Chickpeas: reports from 2012 and implications for 2013 Qld

Kevin Moore NSWDPI Tamworth, Malcolm Ryley DAFFQ Toowoomba, Murray Sharman DAFFQ Brisbane, Joop van Leur, NSWDPI Tamworth

The 2012 northern NSW/southern QLD chickpea season

The 2012 chickpea season in northern NSW/southern QLD followed a wet start to the calendar year (in some places very wet), had above average post-sow rain in June/July and then well below average rain for the rest of the growing season (Dalby’s October/November rain was just below average).  Saturated soil in the early part of the chickpea season resulted in waterlogging and in certain paddocks Phytophthora root rot.  From mid-late August to mid-late October, average daily temperatures dropped below 15^oC and this delayed pod set and caused flower and pod abortion; severe frosts in September killed flowers and pods.  In early September to early October, in some locations, chickpea roots encountered saline conditions which quickly led to death of affected areas.  In general, plant diseases caused few problems across the region.  However, in some districts viruses caused serious losses from mid-late September.  Herbicides and herbicide residues continue to be a concern as recommended plant pack periods or rates are not followed.  Overall, yields were below average – a result of water logging, low temperature, frost, a dry spring and in some locations salinity and virus.

Wet feet (waterlogging) and Phytophthora root rot

Phytophthora medicaginis, the cause of phytophthora root rot (PRR) of chickpea is endemic and widespread in southern QLD and northern NSW, where it carries over from season to season on infected chickpea volunteers, lucerne, native medics and as resistant structures (oospores) in the soil.  The importance of alternative hosts in chickpea PRR was strikingly demonstrated at Moree in 2012.  The paddock had never grown chickpeas and had only had three crops (all wheat) after at least 30 years of Coolatai grass and native medic.  PRR was first diagnosed on 5 July 12 in a few groups of plants; on 2 August, the disease was evident throughout the paddock having killed large areas and on 31 August, the entire crop was sprayed out.  In contrast, no PRR was found in the adjoining paddock which had a long history of cropping (and thus opportunities to control medics).  Indeed every 2012 chickpea crop in which PRR was found also had medics growing in the crop.

PRR and waterlogging are favoured by wetter than normal seasons, as occurred in 2010, or by periods of soil saturation in normal seasons as happened in the early part of 2012.  Waterlogging can be confused with PRR, but differs in that (i) roots die from lack of oxygen whilst with PRR the Phytophthora organism consumes them; (ii) plants are most susceptible during flowering and early pod fill whereas PRR can affect plants of any age (iii) symptoms develop within 2 days of flooding compared to at least 7 days for PRR, (iv) roots are not rotted immediately after the waterlogging event and, (v) initially, plants are not easily pulled from the soil unlike those affected by PRR.

As there are no in-crop control measures for PRR or waterlogging, a critical management tool is avoidance of high risk paddocks (based on previous experience and paddock history). The other key tool for PRR is varietal selection.  Current commercial varieties differ in their resistance to P. medicaginis, with Yorker having the best resistance (MR), PBA HatTrick, Flipper, Jimbour, Kyabra having a lower level (MR-MS) and PBA Boundary having the least resistance (MS). PBA Boundary should not be grown in paddocks with a history of PRR, lucerne, medics or other hosts.  2012 trial data indicates that hybrid breeding lines, generated by crossing chickpea (Cicer arietinum, Jimbour or Howzat) with a wild Cicer species, have significantly higher levels of resistance to P. medicaginis than the most resistant variety, Yorker (Table 1). Although the yields of the hybrids in the absence of Phytophthora infection in this trial were slightly lower than those of PBA HatTrick and PBA Boundary, their improved Phytophthora resistance will compensate for that lower yield in wet years. The results also confirm that PBA Boundary  suffers a higher yield loss from PRR than PBA HatTrick and is there more susceptible. The trial was inoculated with Phytophthora medicaginis, and some plots were soil-drenched with a fungicide to stop root infection; yield loss calculations for each variety/line were based on the difference in yield between the fungicide-treated plots and untreated plots.

