The Beat Sheet

White heads and stem borer in wheat

2009-10-12T08:12:00.005+10:00

Every year we receive reports of white heads in wheat, and while there are several possible causes of this symptom, one suspect implicated in the crime is a small stem boring larva called Ephysteris silignitis (Turner) belonging to the moth Family Gelechiidae.

Rod Collins and Hugh Brier did some investigative work back in 1998. They reared a couple of larvae through to the adult moths and had them identified by ANIC.

Rod Collins made the following observations: “The damage was usually confined to a single tiller per plant at a relatively low incidence through fields. Infected tillers seemed to have flowered normally, but soon after flowering the stem upwards from the last node (and including the head) died and was white in colour with no grain in the head. From a distance, these symptoms appeared to be the same as those of crown rot. However, infected tillers were green and apparently healthy from the last node (including the flag leaf) down. On closer examination, a small entry hole about the size of a pinhead was evident usually at or just below the first node up from the base of the plant. In some cases an exit hole was noted just above the last node.”

“When the stem was split open, you could follow where the larva had been up until the last node, where it was often found feeding on the tip of the stem just above the last node. In some cases, the larva had chewed through the tip and continued to move upwards towards the head. It appeared that once the stem began to dry out, the larva would bore a hole in the stem and exit. Only one larva was found per stem in all the plants that I saw.”

It seems not much is known about this species. It is believed to be a native species, one of three in this genus found in Australia. Ephysteris promptella is recorded as a pest of sugarcane in Australia. Ephysteris silignitis occurs widely in Australia south to about 35 degrees south and is thought to be confined to Australia. It is in the wettest parts e.g. Brisbane and Mt Bellenden Ker and the driest. It is common at Alice Springs. It was suggested that it may feed on grasses but there was no evidence.

Stem borer larva in wheat (Photo by Iain Macpherson)

At this stage the reports of isolated ‘white heads’ do not represent economic loss, but this stem borer is something to be aware of if those scattered white heads are observed in fields. There is no registered chemical control.

Recce for armyworm in winter cereals

2009-10-09T08:13:00.005+10:00

The quick finish for winter cereals this season has resulted in the majority of crops escaping infestations of armyworm. Headers are already into some fields, but there are reports of armyworm making their presence felt in some of the later crops.

Being aware of their presence is one thing; whether to intervene is another.

In some late crops starting to turn, the presence of up to 12 small armyworm larvae per square metre need not necessarily sound alarm bells. This situation requires careful and regular monitoring, but there is every chance the crop will make it to harvest without the need to control armyworm.

If however, the crop lingers and the armyworm develop into medium and large larvae, there is a risk, particularly with barley, that head cutting will result in high yield losses. In this situation, quick action may be required to control armyworm and prevent losses.

The key points are
1) to be aware that armyworm are present and
2) to inspect regularly as the crop approaches maturity so that appropriate action can be taken if head cutting occurs.

For more information on armyworm, see the posting made on 20 October 2008.

Photo: Watch for the early signs of head cutting.

Reminder: Last date for Steward® EC use on chickpeas is 15 October

Winter pulses have had their expected share of helicoverpa infestations over recent weeks and most crops have been sprayed to control grubs. Strategies to minimise the risk of insecticide resistance are available. The following points should be observed.

Under the Insecticide Resistance Management Strategy (IRMS), the last use of Steward® EC for Central and Southern regions is 15 October, while the last use date for Northern (Central Queensland) regions (15 September) has long passed.

Grower and consultants are also reminded that for all pulse crops, not more than one application of Steward® EC per field is allowed for the crops entire growth cycle.

Access the full IRMS for 2009-10 on the Cotton CRC website:
http://www.cottoncrc.org.au/content/Industry/Publications/Pests_and_Beneficials/Insect_Resistance_Management.aspx

Resistance Update on the Road

2009-09-10T08:16:00.009+10:00

The Resistance Roadshow visited regional areas during late August and presented the latest resistance monitoring results for a suite of important pests. Presentations covered resistance to conventional insecticides in cotton aphids, mites, silverleaf whitefly and helicoverpa, and helicoverpa resistance to the Bt toxins in Bollgard II.

Photo: Downs agronomist Bernie Caffery (right) discusses resistance with Sharon Downes and Grant Herron at the Dalby Resistance Roadshow.

Aphids and mites
Dr Grant Herron, Industry and Investment NSW based at Menangle, presented his latest results for cotton aphids and mites. Cotton aphid resistance to neonicotinoids was detected at many locations during the 2008-09 season and was associated with product failures. Grant suggests that neonicotinoid seed dressings are influencing the development of this resistance. His recommendation is to try to avoid foliar neonicotinoids for aphid control if a neonicotinoid seed dressing has previously been used that season. However, if they are sprayed for aphid control Grant highlighted the need to alternate chemical groups for seed dressing and first foliar spray.

Fortunately there appears to be no cross resistance to pirimicarb, organophosphates and endosulfan and these products should perform satisfactorily against neonicotinoid resistant strains. Cotton aphid resistance to Pegasus® was reported for the first time.

Resistance was detected to Comite® in two spotted mite populations. This is one of the most widely used products for mite control in cotton, so the detection of resistance is a serious concern.

Whitefly
Results of resistance testing for silverleaf whitefly (B biotype) by Zara Ludgate, QPIF Toowoomba, indicate no immediate concerns for the two key products, Admiral® and Pegasus®, used to manage this pest in cotton. Surveys of cropping regions during autumn and winter 2009 have so far failed to reveal any Q biotype Bemisia tabaci that was first reported from north Queensland in late 2008 and north-western NSW in 2009.

Helicoverpa
According to Dr Louise Rossiter, Industry and Investment NSW, Narrabri, resistance in Helicoverpa armigera to most conventional insecticides has declined or stabilised. Areas of concern are the continuing high level of resistance to older pyrethroids and moderate resistance to the carbamates. Field performance of these products against H. armigera may be highly variable. There are no conventional insecticide resistance issues associated with H. punctigera.

The 2009-10 Insecticide Resistance Management Strategy is available on the CRC website:
http://www.cottoncrc.org.au/content/Industry/Publications/Pests_and_Beneficials/Insect_Resistance_Management.aspx

Bt resistance
Dr Sharon Downes, CSIRO Narrabri, and Kristen Knight, Monsanto Australia, outlined the results for resistance testing to the two Bt toxins. While the frequency of Cry 1Ac resistance alleles remains at very low levels, the frequency of Cry 2Ab resistance alleles in populations of H. armigera and H. punctigera are higher than expected. Changes in the frequency of resistance alleles are being closely monitored as further upward movement in resistance frequencies could be a trigger for changes to the Resistance Management Plan which aims to preserve the usefulness of this GM technology.

Photo: Kristen Knight (left) of Monsanto discusses Bt resistance with Kate Charleston of QPIF at the Dalby Resistance Roadshow.

Refresher on managing helicoverpa in chickpea

2009-08-07T12:44:00.013+10:00

With the approach of spring, helicoverpa start to become active. In CQ, chickpea crops are attractive to moths, and it is timely to revisit some of the key points related to making decisions about control of this pest in crops.

There is a technical brochure available at the QPIF website, which provides a comprehensive overview of sampling and the use of economic thresholds to guide decision making. The brochure can be accessed via this link: (http://www.dpi.qld.gov.au/cps/rde/dpi/hs.xsl/26_6821_ENA_HTML.htm).

