHARLTON |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
DIGGORA |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
HAMILTON |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
KANIVA |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
MINYIP |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
STREATHAM |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
WUNGHNU |
L |
H |
M |
L |
L |
L |
High blackleg severity in group B. Moderate in group C. |
|
YARRAWONGA |
H |
H |
L |
H |
L |
H |
High blackleg severity in groups A, B, D and G. |
|
WA |
A |
B |
C |
D |
E |
G |
|
|
BADGINGARRA |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
CORRIGIN |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
GIBSON |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
KATANNING |
L |
M |
L |
L |
L |
L |
Moderate blackleg severity in groups A and B. |
|
KENDENUP |
L |
M |
L |
L |
L |
L |
Moderate blackleg severity in group B. |
|
KOJONUP |
L |
M |
L |
L |
L |
L |
Moderate blackleg severity in group B. |
|
S. STIRLING |
L |
L |
L |
L |
L |
L |
Low blackleg severity in all groups. |
|
WILLIAMS |
L |
M |
L |
L |
L |
L |
Moderate blackleg severity in group B. |
Key
|
No data |
|
|
L |
Low blackleg severity compared to national average – continue with current management techniques. |
|
M |
Moderate blackleg severity compared to national average – Monitor crops for disease, see Blackleg management guide. |
|
H |
High blackleg severity compared to national average – high risk of yield loss, see Blackleg management guide. |
Sclerotinia Stem Rot – the new disease challenge
How does the disease develop?
The fungal pathogen that causes sclerotinia stem rot is called Sclerotinia sclerotiorum. This fungus can infect over 300 plant species, mostly broadleaf plants, including many crop, pasture and weed species. This includes plants like canola, lupin, chickpea, sunflower, lucerne, cape weed, and shepherds purse. The main features of the disease are:
- Airborne spores of the fungus are released from apothecia (a small, golf tee shaped structures, 5 – 10 mm in diameter) which germinate from sclerotia in the soil. For this to occur prolonged moist soil conditions in combination with moderate temperatures of 15°C to 25°C are considered ideal. Most sclerotia will remain viable for up to 3 – 4 years then survival slowly declines.
- Spores of the sclerotinia pathogen cannot infect canola leaves and stems directly. They require petals as a food source for spores to germinate grow and colonise the petal. When the infected petal eventually drops, it may become lodged onto a leaf, within a leaf axil or at branch junctions along the stem. If conditions are moist the fungus grows out of the petal and invades healthy plant stem tissue which will result in a stem lesion and production of further sclerotia within the stem which will be returned to the soil after harvest.
- Sclerotia also have the ability to germinate in the soil, produce mycelium and directly infect canola plants in close proximity, causing a basal infection.
- Weather conditions during flowering play a major role in determining the development of the disease. The presence of moisture during flowering and petal fall will determine if sclerotinia develops. Dry conditions during this time can quickly prevent development of the disease, hence even if flower petals are infected, dry conditions during petal fall will prevent stem infection development.
Research findings in 2013
A number of commercial canola crops were monitored for the development of sclerotinia stem rot in 2013. These crops were around Cootamundra and south of Henty, in traditionally high disease risk districts. Results from observations within these crops found a very strong relationship between leaf wetness and stem rot development. While the level of stem rot development varied between the crops south of Henty and those at Cootamundra, it was found those extended periods of continual leaf wetness of at least 48 hours or longer were critical ‘trigger’ points for stem rot development in both regions.
There was also two distinct phases identified in the development of the disease. It was found that petal infection provided the first phase in the initial establishment of stem rot within the crop. The second phase occurred once canopy closure occurred and a humid microclimate was established, with the retention of infected plant tissue under the crop canopy providing opportunities for continued disease development later in the season. This tissue included lower leaves and senescent leaves that became colonised and later adhered to stems, causing stem lesion development and yield loss. This work will continue in 2014 to collect and collate data which will be used to develop a disease prediction model for NSW.
Where did the disease occur in 2013?
In 2013 epidemics of sclerotinia in southern NSW and northern Victoria were observed in traditionally high rainfall districts. These included districts east of Cootamundra, Young and Cowra, south of Henty, around Corowa and Howlong and districts along the Murray River. Infection levels observed in some crops were as high as 30 – 60%. In other districts, crop infection levels were generally low.
Why did we observe higher levels of sclerotinia stem rot in 2013?