Table 1: Yields of commercial chickpea varieties and breeding lines in the absence of phytophthora root rot, and % yield losses due to phytophthora root rot from a 2012 trial at Warwick QLD

^A D lines are hybrid crosses between C. arietinum and a wild Cicer species

For detailed information on control of PRR in chickpea visit: http://www.pulseaus.com.au/pdf/Chickpea%20Phytophthor%20Root%20Rot%20Management.pdf

Viruses in 2012 chickpea crops

Chickpea crops are susceptible to many plant viruses. The effects on plants include stunting, reddening, chlorosis, distortion, shoot tip wilting, reduced yield and grain quality and often death. Infections occurring early in the cropping cycle generally result in more severe disease outbreaks and yield losses. All are spread by flying insect vectors and almost all can be separated into two main groups, those that are transmitted by aphids persistently and those that are transmitted non-persistently. Persistently transmitted viruses (eg Beet western yellows virus – BWYV, Bean leaf roll virus – BLRV) include the luteoviruses and poleroviruses where the aphids can retain and transmit the viruses for many weeks but require up to 1-2hrs of feeding to transmit. Non-persistently transmitted viruses (eg Alfalfa mosaic virus – AMV, Cucumber mosaic virus – CMV) are only carried by aphids for a few hours but can be transmitted in less than a minute of feeding. Virus disease outbreaks in chickpeas are sporadic and difficult to predict from season to season or between locations. Major outbreaks of virus diseases in chickpeas occurred in the early 1990s (when losses in many chickpea crops on the Liverpool Plains reached 100%) and most recently in 2012 in several regions of NSW.

In 2012, virus infection was found in almost all chickpea crops inspected in NSW from the QLD border in the north, west to Nyngan, south to Dubbo and east to Yallaroi and Tamworth.  The incidence of virus infection was generally about 10% but was as high as 40-60% in several crops with severe outbreaks found from Breeza in the south, west to Burren Junction and north to Edgeroi. Overall, the most prevalent virus was BWYV and in some locations more than 90% of symptomatic plants were infected with BWYV. There are related virus species that also react with the BWYV assay as is discussed further below, so it is possible there was a mix of BWYV-like viruses present. BWYV has a very wide host range and is transmitted by several aphid species. Other viruses also detected in chickpea crops generally at much lower incidences included the non-persistently transmitted Soybean dwarf virus (SbDV), AMV and CMV.  In three crops in the south eastern region (Tamworth, Winton and Blackville) more than 50% of symptomatic plants were infected with AMV – all three crops were close to lucerne stands.  A detailed survey of chickpea crops on the Liverpool Plains found:

• Very high BWYV infection with some AMV and CMV.
• Non-symptomatic plants still showed high BWYV infection (but little or no AMV or CMV).
• The massive BWYV infection in 2012 related to large areas canola and (probably) drought-induced aphid movement. 

There were anecdotal reports of very high numbers of aphids migrating (aphid flights) in early September 2012, preceding the major virus outbreaks in chickpeas. It is possible that the high incidence of BWYV detected in some canola crops and turnip weed may have played a role in the movement of this virus into chickpea crops.

BWYV was detected in weed species from across a large region of NSW and was found in Marshmallow (Malva parviflora) from many locations across NSW including Wagga Wagga, Griffith, Hillston, Burren Junction, Narrabri, Gunnedah and Moree. Other host species included Shepherd’s purse (Capsella bursa-pastoris) and Turnip weed (Rapistrum rugosum).