In this article, I wanted to discuss a couple of the recommendations which some growers and agronomists have found a bit challenging. The first is adjusting larval density estimates to focus on those larvae that will cause damage.

1) Excluding very small larvae from threshold calculations. Estimating the number of very small larvae (VS) is time consuming to do in the field, and the accuracy of the estimate can be affected by the age and condition of the crop (figure 1). Whilst VS larvae are indicative of the size of the potentially damaging population in 2-3 weeks time, they are not relevant to a decision about controlling the current population to prevent yield loss.

Figure 1. Comparison of visual, beat sheet and absolute sampling for helicoverpa larvae on (a) dryland chickpea at late flower - pod fill, and (b) irrigated chickpea at mid flower to early pod set. VS=very small, S=small, SM=small-medium, ML=medium-large, L=large larvae. (Source: Melina Miles, 2004)

2) Applying a 30% mortality factor to small larvae. Essentially, applying mortality is acknowledging that a proportion of the population will not cause any yield loss because they do not survive to become medium and large larvae. Natural mortality is likely a result of dislodgement from the plant, disease, cannibalism, and predation. Remember that 80-90% of crop damage is caused by medium and large larvae.

This recommendation is based on trial work, largely in CQ, in which we followed infestations of helicoverpa and determined what proportion of the starting population of small larvae survived to large. The level of survival was variable between fields, but on average we determined that 70% of small larvae died before they reached large. In making a recommendation to include natural larval mortality we have been conservative, using only 30% mortality (the highest level of survival we found at any of the sites we monitored).

In practice, in calculating how many larvae per m2 are likely to contribute to yield loss in the crop if left untreated, the following equation be used:

Where S = small, M= medium and L= large larvae

3) One well timed spray should be enough. The timing of a spray should control larvae before they cause damage, but not simply be applied if larvae in the crop exceed threshold. Infestations of helicoverpa in vegetative and flowering crops have been shown to cause no yield loss in chickpea.

A spray may need to be applied during late flowering if targeting small and medium larvae to prevent them causing damage to a podding crop when they are medium-large. However, treating a population to prevent damage to a vegetative or flowering crop offers no yield benefit.

Trial results show that, in chickpea, helicoverpa larvae cause very little damage to buds and flowers (Figure 2). This is quite different behaviour from that seen in mungbeans, for example, where larvae show a clear preference for buds and flowers. Chickpea leaves appear more palatable to helicoverpa larvae than that of mungbeans.

Figure 2. Feeding preferences of small and large helicoverpa larvae on chickpea.

These observations are supported by yield data from time of spraying trials which shows no significant benefit in yield from applying a spray at flowering. Yield loss starts to occur when larvae are present in podding and filling crops (Figure 3).

Figure 3. The impact of helicoverpa on chickpea yield when controlled at different stages of crop maturity. Bars with the same letter are not significantly different from each other.

Remember
Only one application of Steward® (indoxacarb) per crop, with a cut off for use of 15 September in CQ, and 15 October in cooler regions. Currently there is no evidence of resistance to indoxacarb in helicoverpa populations from cotton, but there has been no testing of larvae from CQ, or from chickpea. I would encourage you to make collections from chickpea to send to Dr Louise Rossiter, NSW DPI, Narrabri. Contact Louise on 02 6799 2428 or louise.rossiter@dpi.nsw.gov.au.

Revised ready-reckoner for calculating the economic threshold

Economic Thresholds (larvae/m2) for Conventional Pesticides

Cross-reference the cost of control versus the crop value to determine the economic threshold (ET).
If the cost of control = $25/ha and the crop value =$450/t, the ET = 2.8
If the cost of control = $10/ha and the crop value =$550/t, the ET = 0.9
The lower the cost of control, and the higher the crop value, the lower the threshold.
(Table compiled by Gordon Cumming, Pulse Australia)

Your opinions and experience are important, so leave a comment on this article or helicoverpa management issues.

Maybe you would like to respond to the following questions:

- How are you calculating your economic thresholds? Are you using the equation suggested by QPIF, or is it close enough to 2 or 3 that you are just using that?

- Is the revised ready-reckoner any easier to use?

- Would a threshold calculator be a useful tool? i.e. where you put sampling and crop data in (larval number and size, grain price, cost of control) and it calculates the threshold.

Article by Melina Miles

Winter cereal aphids – background to the potential impact of infestations

2009-08-02T22:58:00.005+10:00

The most critical issues we face in managing cereal aphids currently is the lack of local knowledge about the likely impact of infestations on yield and quality (the damage thresholds).

In this article, I will not go over aphid basics i.e. identification and sampling. You can follow the links to previous articles to read about these (http://thebeatsheet-ipmnews.blogspot.com/2008/09/cereal-aphids-in-wheat-and-barley.html).

In this posting, I want to discuss what is known, from overseas research, and what we might draw on from this work to help us make decisions about aphid management and control. This review may provide some useful information, in the absence of any locally generated data on aphid impacts. Surprisingly, there has been very little work done on cereal aphids in Australia.

General points
The literature, largely from North America and Europe, indicates that there can be significant differences in the way different cultivars respond to the impact of aphids. For this reason, it is important to use this information as general information that may assist in understanding how your crop may be responding to an aphid infestation. In the absence of local data, it is a useful starting point.

Aphids have a requirement for nitrogen (N) to complete development and reproduce. Honeydew is a by-product of their feeding. Essentially aphids compete with the plant for available N, which can impact on the plant in at different stages of crop development.

Early aphid infestations (from seedling)
Root and shoot growth may be impaired as a result of aphids competing for N. Inadequate N for the crop may make the crop more vulnerable to the impact of an aphid infestation.

Prolonged infestation can reduce tillering and result in earlier leaf senescence. Controlling aphids generally results in a recovery of the rate of root and shoot development, but there can be a delay.

Late aphid infestations
There is no impact on yield after grain has filled and is maturing (soft-hard dough).

Infestations that occur during booting to milky dough, particularly where aphids are colonising the flag leaf, stem and ear, result in yield loss. Generally, the distal grains in the head fail to fill. Infestations at this stage in which aphids colonise the leaves, particularly lower in the canopy, tend to result in grain with reduced N (protein) rather than a loss in yield. Aphids are intercepting the N being relocated from leaves to the filling grain.

The relative impact of timing and location of infestation makes sense if you review it along with what is known about the contribution of different parts of the crop to yield. The figure below illustrates the contribution of the upper leaves, stem and ear to the yield of wheat and barley (GRDC Winter Cereal Crop Growth Guide 2005 http://www.grdc.com.au/director/events/grdcpublications?item_id=8D607A46EDDFD98A822CFAEC7FCC4EC2&pageNumber=1).

Ongoing research
There is currently research being conducted on cereal aphids, by QPIF and the Northern Grower Alliance (NGA).

In 2008, initial trial work by QPIF and NGA showed different results (see the GRDC Update, Goondiwindi, 2009 papers for NGA results. Briefly, NGA trial work showed an overall yield benefit of around 10% from using seed dressings containing imidacloprid. QPIF results showed no difference from seed treatment, but a yield benefit where a foliar treatment (pirimicarb) was applied at head emergence.

Article by Melina Miles

Slaters and other winter cereal establishment pests

2009-05-28T14:18:00.012+10:00

In recent days we have received a number of reports of slater activity in winter cereal crops in southern Queensland and northern NSW.

Slaters are not generally regarded as a pest of broad acre agriculture and tend to feed on decaying vegetation and dead animal matter. Overall they perform an important recycling role in the environment however on rare occasions they can also attack seedlings of broad acre crops.