The weather conditions during the winter of 2013 could be considered ideal for the development of sclerotinia stem rot. Mild winter temperatures resulted in many canola crops flowering 3 – 4 weeks earlier than would be considered ‘normal’ for southern NSW and northern Victoria. Canola crops were observed to be flowering as early as the middle of July. These flowering crops also coincided with good rainfall throughout late July and August, which provided ideal conditions for apothecia development and release of ascospores. Frequent rainfall events throughout August provided long periods of leaf wetness and ideal conditions for infected petals to drop into wet crop canopies and allow infection to occur.
What are the indicators that sclerotinia stem rot could be a problem in 2014?
- Epidemics of sclerotinia stem rot generally occur in districts with reliable spring rainfall and long flowering periods for canola.
- Use the past frequency of sclerotinia stem rot outbreaks in the district as a guide to the likelihood of a sclerotinia outbreak. Paddocks with a recent history of sclerotinia are a good indicator of potential risk, as well as those paddocks that are adjacent.
- The commencement of flowering can determine the severity of a sclerotinia outbreak. Spore release, petal infection and stem infection have a better chance of occurring when conditions are wet for extended periods, especially for more than 48 hours. Canola crops which flower earlier in winter, when conditions are cooler and wetter, are more prone to disease development.
If I had sclerotinia in my canola crop last year, what should I do this season?
The biggest challenge in managing sclerotinia stem rot is deciding whether or not there is a risk of disease development and what will be the potential yield loss. Research in Australia and Canada has shown that the relationship between the presence of the pathogen (as infected petals) and development of sclerotinia stem rot is not very clear due to the strong reliance on moisture for infection and disease development.
Important management options include:
- Sowing canola seed that is free of sclerotia. This applies to growers retaining seed on farm for sowing. Consider grading seed to remove sclerotia that would otherwise be sown with the seed and infect this season’s crop.
- Separate this season’s paddock away from last year’s canola stubbles. Not only does this work for other diseases such as blackleg, but also for sclerotinia.
- Rotate canola crops. Continual wheat/canola rotations are excellent for building up levels of viable sclerotia in the soil. A 12 month break from canola is not effective at reducing sclerotial survival. Consider other low risk crops such as cereals, field pea or faba bean.
- Follow recommended sowing dates and rates for your district. Canola crops which flower early, with a bulky crop canopy are more prone to developing sclerotinia stem rot. Bulky crop canopies retain moisture and increase the likelihood of infection. Wider row spacing’s can also help by increasing air flow through the canopy to some degree until the canopy closes.
- Consider the use of a foliar fungicide. Weigh up yield potential, disease risk and costs of fungicide application when deciding to apply a foliar fungicide.
- Monitor crops for disease development and identify the type of stem infection. Main stem infections cause the most yield loss and indicate infection events early in the growing season. Lateral branch infections cause lower levels of yield loss and indicate infection events later in the growing season.
When is the best time to apply a foliar fungicide?
Research in Australia and Canada has shown that an application of foliar fungicide around the 20% - 30% flowering stage (20% flowering is 14 – 16 flowers on the main stem, 30% flowering is approx. 20 flowers on the main stem) can be effective in reducing the level of sclerotinia infection. The objective of the fungicide application is to prevent early infection of petals while ensuring that fungicide also penetrates into the lower crop canopy to protect potential infection sites (such as lower leaves, leaf axils and stems). Timing of fungicide application is critical.
In 2013 some commercial crops which received an application of foliar fungicide still developed stem rot later in the season. This is not unexpected as the fungicide will have a limited period of protection during a time of rapid plant growth and the main aim of foliar fungicide applications is the prevention of main stem infections, which cause the greatest yield loss. Development of lateral branch infections later in the season is not uncommon, and will cause lower yield loss.
Consult the Sclerotinia Stem Rot in Canola factsheet for further information. This publication is available from the GRDC website.
Diseases of Pulse Crops in 2013
Disease issues which appeared in pulse crops in the southern region in 2013 include:
Phytophthora root rot of lupin:
Wet winter conditions throughout July and August favoured the appearance of this disease in some districts. Lupin crops sown in lower lying paddocks or paddocks with a hard pan layer which favoured waterlogging, may have developed patches of dying plants in spring. The pathogens which cause phytophthora root rot (also known as sudden death in the past) only require a brief period (short as 8 hours) of waterlogging for infection of roots to occur.Bacterial blight of field pea:
This disease was not seen or reported in any commercial field pea crops in 2013. The mild winter temperatures experienced in 2013 did not favour development of bacterial blight, with a strong relationship between frost events and the development of the disease.
Powdery mildew of field pea:
Mild winter temperatures and rapidly growing crops resulted in outbreaks of powdery mildew in some districts further west and north of Wagga Wagga. Traditionally this disease does not appear in crops till later in the season during late flowering and pod fill. Mild daily temperatures in combination with cool nights, which favour dew formation, are ideal for this disease to develop. There are a number of foliar fungicide options available and resistant varieties to manage this disease.