A newly identified polerovirus (aphid transmitted) was identified from NSW and QLD chickpea crops in 2011 and 2012 from locations near Kingsthorpe, North Star, Burren Junction, Edgeroi and Breeza. It has also commonly been found infecting the weed Phasey bean (Macroptilium lathyroides) from QLD and is hereafter referred to as Phasey bean virus (PhBV) but further characterization is required for a more appropriate name. This virus is distinct from BWYV but appears to cause very similar disease symptoms in chickpea and some of the currently used antibody-based diagnostic tests can not distinguish the two viruses. Under experimental conditions, PhBV was transmitted into chickpeas and other pulse crops using the cowpea aphid (Aphis craccivora). Although the relative importance of PhBV in chickpea crops is still uncertain, it appears to have been responsible for approximately 30% of the infections thought to be BWYV. These results and a recent report of a closely related polerovirus from Tasmania suggest that it has a very wide geographical range. It may be either endemic or have been present in eastern Australia for some time and may be commonly infecting leguminous species including pulse crops throughout these areas. Some aspects of the biology of this new chickpea infecting virus remain uncertain, including the alternative weed hosts that may act as a source for the virus around pulse crops in NSW and the range of aphid species that are causing transmission into crops.

Visit http://www.pulseaus.com.au/pdf/Virus%20Contol%20in%20Chickpea.pdf for detailed information on reducing losses from viruses in chickpeas.   Currently, the best strategies to manage chickpea viruses are agronomic ones:

• Retain standing stubble which can deter migrant aphids from landing. Where possible, use precision agriculture to plant between stubble rows. This favours a uniform canopy which makes the crop less attractive to aphids. 
• Plant on time and at the optimal seeding rate – these result in early canopy closure which reduces aphid attraction (see paper by Verrell in these proceedings)
• Ensure adequate plant nutrition (see paper by Verrell in these proceedings)
• Control in-crop, fence-line and fallow weeds – this removes in-crop and nearby sources of vectors and virus. 
• Avoid planting adjacent to lucerne stands – lucerne is a perennial host on which legume aphids and viruses, especially AMV and BLRV survive and increase.
• Seed treatment with insecticides e.g. imidacloprid are not effective for non-persistently transmitted viruses but may be effective for luteoviruses. Unfortunately, local data supporting seed treatment is lacking.
• Given the high incidence of BWYV sometimes found in canola, consider growing chickpeas (and other pulse crops) away from canola.

Salinity

A 2001 survey found that 48,000ha of agricultural land in QLD was seriously affected by salinity, mostly in the coastal or sub coastal areas. Biggs and Power (2003) stated that 9,500 ha was affected by salinity in the QLD Murray Darling basin alone. The National Land and Water Resources audit estimated that over 3 million ha in QLD could be affected by salinity by 2050. In 2012 some chickpea crops in southern QLD and northern NSW, particularly those growing in the brigalow belt or near rivers, were seriously affected by salinity – in the worse cases, large patches or entire crops died in September/ October. Chickpeas are highly intolerant of salinity (Table 2), with symptoms including plant stunting due to poor root growth and nutrient uptake (especially calcium and magnesium), yellowing and death of leaf margins followed by the entire leaf; yellowing and sometimes reddening of stems and reduced flower production and pod setting.  It is believed wet conditions in 2010 and 2011 caused the water table to rise carrying salts with it.   Soil tests in one 2012 salt affected crop of PBA HatTrick south of Walgett showed chloride levels of 1,770 mg/kg (1:5 H2O) and EC_se of 9.8 dS/m at 15-60cm.

Table 2: Relative salt tolerance of some field crops^A

^A Adapted from Table 46 (pages 125-126) in The Salinity Management Handbook, 2^nd edition. Department of Environment and Resource Management, Brisbane, 2011, and from Maas and Hoffman (1977).

^B Electrical conductivity of saturated extract; dS/m = decisiemens per metre

^C productivity decrease per unit increase in salinity is not linear.