Slaters are woodlice and they are crustaceans, not insects. They have a hard skeleton on the outside and many pairs of jointed legs. The native slater species doing the damage to cereal crops is Australiodillo bifrons. This species has a light brown oval shaped and flattened body with a dark brown stripe in the middle of the back. Both males and females have a characteristic split on the frontal plate. Males tend to be larger than females and can grow as large as 9 mm long and 6.5 mm wide.

This native slater is commonly found in low lying swampy regions and tends to be more active after rain periods. They need damp conditions and will die if exposed to open and dry situations.

Slaters are an agricultural pest in South Africa where they are generally controlled by cultivation. Changing farming practices such as minimum or non tillage seem to have worsened the slater problem, especially if there is also a large amount of stubble present in fields.

Slaters are known to do damage to seedlings of wheat and oats and there is also evidence of slater activity in canola in western and southern Australia. It is not known if other crops are hosts for slaters.

Damage: Slater damage looks similar to snail and slug damage with rasping and shredded appearance to leaves. Feeding damage can also appear as irregular patches removed from the leaves resulting in distinctive ‘windows’ of transparent leaf membrane. Thousands of seedlings can be eaten in a short time by swarms of slaters.

Control: There are no registered pesticides for the control of slaters in winter cereals. Non chemical approaches such as providing alternative habitats may decrease slater numbers in crops. Shelterbelts containing a complex understorey of vegetation and soil litter may be more attractive to slaters. Such environments also harbour many natural enemies of broad acre insect pests which can also keep slater populations in check.

Other winter cereal seedling pests

Cutworms
Several species of cutworms attack establishing cereal crops in Queensland and NSW. As their name suggests cutworm larvae sever (cut) the stems of young seedlings at or near ground level, causing the collapse of the plant.

Cutworm larvae are up to 50 mm long, hairless with dark heads and usually dark coloured bodies, often with longitudinal lines and/or dark spots. Larvae curl up and remain still if picked up.

Damage: Young caterpillars climb plants and skeletonise the leaves or eat small holes. The older larvae may also climb to browse or cut off leaves, but commonly cut through stems at ground level and feed on the top growth of felled plants. Caterpillars that are almost fully grown often remain underground and chew into plants at or below ground level.

Monitoring and control: Inspect crops twice weekly in seedling and early vegetative stage. The best time to monitor is late afternoons and evenings when larvae feed. Chemical control is warranted when there is a rapidly increasing area or proportion of crop damage. If distribution is patchy, spot spraying may suffice. Chlorpyrifos and various pyrethroids are mainly used to control cutworm.
Cultural control measures include weed control – at least 3-4 weeks prior to sowing.

Blue oat mite
The blue oat mite is an important pest of seedling winter cereals. When infestations are severe the leaf tips wither and eventually the seedlings die. Eggs laid in the soil hibernate over winter, allowing populations to build up over a number of years, causing severe damage if crop rotation is not practised.

Adults are 1 mm long and have 8 legs. Adults and nymphs have a purplish-blue, rounded body with red legs. They move quickly when disturbed. The presence of a small red area on the back distinguishes it from the redlegged earth mite.

Damage: Adults and nymphs pierce and suck on leaves resulting in silvering of the leaf tips in cereals. When heavy infestations occur, the leaf tip withers and the seedling can die. In canola, leaves are mottled or whitened in appearance.

Monitoring and control: Check from planting to early vegetative stage, particularly in dry seasons. Blue oat mites are most easily seen in the late afternoon when they begin feeding on the leaves.
Where warranted, foliar application of registered insecticide may be cost-effective if applied within 2-3 weeks of emergence in autumn.

Know your seedling pests
Correct identification of pests feeding on cereal seedlings is important as this will influence selection of control options.

Article by Kate Charleston and David Murray

Fleabane alert
Following the recent rain, the first main flush of fleabane for the year will start to emerge shortly. Whilst this weed is often regarded as one of the most difficult-to-control weed, it is much easier to control when it is a small seedling. So, growers need to be alert and think about spraying soon.

To assist growers and consultants, the weeds team has recently published a brochure on fleabane, and it is available from the Queensland Primary Industries & Fisheries website http://www.dpi.qld.gov.au/cps/rde/dpi/hs.xsl/26_4251_ENA_HTML.htm

The team is also developing a best practice herbicide guide, using feedback from consultants on what is being used successfully for in-crop and fallow control across the region.

If you wish to be part of this short survey, please contact Michael Widderick on Michael.widderick@dpi.qld.gov.au or Steve Walker on steve.walker@dpi.qle.gov.au

New whitefly found!

2009-04-28T14:17:00.006+10:00

Detection of Q biotype Bemisia tabaci in Australia

The presence of Q biotype Bemisia tabaci species complex in Australia has been confirmed by Dr Robin Gunning, NSW DPI. Q biotype was collected from vegetables in the Bowen/Burdekin region during late 2008, as well as from cotton in southern Queensland (Goondiwindi) and north-western NSW (Wee Waa) during 2009. It is likely that Q biotype is more widely distributed than just these regions.

What are the implications?
Overseas studies indicate Q biotype has the capacity to develop resistance to many insecticides including insect growth regulators (IGRs) such as Admiral® and neonicotinoids like Confidor®. High levels of resistance to Admiral® have been detected in horticultural crops in a few locations in north Queensland and some field control problems have been observed for Admiral®.

Resistance testing from cotton production areas for the 2008-09 season has not shown any alarming resistance levels to Admiral® to date.

Overseas where populations are predominately of Q biotype, moderate to high resistance has developed to Admiral®. Where populations were mostly B biotype, Admiral® has retained high efficacy. This has been the case in Queensland where, according to Dr Gunning, B biotype populations remain susceptible to Admiral® and have a higher susceptibility to neonicotinoid insecticides, compared to Q biotype populations. At this stage Q biotype is showing markedly less resistance to pyrethroids than the B biotype.

In Israel, Q biotype has not developed resistance to Pegasus® despite several years of reliance on this product. In horticultural areas, significant resistance to Pegasus® was not found in either biotype.

Integrated Pest Management
Practicing good IPM principles can discourage Q biotype numbers from building up. Under natural conditions, B biotype will out-compete Q biotype. However, in an environment of high insecticide use, the more insecticide resistant Q biotype tends to displace B biotype, and once this shift occurs B may not recover to its former levels. Limiting the amount of chemical used against insect pests may favour the dominance of B over Q.

Q biotype, like B, has the capacity to vector the virus that causes cotton leaf curl disease. This disease is not present in Australia. The main risk is that any new whitefly incursions, whether Q or B biotype, could carry viruses that are not present in Australia.

Identification
Q biotype and B biotype can not be distinguished visually. They can only be distinguished by looking at small differences in their DNA or biochemical make-up.

WE NEED YOUR HELP
In order to determine the distribution of Q biotype, we are asking growers and consultants to send in whitefly specimens to the Queensland Primary Industries and Fisheries, Toowoomba. Please refer to details at the bottom of this article.