Manuring Pulse Crops and Disease Management – striking the balance
A number of disease issues developed in 2013 with the increasing trend toward using pulse crops as manure in southern NSW and northern Victoria, in particular blackspot (or ascochyta blight) of field pea. If pulse crops are to be used successfully for manuring purposes the balance has to be made between dry matter production and disease management. Essentially, many pulse crop species have not been developed as manure crops and the agronomy and disease management packages which accompany these crops traditionally focus on grain production.
Field pea is sensitive to early sowing. Early sown field pea crops are more prone to developing blackspot if conditions are wet in winter or developing bacterial blight if conditions are dry and frosty. The traditional sowing window for this crop in our region has been developed around maximising yield and avoiding disease, in particular avoiding spore release from old field pea stubble. The pathogens which cause blackspot can survive between seasons on seed, in soil and on stubble, which means an integrated approach must be taken to manage this disease. Effective disease management options include:
- Use of a fungicidal seed dressing: To reduce seed transmission of the disease and provide early seedling protection (products such as P-Pickle-T).
- Crop rotation: A break of at least three years to ensure adequate time between field pea crops for soil-borne spore populations to decease.
- Paddock selection: Do not sow this year’s field pea crop adjacent to last year’s field pea stubble which will release air-borne spores onto new season’s crops. Leave a distance of at least 500m between last year’s stubble and this year’s field pea crop.
- Time of sowing: Do not be tempted to sow crops too early outside the recommended sowing window for your district. Early sowing will expose crops to early season spore showers and allow crops to develop a dense canopy by mid – late winter, which further favours disease development. Early sown crops are also more prone to bacterial blight by increasing exposure to frost events.
Why did we see high levels of blackspot last year?
High levels of blackspot were observed in field pea manure crops in several districts in southern NSW last year. These blackspot epidemics were due to a combination of factors which favoured development of the disease. These factors included:
- Crops sown extremely early.
- Field pea crops sown adjacent to the previous season’s field pea stubble.
- Dry conditions over summer which did not allow any spore maturation or release to occur prior to sowing.
In previous seasons the wet conditions over summer had aided breakdown of field pea stubble and accelerated spore maturation and release in the field prior to sowing. This had allowed early sown field pea crops to effectively ‘escape’ early spore showers from the previous season’s stubble and develop only low levels of disease.
Disease prediction using Blackspot Manager
Primary infection of blackspot can be reduced if field pea crops are sown after the majority of blackspot spores have been released from infected field pea stubble. Consequently pea growers have generally been advised to sow pea crops 2-3 weeks after opening rains so newly emerging crops can avoid these spores. However the timing of the spore release varies depending on seasonal conditions over summer and autumn.
For several years field pea producers in Western Australia have had access to ‘Blackspot Manager’, which is a computer based model which predicts the best time to sow new season field pea crops to avoid disease. Blackspot Manager calculates the timing of spore release from old stubble using seasonal rainfall and temperature data, and identifies whether the delay in sowing is necessary in the current season or whether it is safe for peas to be sown during the autumn. The optimum sowing dates calculated also consider agronomic factors, which may vary by region and season, and other production issues such as frost and yield penalties from late sowing.
In recent years this service has been extended to field pea producers in South Australia and Victoria. Over the past two years data from southern NSW has been supplied to researchers in DAFWA to validate the model for this region. It is hoped that this service will be available this coming season. For more information refer to http://www.agric.wa.gov.au/cropdisease
Choosing appropriate crops to manure
To overcome the issues relating to disease development in manure crops the choice of manure crop should be considered and how this fits within the farming system. For some producers time of sowing will be seen as an important consideration. For producers who prefer to sow manure crops early, crops such as lupin or high density legume species would be suitable. For a later sowing time, field pea or vetch would be appropriate. Even within some of these crop types there are varietal differences that can be exploited if dry matter production (and hence nitrogen fixation) is the objective. For example, within field pea, varieties such as Morgan or PBA Hayman produce more dry matter, even when sown within the recommended sowing window, than other high yielding varieties.
Unfortunately there exists a conflict between dry matter production and disease development. Crops sown to maximise dry matter production will succumb to disease more readily and the manure crop fails to reach potential. Advisers and producers need to weigh up the potential risks, limitations and costs of the various manure crop options and choose a crop that best suits their particular farming system.
Contact details
Kurt Lindbeck
NSW Department of Primary Industries, Wagga Wagga Agricultural Institute
02 69 381 608
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