Salts occur naturally from weathering of rocks and from the atmosphere, and there is a balance between inputs and losses of salts in natural systems. Changes in the water balance from land clearing, cropping and irrigation alters that balance, causing excessive deep drainage and salt mobilisation. High salinity is caused by shallow water tables and an increase in the amount or concentration of salts in the upper soil layers.  The groundwater does not need to be highly saline for salinity to occur, because when slightly saline water is close to soil surface, evaporation concentrates the salts. Dryland salinity is mostly caused after clearing of trees and other deep rotting vegetation when slightly saline groundwater rises towards the surface. Salinity can also develop as a result of poor irrigation practices, because it has the capacity to mobilise salts at a greater speed due to more water in the subsoil. Sodicity, the result of high sodium levels in the soil is often confused with salinity, but both can occur together particularly in brigalow soils.

Options for management of dryland salinity are limited and include – planting, regenerating and maintaining native vegetation and good ground cover in recharge, transmission and discharge zones, increasing groundwater use in recharge areas by pumping water from bore, installing subsurface drainage, and growing salt-tolerant crops in problem paddocks.

Ascochyta management

The impact of Ascochyta blight (AB; caused by Ascochyta rabiei) in 2012 was minimal across northern NSW and Southern QLD.  Of 363 chickpea crop inspections conducted in northern NSW between June and November, AB was confirmed in only 10 crops with only one of these north of Coonamble (cv Flipper at North Star).  Six crops were PBA HatTrick.  AB was found for the first time in a commercial crop of PBA Boundary at Narromine in mid-September demonstrating that whilst this variety has the best AB resistance currently available in desi varieties for the Northern Region, it still requires management.  The reasons for the small number of AB infected crops in northern NSW and Southern QLD in 2012 were (i) lower than normal rainfall during the season, and (ii) replacement of susceptible varieties with more resistant ones.  However, in July, outbreaks of AB occurred across many parts of central Highlands of QLD, including localities where the disease had not been detected previously. Conditions were ideal for AB, with cooler than normal weather and three or four rain events, each of 3-6 days duration during late May to mid July. Subsequently, the lack of rainfall in the months after these initial outbreaks minimised the impact. These outbreaks confirmed that AB must be considered to be a potentially significant disease in central QLD in some years, which warrants the adoption by growers and industry of integrated disease management practices. These practices include – use of clean, fungicide-treated planting seed sourced from central QLD and avoidance of growing chickpea in or beside paddocks in which AB has been detected in the past 2 seasons.  All of the current commercial varieties adapted to central QLD (PBA Pistol, Kyabra, PBA Moti, and Jimbour) are highly susceptible to A. rabiei, and our future recommended fungicide strategies for these varieties will be the same as for susceptible varieties in southern QLD and northern NSW.

In northern NSW and southern QLD, initial Ascochyta inoculum for the 2013 season will be quite low and the greatest risk of this disease to chickpea growers will be complacency.  If you are growing susceptible varieties eg Jimbour, Kyabra, the first Ascochyta foliar fungicide is critical – don’t miss it.  All 2013 chickpea growers are urged to follow the disease recommendations in the Varietal Management Packages, VMPS for each variety and seek other information from the following sites:

http://www.pulseaus.com.au/pdf/Chickpea%20Ascochyta%20Blight%20Management.pdf

http://www.pulseaus.com.au/pdf/Chickpea%20Integrated%20Disease%20Management.pdf

http://www.dpi.nsw.gov.au/__data/assets/pdf_file/0011/272945/Winter-crop-variety-sowing-guide-2013.pdf

Chickpea seed discoloration

From October 2012 onwards, reports of higher than normal levels of discolored seed began to come from central QLD crops prior to harvest. Later, the disorder was reported from southern QLD and northern NSW. Fifty (50) samples of seed harvested from commercial crops showed that of the 23 samples from central QLD, 65% had discolored seed, and in southern QLD 82% of the 27 samples had discolored seed. The % of affected seed ranged from 0% to 8.6% with an average in affected samples in central QLD of 1.6% and in southern QLD of 0.8%. The varieties PBA HatTrick, PBA Boundary and Kyabra generally had lower levels of discolored seed than other varieties. The last report of significant seed discoloration in chickpea crops in northern NSW and southern QLD was 2003.