FURTHER READING

Whiteflies http://www.dpi.qld.gov.au/cps/rde/dpi/hs.xsl/26_10277_ENA_HTML.htm

The Cotton Industry Biosecurity Plan Appendix 3 provides information on Q biotype (page 32) and Cotton Leaf Curl Virus (page 40).
http://www.planthealthaustralia.com.au/project_documents/uploads/Section%209%20Appendix%203%20Pest%20Risk%20Reviews.pdf

Follow this link to the Fact Sheet on Q biotype whitefly. http://www.dpi.qld.gov.au/cps/rde/dpi/hs.xsl/26_13554_ENA_HTML.htm

WHITEFLY SAMPLING
Adult whiteflies: In fields where whitefly are present, collect a minimum of 30 adults from random plants throughout the crop. Place these in 65% alcohol (watered down methylated spirits) in a leak proof vial or bottle. Note that >70% alcohol is classified as a dangerous liquid and should not be sent via post or courier.

Immatures: In fields where whitefly are present, collect a minimum of 30 leaves from random plants throughout the crop. Aim to collect leaves that have large immature scales (4th instar/red-eye nymphs) on their underside. Collect only 1 leaf/plant. Pack the leaves into a paper bag and then inside a plastic bag.

For live material, send by overnight courier to:
Richard Lloyd
DEEDI, Primary Industries and Fisheries
203 Tor St, Toowoomba Q 4350
Ph: (07) 4688 1315

Ensure samples are clearly labeled and include the following information:
Collectors Name, Phone No., Fax No., Email address
Farm Name, Field, Postcode, Region (e.g. Gwydir)
Date of Collection, Host Plant (Crop)
Comments

Article by Zara Ludgate and David Murray

What is eating my soybean pods?

2009-03-06T14:22:00.006+10:00

Field crickets
High numbers of field crickets have been reported across the Darling Downs in the last couple of weeks, with some large aggregations of adults attracted to lights at night. Are these crickets doing damage to crops? The answer may well be yes.

Field crickets are generally a pest of pastures but can also attack soybeans, cotton, sugarcane and sunflowers. There are two species commonly found – the brown field cricket and the slightly larger black field cricket. The brown field cricket is the most prevalent at present. Crickets hide during the day in cracking soils or under clods of dirt or crop residue and emerge at night to feed on crops. They usually feed on seedlings but high populations in late summer may feed on more mature plant structures such as sunflower heads and soybean pods.

How do I know that crickets are damaging my soybean pods?
Crickets are not the only pest of soybean pods. Mice can also do considerable damage to pods, with soybeans often the last of the summer crops to mature and as a consequence the only source of food on offer for mice.

Cricket adults and large nymphs chew into pods to reach the seeds but this damage looks similar to damage done by mice. So how do you determine what pest is doing the damage?

The best time to check for crickets is to inspect crops at dusk or later into the night when crickets are most active. Field cricket activity can also be monitored with light traps. Another way to determine whether crickets are present is by using hessian bags placed out overnight at regular intervals across the paddock. In the morning check for the presence of crickets sheltering under the bags.

The best way to determine whether mice are damaging the crop is to go out at night to check for their presence or use mouse bait cards (as described on the DPI&F website).

Mice damage to soybeans can be an ongoing and costly problem as soybeans provide good groundcover for mice and excessive grain losses immediately prior to or during the harvesting operation are likely to increase mouse populations given an ongoing food source. This will also impact on early follow-on cereal crops such as wheat and barley.

For further information on how to manage mice in your crop please visit the DPI&F website through the link provided below:

http://www.dpi.qld.gov.au/cps/rde/dpi/hs.xsl/4790_8283_ENA_HTML.htm

How do I control the crickets if they are damaging my soybeans?
Past experience has shown that foliar insecticide applications do not provide control of crickets. They shelter by day and are found low down under dense canopy at night, making spray contact difficult.

Chlorpyrifos-treated cracked grain baits are registered for cricket control in soybeans, but this is mainly used to protect seedling crops. The bait is prepared by mixing 100 mL chlorpyrifos (500 g/L EC formulation) and 125 mL sunflower oil, and adding this to 2.5 kg of cracked wheat or sorghum/ha. The bait is broadcast evenly by air or ground.

Article by David Murray and Kate Charleston

Mirids in Mungbeans

2009-02-26T11:10:00.007+10:00

Mirids can cause significant crop losses in mungbeans with yield reductions of up to 25-50% common where high mirid populations (eg 10/m2) are left uncontrolled. Mirids can reduce yields by 60 kg/ha for every mirid/m2 of crop.

Mirids can be present in mungbeans at any stage from seedlings to podding. Budding, flowering and early-podding crops are at greatest risk. Low populations of green mirids are often present in vegetative crops but there is no evidence they cause ‘tipping’ of vegetative terminals or have any impact on yield. Mirid populations usually increase with the onset of budding and peak during late podfilll. Generally around 80% of mirids present in flowering/podding crops are nymphs.

Damage caused by mirids
Mirids are sucking insects and feed by piercing the plant tissue and releasing a chemical that destroys cells in the feeding zone. This causes plant tissue to discolour and die. Mirids prefer to feed on flowers, buds and young pods, causing these to abort (shed).

Severe mirid damage in mungbean results in fewer harvestable pods being set, however similar symptoms can also be caused by thrips, high temperatures and moisture stress. Mirids may also attack more mature pods, damaging the seeds inside without causing shedding. If pod set is greatly reduced and mirid populations are relatively low (e.g. 0.5 - 1.0 /m2) - loss may be due to other factors.

Mungbeans always set many more buds and flowers than they can convert to harvestable pods. Typically (in dryland crops) only 33% of buds/flowers are converted to harvestable pods. This % may be even lower in moisture stressed crops.

Recent Helicoverpa threshold trials show that mungbeans can compensate for considerable early damage (at budding/flowering) of up to 80% bud loss with no yield loss. However this only occurs when there is adequate moisture and no subsequent pest damage and there will be a delay in crop maturity. . The loss of replacement buds through continuous pest pressure will lead to reduced crop yields.

Medium and large mirid nymphs (instars 3-5) are as damaging as adults. Although small mirid nymphs (instars 1-2) are less damaging, they soon develop into larger more damaging pests (within 3-5 days respectively at 300C).

Identification of mirids
Three mirid species attack mungbeans and include green, brown and (less frequently) crop mirids. All these species are equally damaging.

For more information on the identification of these mirid species please refer to the Integrated Pest Management website by clicking onto the link provided below:

http://www.dpi.qld.gov.au/cps/rde/dpi/hs.xsl/26_5085_ENA_HTML.htm

Where do the mirids come from?
Mirids move into summer crops when alternative host plants dry off and they seek a fresh food source. Lucerne and sunflowers can be important sources of mirids in mungbean growing areas.

Influxes of mirid adults often coincide with northwest winds in spring. There is also evidence of long distance migration, possibly from inland areas, associated with weather fronts. This may be the cause of some of the widespread and repeated influxes of mirids sometimes observed in grain and cotton growing regions early in the summer.

Monitoring for mirids
- Mirids are very mobile and in-crop populations can increase rapidly from budding onwards.
- Crops should be inspected weekly during the vegetative stage (to pick the start of budding) and twice weekly from budding onwards until post flowering.
- In row crops, the preferred method is beat sheeting, as this method is the most effective for helicoverpa and pod-sucking bugs.
- Sample 5 one-metre lengths of row (not consecutive) within a 20 m radius, from at least 6 sites throughout a crop.
- Avoid sampling during very windy weather as mirids are easily blown off the sheet.

Action level
Mirid thresholds for budding/flowering mungbeans are 0.3-0.6 per m2 for aerial and ground rig respectively. While mirid thresholds are very low, this is more a reflection of the cheapness of the preferred mirid pesticide – dimethoate ($4/ha + application). The above thresholds are based on the cost of dimethoate. Were indoxacarb to be used, the threshold for an aerial sprayed crop would be 1 mirid/m2.

Note that these thresholds are the break even point (cost of control = value of likely damage) and that action only needs to be taken if populations exceed these levels. However in practice, mirid populations usually increase rapidly from budding onwards and spraying at-threshold populations would be justified. Because of the crop’s ability to compensate for early damage, spraying low mirid populations (i.e. at threshold) at very early budding can be delayed slightly with no risk to yield or harvest maturity. Taking this approach reduces the need for a subsequent mirid spray.

Cultural control
A crop that has a short flowering period reduces the risk of mirid damage. Flowering periods can be shortened by planting on a full moisture profile and by watering crops just before budding. Consider planting crops in at least 50 cm rows (as opposed to broadcast planting) to facilitate easier pest sampling.

Chemical control
· Dimethoate at 500 mL/ha (all summer pulses) or
· Indoxacarb at 400 mL/ha (mungbeans and soybeans only).

Dimethoate is often applied at lower than label rates (eg 200-250 mL/ha). These rates give excellent mirid control but have far less impact on many beneficials. Trail results have shown that the addition of salt (0.5% NaCl) as an adjuvant, improves the effectiveness of dimethoate at lower rates. The amount of salt used (0.5%) has no phytotoxic effect on summer pulse crops. ‘Hard’ water can markedly lower the effectiveness of dimethoate and should be countered by adding a buffering agent such as LI700.

Indoxacarb is not recommended at high mirid pressure (>2/m2) because it is less effective than dimethoate. Because it is restricted to one (1) spray application per crop, indoxacarb is best reserved for Helicoverpa control during podding. As mentioned previously, the mirid thresholds are considerably higher if indoxacarb is used.

Natural enemies
There are no beneficial species that are recognised to be regulators of mirid populations in mungbeans. However damsel bugs, big-eyed bugs, predatory shield bugs, as well as lynx, night stalker and jumping spiders are known to feed on mirid adults, nymphs and eggs. Naturally occurring fungi (eg Beauvaria) may also infect and kill mirids, but are rarely observed in the field.

Article by Kate Charleston and Hugh Brier

Late season pests of pulses and cotton

2009-02-20T14:04:00.006+10:00

Go soft early
Go soft early is a basic integrated pest management (IPM) strategy to avoid using non selective pesticides for as long as possible. This approach encourages a build up of predators and parasites to keep early pests in check and buffer the crop against attack later in the season. This strategy is particularly important in soybeans because of the risk of flaring whitefly. With no registered pesticides against whitefly in soybeans the going soft early approach maximises the chances of whitefly parasitism by the recently released wasp Eretmocerus hayati.

Green Vegetable Bugs (GVB) in soybeans
Intervention with non-selective pesticides may be required during pod-fill to control pod-sucking bugs. The most abundant pod sucking bug in soybeans is the green vegetable bug.

GVB are often found in the crop prior to pod-fill, but at this stage they are not causing economic damage. By delaying insecticide use untill pod-fill reduces the risk of flaring whitefly, because as pod-fill progresses, the leaves become progressively less attractive to this pest.

GVB is primarily a pod feeder with a preference for pods with well-developed seeds. Summer pulses remain at risk until pods are too hard to damage (very close to harvest). Damaging populations are typically highest in late summer crops during late pod-fill (when nymphs have reached or are near adulthood).

While many cultivars can compensate for yield loss caused by moderate bug populations, seed quality is adversely affected, particularly for culinary beans which have very low damage tolerances (a maximum of 2% damaged seed). Bug damaged seeds have increased protein content but a shorter storage life (due to increased rancidity). In soybeans, bug damage also reduces seed oil content.

The only effective registered pesticides for pod-suckers are non-selective and include deltamethrin (Decis) and trichlorfon (Dipteryx). Of these, deltamethrin is the most effective. Note that dimethoate and methomyl are also registered but were found to be ineffective against pod-sucking bugs.

To mitigate the impact of non-selective insecticides, delay spraying podsuckers until early podfill.

Green Vegetable Bugs (GVB) in cotton
Cotton is susceptible to damage from GVB from flowering through until one open boll per metre. Damage symptoms from GVB cannot be distinguished from damage done by mirids which include warty growths and brown staining of lint in developing bolls.

Thresholds for GVB in cotton are 1 adult/metre from flowering through to harvest. For insecticide options please refer to the cotton pest management guide which can be found on the Cotton Catchment Communities CRC website.

http://www.cottoncrc.org.au/content/Industry/Publications

Grass blue butterfly
Large populations of grass blue butterfly have been observed in soybeans this season. The green slug-like caterpillars feed mainly on leaves. They may be confused with the larvae of the hoverfly which is a beneficial insect and often found near aphid colonies on which they feed.

While control is rarely needed, high numbers can damage terminals resulting in plant branching and pods being set closer to the ground. This can indirectly impact on yield as low-set pods are more difficult to harvest. There are no insecticides registered for this pest specifically but anecdotal evidence (from a leading Goondiwindi grower) suggests they are readily controlled with Bt (Dipel).

Pale cotton stainers
Cotton stainers are occasional pests of cotton that feed on developing and mature cotton seed. While previously controlled by broad spectrum insecticides for other pests in cotton, the reduction in chemical use, especially on Bollgard II®, may lead to increased populations which may need to be managed.
For more information about pale cotton stainers click on the link provided below:

http://www.cottoncrc.org.au/content/Industry/Publications/Pests_and_Beneficials/Sucking_Pests.aspx

Article by Kate Charleston and Hugh Brier

Soybean Moth causing major damage in soybeans

2009-02-13T08:16:00.009+10:00

Soybean moth Aproaerema simplexella is a very common pest of soybeans but is usually only present in very low numbers. This season high numbers of moths and caterpillars were found especially in the Wide Bay Burnett region with some fields sustaining extensive damage. Similar leaf miners attack many horticultural crops, but are species other than A. simplexella. Weed hosts of soybean moth include emu foot (Psoralia tenax)

Identification
The soybean moth is a small narrow winged moth, up to 6 mm long. The forewings are dark brown/grey, each with a white bar across them, and pale brown hind wings. In soybean crops with large infestations, moths can be seen flying up from the foliage when disturbed.

Small elongated eggs are laid on both the top and underside of leaves, generally near leaf veins. Larvae of soybean moth are pale green/grey and grow up to 7 mm (in length).

Other caterpillars that mine and web soybean leaves include soybean leafminer Lithocolletis aglaozona, a much smaller and less abundant species, and legume webspinner Omiodes diemenalis, common in coastal regions, but which is much larger (up to 15 mm) and is bright green. Legume webspinners also web leaves together, rather than mining leaves (feeding inside them).

Similar leaf miners also attack many horticultural crops, but are species other than A. simplexella. Weed hosts of soybean moth include emu foot (Psoralia tenax).

Damage
Larvae initially feed inside leaves (i.e. in leaf mines) and emerge after approximately four days to feed externally, folding and webbing leaves together to form a protective shelter. Infested leaves are often crinkled and pulled together in the middle and this together with webbing of leaf is the most obvious symptom of damage. In low numbers the larvae only cause cosmetic damage. While larvae normally feed on leaves only, extremely high populations will also graze on the surface of pods, after they have denuded the crop of leaves.

Soybean moth infestations are favoured by hot, dry weather, with crops under severe moisture stress most at risk. Large populations can cause extensive damage by stripping all leaves from crops and so reduce photosynthesis and grain production.