Affected seeds display a range of symptoms, including black speckling consisting of small spots, “tiger stripes” consisting of short stripes usually in the groove between the lobes, to large discoloured areas (up to 30%) over the seed coat. The discoloration is confined to the seed coat, and buyers in northern NSW, and southern and central QLD did not reject deliveries with discolored seed.

Work on a similar problem in Western Australia in 1999 showed that there were varietal differences, with cv. Tyson being resistant and cvv. Sona and Heera being susceptible (Vietch 2001). Data from CQ/SQ variety trials in 2012 indicate that there are differences in the incidence of seed discoloration between varieties and breeding lines and that there are advanced breeding lines with high tolerance to the disorder.

The available evidence indicates that the seed discoloration is not due to a pathogen, rather it is a result of an environmental factors. No pathogenic fungi or bacteria were isolated from discolored seed, nor were there symptoms of virus infection on plants with discolored seed. Examination of chickpea plants showed that pods with discolored seeds occurred on some, but not all of the branches and in most cases there were only one or two affected pods on a branch.   On an individual plant, affected pods were not necessarily at the same height above ground level, but affected pods were often at the same height on different plants. It is apparent from these observations that discoloration of chickpea seed is a result of the influence of an environmental stress or combination of stresses at a specific stage of seed development. The most likely culprits are high temperatures perhaps in combination with low soil moisture levels, conditions which were widespread in the 2012 season.

Management –As the most probable cause of chickpea seed discoloration is a genotype x environment x seed development interaction, and the fact that significant levels of discoloration are infrequent and almost impossible to predict, little can be done to prevent the disorder.

Glyphosate and seed germination

Desiccating chickpea crops too early can result in a significant reduction in germination of seed kept for planting. Desiccation is a valuable tool for maximising yield and quality where the crop is uneven in maturity due to late season rain, low plant populations, poor heliothis management, wheel tracks, weeds etc. As chickpea seeds mature progressively from the bottom to the top of the plant, deciding when to desiccate in these situations can be difficult. Experience has shown that the optimum time to apply a registered desiccation product is when 90-95% of seeds are physiologically mature (i.e. less than 35% moisture content). The best rule of thumbs is – desiccate when all seeds in the top 25% of the canopy are starting to turn yellow from the “beak” and when all of the seeds in the bottom 75% of the canopy are fully yellow and the pods are a very light green-yellow colour. The withholding period for Roundup® PowerMax from application to harvest is 7 days.

References

Biggs, AJW, Power RE (2003). A review of salinity occurrences in the Queensland Murray-Darling Basin, 2002. Department of Natural Resources and Mines, Toowoomba.

Brough DM ( 2007) Salinity. Pp. 94-98 in State of the Environment Queensland 2007. Department of Environment and Heritage Protection, Queensland.

 Irvine SA, Doughton, JA 2001. Salinity and Sodicity, implications for farmers in central Queensland. In Proceedings of the 10^th Australian Agronomy Conference.

Pulse Australia. Chickpea harvest and seed storage, including desiccation timing guidelines.

Veitch C. (2001). Crop updates – desi chickpea seed discoloration. Western Australia Department of Agriculture. www.agric.wa.gov.au/PC_90973.html

Verrell, A. (2013).  Virus in chickpea in northern NSW 2012 – the effects of sowing date, plant density and nutrition on virus symptoms in chickpea.  Proceedings 2013 GRDC Grains Research Update, Goondiwindi.

Acknowledgements

Thanks to the growers and agronomists for help with the crop inspections and submitting specimens, to Woods Grains, Goondiwindi for planting material for trials and to chemical companies who provided products for research purposes and trial management.  Thanks to Steve Harden for trials designs and to Paul Nash, Gail Chiplin, Willy Martin and Kris King for technical support.

Contact details

Kevin Moore
NSW Department Primary Industries
Ph: 0488 251 866
Fx: 02 6763 1100
Email:  kevin.moore@dpi.nsw.gov.au

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