Monitoring and thresholds
Monitor crops regularly for the early warning signs of rare plague events. Look for numerous small, pale patches (leaf-mining) on the leaves and large numbers of soybean moths in the crop or around lights at night.

The indicative threshold is based on defoliation, with. 33% pre flowering and 15-20% defoliation during early to mid pod-fill.

Control
In most years control is not required but large infestations in the Bundaberg region will need chemical control to prevent total crop loss. Check thoroughly before spraying, as larvae may have already pupated (as black pupae within the webbing) or reached full size (7 mm) and stopped feeding.

There are no specific registrations for the control of soybean moth. However pesticides effective against Helicoverpa (except Helicoverpa virus and Bt), and targeting that pest in soybeans will most likely also give reasonable control of soybean moth.

DPI&F plans to investigate the effectiveness of a number of pesticides registered in soybeans (against other pests,) as part of GRDC-funded pulse IPM research project DAQ00086. The hope is to identify at least a moderately selective pesticide to preserve soybean moth parasites such as Temclucha sp., a small Ichneumonid wasp (8 mm). This species has been observed in very high numbers in some crops infested with soybean moth.

Article by Kate Charleston and Hugh Brier

Silverleaf whitefly update

2009-02-11T15:50:00.004+10:00

There are reports of large infestations of silverleaf whitefly (SLW) from the Narrabri/Moree region. Exponential growth in whitefly numbers coupled with honeydew on leaves indicates that the whitefly are probably SLW and not East Australian native Bemisia or Greenhouse whitefly (GHW).

It is thought that increased host plant availability from a wet winter/spring, warm conditions and a decline in natural enemies due to the use of broad spectrum insecticides has contributed to the high SLW numbers in this more marginal area of occurrence.

DPI&F entomologists will be visiting Narrabri on Thursday, 11 February 2009 and Moree on Friday 13 February 2009 where they will meet with growers and consultants and speak about SLW and the management options that are available.

Cotton fields around St George are at or reaching high densities of SLW. Reports indicate Admiral® has been applied on many fields to suppress SLW populations. Parasitism levels of 50% and 70% were recorded from two fields in the St George area. This should help to keep SLW in check later in the season even if they start to re-infest crops post Admiral® spray.

SLW numbers in Biloela and Theodore are reportedly dropping off. This may be in part due to parasitism levels. Recent testing for insecticide resistance in populations of SLW from Biloela and Theodore show no alarming results for Admiral®. These results were expected due to the minimal use of Admiral® in central Queensland this season.

GHW are in moderate densities in the Norwin region on the Darling Downs. While GHW will produce honeydew it does not normally cause the same problems as SLW which has a wider host range, higher reproductive rate, develops resistance to insecticides rapidly, and is adapted to high temperatures. Where populations are a mix of SLW and GHW, consider treating as if all are SLW.

The report on managing silverleaf whitefly by Richard Sequeira and Tracey Farrell can be accessed through the cotton CRC using the link below: http://www.cottoncrc.org.au/files/5743fee2-f978-4a79-a9d1-9b1800e899cf/Whitefly_Management[1].pdf.
This document outlines sampling, thresholds and management options for SLW. Remember only one spray of Admiral® is allowed per season.

Article by Zara Ludgate

Latest helicoverpa thresholds for mungbeans

2009-01-30T16:10:00.008+10:00

Revised thresholds for helicoverpa in flowering/podding mungbeans are based on a rate of damage of 35 kg/ha per larva per square metre in podding crops. The new thresholds are nearly double the old threshold of 1/m2, and make allowances for variations in control costs and crop value. For a typical scenario with pesticide control (including aerial application) costing $40/ha and an anticipated crop value of $600/t, the new threshold (see chart) is 1.9 larvae/m2.

Helicoverpa threshold table for mungbeans 2008
Based on data from 2006/07 threshold trial
Assumes yield loss of 35kg/ha for every larva/m2. No allowance for larval mortality, but this most likely cancelled out by sampling inefficiency with a beat sheet. Yield loss is probably at the upper end of that likely as the trial showed no yield loss for up to 8 larvae/m2 at flowering. Very high control costs included in table reflect extremely high application costs in coastal crops.

Cross-reference the cost of control versus the crop value to determine the economic threshold (ET).

If the cost of control = $35/ha and the crop value =$450/t, the ET = 2.2

If the cost of control = $25/ha and the crop value =$650/t, the ET = 1.1

The lower the cost of control, and the higher the crop value, the lower the threshold.

Note that the thresholds are at the break even point, where the cost of control = the value of the likely damage, i.e. where the benefit: cost ratio is 1:1, or in other words where there is not net gain if you spray and no net loss if you don’t spray. Hence control is only recommended if the population exceeds the economic threshold, in other words if the benefit:cost (B:C) ratio is greater than 1.

While IPM guidelines traditionally recommended a B:C ratio of 2:1, most growers using the control cost scenario (above $40/ha) are unlikely to tolerate another $40 of damage/ha before taking action. Therefore use the above table as follows: Decide how much extra potential damage (in $/ha) you are willing to accept before taking action. For example if you are only willing to accept another $10 of damage/ha before taking action, and control costs and likely crop values are $40/ha and $600/t respectively, then adjust your control costs up to $50/t, and cross reference with the above crop value to give an action threshold of 2.4 larvae/m2.

While early reproductive damage at flowering may be totally compensated for, significant early damage can delay harvest maturity, and may reduce ‘commercial harvest yield’, i.e. the yield in crops where desiccants are used to dry out green pods lagging behind the main crop of black pods. For this reason, the threshold is conservatively set from flowering to podfill.

Recent data suggest early moderate damage can be totally compensated for with no delay in harvest, in well growing crops with plentiful moisture. In such crops, growers might consider using a helicoverpa NPV product such as VivusMax for low-moderate populations (eg 2/m2) provided they are able to guarantee thorough coverage, include an Aminofeed adjuvant and are targeting small larvae (ideally not greater than 5 mm long).

In view of the recent changes to the Helicoverpa threshold in vegetative soybeans, a provisional threshold of 4-5 larvae/m2 has been set for vegetative mungbeans, in lieu of the old 33% defoliation threshold (which still holds for loopers). This is because helicoverpa are also likely to target the mungbean’s auxiliary buds which are the precursors to floral buds.

The threshold is set lower than the vegetative soybean threshold because mungbean plants are smaller than soybeans. Note that this vegetative mungbean threshold is provisional and has to be verified in replicated field trials.

Helicoverpa and mirids
Recently we received a number of reports of flowering mungbean crops with above threshold mirid populations and low numbers of Helicoverpa. In such instances, dimethoate (250 mL/ha) plus NPV can be mixed with no risk of incompatibility. However it is critical to add a buffer such as LI700 to tank mix water to keep the pH below 7, as both dimethoate and NPV are deactivated in alkaline water (pH >7).

Note that dimethoate is recommended at the lower 250mL/ha rate as this has proven efficacy in DPI&F’s trials and has far less impact on beneficials than the full registered rate of 500mL/ha. Preserving as many beneficials as possible will complement NPV’s impact on helicoverpa larvae and will reduce the risk of subsequent sprays to control this pest..

Article by Hugh Brier

Silverleaf whitefly in cotton – an update

2009-01-23T11:17:00.005+10:00

Silverleaf whitefly (SLW) is a serious pest of cotton. It reduces yield and quality of cotton due to feeding damage and excretion of honey dew. It is a difficult pest to manage due to its ability to rapidly increase in numbers and the development of resistance to many insecticides.

Resistance testing for the 2007-08 season indicated no alarming results for Admiral® (pyriproxyfen) or Pegasus® (diafenthiuron) in cotton areas. The IRMS guidelines for Admiral® require that only 1 spray may be applied per season. Apart from the cost involved, more then one Admiral® spray has been shown to increase the development of resistance.

As part of the management strategy for whitefly it is important to know what species or biotypes are present as these will significantly impact on the management decision that is required. Refer to past beatsheet articles to read more on identifying the different species and biotypes of whitefly.

We are seeing mixed populations of whitefly across Queensland. On the Darling Downs, greenhouse whitefly (GHW) has made up >90% of the population in the Norwin area. A sample from Theodore showed the whitefly population was made up of 70% SLW and 30% GHW.

In Emerald, a limited number of Pegasus® sprays have been applied for SLW. Pegasus® is best used for early season suppression of SLW at low insect densities or as a late season knock down to prevent honey dew contamination of open bolls.

Very few (if any) Admiral® sprays have been applied so far this season. Admiral® may be applied after 1450 day degrees if SLW numbers reach high densities. Remember that only one Admiral® spray may be applied per season to limit the potential for resistance to develop.

An excellent publication has been produced through the Cotton CRC by Richard Sequeira and Tracey Farrell That outlines thresholds and sampling methods for SLW in cotton in more detail (www.cottoncrc.org.au/files/5743fee2-f978-4a79-a9d1-9b1800e899cf/Whitefly_Management.pdf). This should be referred to when making management decisions for this pest.

In Emerald, there were high levels of natural control of whitefly from the parasitic wasps, Eretmocerus hayati and Encarsia formosa. Parasitism levels of 40% and 75% were recorded in two fields in the Emerald Irrigation Area. In a field at Biloela, parasites were so abundant that the tiny wasps were clearly visible walking around on leaves.

It is likely that the high parasitism levels recorded in Emerald and Biloela are due, in part, to very limited mirid sprays earlier in the season. This has avoided flaring SLW and allowed beneficials to multiply and offer a free service to growers and consultants in controlling whitefly.

Whitefly numbers are reportedly building up at St George/Dirranbandi. DPI&F entomologists will be visiting the area next week to collect samples for resistance monitoring and check parasitism levels.

While whitefly are definitely starting to make their presence felt in cotton fields across Queensland, their presence does not necessarily warrant action. Monitor fields often as whitefly can build up exponentially, identify what species/biotypes are present, use the available thresholds and avoid flaring whitefly by minimising the use of disruptive insecticides and maintaining beneficials in the system.

Article by Zara Ludgate

New Helicoverpa thresholds in vegetative soybeans

2009-01-15T11:13:00.008+10:00

The new economic threshold for Helicoverpa in vegetative soybeans is 8 larvae per sqare metre and replaces the old 33% defoliation threshold. The new threshold is based on field trials conducted by John Rogers (formerly with DPI&F at Kingaroy). These field trials show that approximately 7.5 larvae per square metre can be tolerated with no yield loss, but that severe yield losses can occur once this critical population (the inflection point) is exceeded.

The new threshold (8 larvae/m2) is based on the maximum number of larvae that can be tolerated before there is an economic reduction in yield. The closeness of the threshold and the inflection point is a measure of the severity of the yield losses that can occur once this critical population is exceeded.

Previous thresholds were based on the maximum defoliation (33% and widely cited in the scientific literature) that can be tolerated without reducing soybean yield. In John Rogers’ trials, Helicoverpa populations equivalent to the new threshold (8/m2) inflicted significantly less than 33% defoliation. Note that the threshold may be influenced by crop size, with fewer larvae tolerable in very early or very small crops, and more larvae acceptable in larger more vigorous late-vegetative crops.

The reason yield loss occurs below 33% defoliation is because of Helicoverpa’s feeding behaviour - they are not called budworms for nothing. As well as feeding on leaves, they also feed on the soybean plant’s vegetative terminals and auxiliary buds, the latter which are the precursors to floral buds.

Previous vegetative thresholds allowed for vegetative terminal loss (tipping) with 25% terminal loss the cited critical level above which action was required. The new thresholds are below the old terminal-loss guidelines as populations of 8 larvae/m2 destroyed fewer than 25% of terminals in John Rogers’ trials.

The crop’s ability to tolerate 7.5 larvae/m2 during the vegetative stage without yield loss, means that Helicoverpa nucleopolyhedrovirus [NPV] (e.g. VivusMax®) can still be safely used prior to flowering, provided it targets appropriately small larvae (<7 mm long). This is because NPV only has to keep populations below this critical level, rather than achieving ≥90% control that would be required if yield loss commenced as soon as populations exceeded 0/m2.

Immediate intervention with a more robust larvicide may be required against extremely high populations (e.g. > 20/m2). While indoxacarb (Steward®) could be used at this stage, only one application is allowed per field per crop growth cycle, and this product is best saved for later in the season when it is most needed.

Until data to the contrary is available, the 33% defoliation vegetative threshold is still valid for
loopers and cluster caterpillars which are primarily foliage rather than bud feeders. However, cluster caterpillars are more likely to attack soybean pods than loopers, but not as savagely as Helicoverpa.

John Rogers’ studies illustrate the link between a pest’s feeding behaviour and its impact on crop yield. The studies also highlight the importance of having ‘species specific’ data, and that a ‘one threshold model fits all’ approach is not always appropriate. Further trials are planned to study the feeding behaviour and damage potential of cluster caterpillars and all the major looper species attacking soybeans. However, such detailed research is likely to take at least 3-4 years to complete.

Helicoverpa damage in soybeans

A- vegetative damage
B - damage to terminals results in
C - reduction in pods and yield

Article by Hugh Brier (DPI&F Kingaroy), John Rogers (formerly DPI&F Kingaroy and Kate Charleston (DPI&F Toowoomba)

Insurance spray for mirids in Bollgard II

2008-12-19T11:32:00.004+10:00

With the Festive Season almost here and an irrigation pending, is there value in applying a spray for mirids for peace of mind? This decision to apply an insurance spray needs to be carefully considered because it has the potential to cost much more in the long run.

In order to make this decision, we need to understand the population dynamics of mirids, their damage potential and what is currently happening to the crop.

Where do mirids come from?
Mirids overwinter as adults on wild hosts. In the spring with rising temperature, mirid populations start to build up on alternative hosts around cotton growing areas. Wet winters usually contribute large reserves of alternative hosts which support mirids. Once these hosts hay off and lose their suitability, the mirids can invade seedling or squaring cotton. Mirids sometimes build up in western Queensland, from where they allegedly move on north-westerly winds and invade cotton areas. Mirid adults also continually move into cotton from surrounding hosts and move out of the cotton to alternative hosts throughout the season.

What is happening this year?
Although the past winter was relatively wet, mirid numbers have been very low across the cotton growing areas so far this year. The reason for this phenomenon is not well understood. It could be that alternative hosts are still fresh enough to continue to support mirids, or the severe and prolonged winter may have had a detrimental effect on mirid populations.

Do we need to be worried about square loss at this stage?
If square retention is above 70% and mirid numbers are very low (as is the case for many crops this year), there is no need to be unduly concerned. Squares may be lost for various reasons e.g. insect feeding and physiological reasons.

If mirid numbers are very low and retention is less than 70%, square loss may be due to reasons other than insect feeding and a spray will not help. Square loss at this stage will be compensated fully provided plants do not further suffer from water or nutrient stress or from any other insects.

Figure 1. Compensation for mirid damage at squaring stage

Figure 1 shows plant mapping data in Bollgard® II.

Observation 1 shows the percentage fruit loss for 3 treatments at the squaring stage while observation 2 shows the same 3 treatments at cut out. The treatments consisted of the following:

In treatment 1 (blue) plants were sprayed on a regular basis starting from when 60% of plants had their first flower until cut out. Treatment 2 (red) was not sprayed for mirids at any time during the season. Treatment 3 was sprayed regularly throughout the season to provide total protection against mirids.

In observation 1 – the squaring stage – there is a significant difference between treatment 1 (sprayed from squaring) and treatment 3 (sprayed all season) with 18% fruit loss for treatment 1 and 8% loss in treatment 3.

However if we now look at what happened in these treatment at cut out, we find that there is no significant difference between treatment 1 and 3. What this shows is that in this experiment the plants had the ability to make up for the early season fruit loss.

Consequences of unwarranted mirid sprays at this stage
There are several consequences of an insurance spray.
- It is an unnecessary expense.
- Disruption of benefcial communities at this early stage could flare secondary pests such as aphids, mites and whitefly.
- Sprays at this stage, whether high or low rate, will not provide long residual protection. Mirids could move in from alternative hosts at any time, and it is best to save the spray for when it is really warranted.

Article by Dr Moazzem Khan

Thresholds and changing sorghum crop value

2008-12-16T08:47:00.012+10:00

Growers and consultants need to revise the thresholds used for important insect pests of grain sorghum such as corn earworm and sorghum midge in light of lower prices currently being offered.

With new crop grain sorghum prices below $200 per tonne, the break even cost of control means that higher pest numbers (density) are needed before control becomes economic, compared to thresholds used last season when grain values were much higher.

The use of a benefit:cost ratio is also an important consideration. The break even point is where the cost of control is equal to the loss caused by the pest. The benefit:cost ratio will vary according to individual preferences, and needs to be factored in to calculations.

Corn earworm
One corn earworm larva is estimated to consume 2.4 g of sorghum during its lifetime. The economic threshold (i.e. the number of larvae/head where the cost of control is equal to the value of the grain saved) can be calculated using the formula:

No. larvae/head = (C x R) ÷ (V x N x 2.4)
where
C = cost of control ($/ha)
R = row spacing (cm)
V = value of crop ($/tonne)
N = number of heads/metre of row
2.4 = weight of sorghum (grams) lost/larva

The value of crop loss caused by corn earworm larvae in grain sorghum, for a range of larval densities and grain prices and based on 10 heads/metre of row on 1 metre row spacing, are presented below.

When sorghum is valued at $300/tonne, one larva/head could cause $72 crop loss/ha. If the price drops to $150 per tonne, one larva/head causes just $36 crop loss/ha, or 50% less economic damage. This example demonstrates just how important it is to consider each case on its merits, and in particular to consider the cost of control, as it too can vary widely depending on whether aerial or ground spraying is used.

Sorghum midge

As with corn earworm, decisions to spray for midge are greatly influenced by crop value and it is possible to calculate theoretical yield loss estimates for a particular crop scenario (see table). These yield loss estimates are based on extensive field trials by DPI&F that determined the average yield loss/midge/day on different rated midge hybrids. For a susceptible hybrid, one midge is estimated to cause 1.4 g yield loss/day. (Photo: D. Ironside)

Listed below are estimates of midge damage at different grain prices without chemical treatment.

The yield loss estimates in the table assume that spraying results in a 100% kill and that there is no midge damage prior to chemical application. It also assumes that the same average midge pressure persists over 4-5 days. In reality research has shown that one well timed insecticide for midge (put on from panicle emergence and before midge even enter the crop) will still only prevent 70-80% damage protection in lower rated sorghum hybrids. In 8+ rated hybrids, yield losses can be reduced by over 90% with this spray timing.

If the total cost of applying a synthetic pyrethroid by plane is around $20/ha, we can see that at a grain price of $150 per tonne, it is simply not economic to spray mid to high rated hybrids at 1 midge/head and 8+ hybrids at 3 midge/head. It should be emphasised that 8+ is the highest rating that can be assigned to midge resistant hybrids. There are some 8+ lines that would have a considerably higher rating if the scale was extended, and are practically immune to midge damage.

The above information shows the importance of calculating thresholds for the current situation, rather than relying on a fixed value from one year to the next.

Bring on NPV against grubs on grain sorghum

2008-11-26T11:33:00.016+10:00

Every year caterpillars of the corn earworm (helicoverpa), Helicoverpa armigera, cause losses to sorghum crops. Regular inspection during flowering is important to detect caterpillar infestations and properly time control measures.

Pre-flowering heads of grain sorghum are very attractive to egg-laying moths of the corn earworm. On any individual head, most eggs are laid prior to the start of flowering, as indicated by the presence of yellow anthers.

By the end of flowering, when brown anthers are present at the base of the head, eggs will have hatched and most larvae will be less than 7 mm in length.

A timely spray application of the naturally occurring biopesticide NPV (nucleopolyhedrovirus) remains the best control option for grain sorghum crops under attack from corn earworm.

NPV performs exceptionally well on grain sorghum, with well timed sprays usually achieving greater than 90 per cent control while leaving beneficial parasites and predators to mop up survivors.

If the spread of flowering in a crop is large, it may be better to spray earlier rather than wait until 50% of the crop is at the brown anther stage. This is because caterpillars on the earliest flowering heads may be larger than the ideal size to target with NPV, and they will cause some damage if not adequately controlled with NPV.

Research has shown that early application of NPV creates a disease outbreak and secondary NPV infection will control most caterpillars on late flowering heads.

Other issues to ensure good results with NPV

Water quality
Water used in spray mixes should have a pH of 7. Alkaline water will seriously reduce the performance of NPV, so buffer water with Li700 or equivalent to neutralise pH.

Water volumes
For high-volume, water-based sprays, a minimum of 30 L water/ha is recommended for aerial application, and 100 L water/ha for ground rig application.

Coverage
NPV must be ingested to be effective, so the challenge is to achieve good coverage of the target. This means paying particular attention to water volumes, nozzles, operating pressure, weather conditions, etc. You want to spread NPV over as much of the head as possible to ensure caterpillars have a high chance of picking up a lethal dose as they feed on the head.

Additives
Additives such as Amino Feed, etc. are not recommended when NPV is applied to grain sorghum.

Paying attention to the detail will ensure the best results from NPV.

Article by Dr. Dave Murray

New Beat Sheet contributors

The beat sheet blog team has been expanded and includes two new contributors, Kate Charleston and Zara Ludgate.

Kate is the development extension officer with the entomology team in Toowoomba. Her role is to provide information about IPM in field crops as well as training to growers and industry in managing insect pests according to IPM principles.

Kate joined entomology in June 2008 and has previously worked as a research scientist, agronomist, plant health inspector and extension officer. She started her career with the Department of Primary Industries in Tasmania and joined the Queensland Government in 1999 as an extension officer in a sugar project at South Johnstone in Far North Queensland. Kate has worked with sugarcane, cotton and pulse crops and has considerable training and extension experience.

Zara has just started her career in entomology research. She comes from a rural background and completed a degree in plant and soil science at St Lucia, Brisbane in 2007. For her honours year she investigated the effect of a plant defence compound on the fitness of diamondback moth.

While in Brisbane she provided technical support for research into bio-pesticide production for helicoverpa and green vegetable bug management. She is now based in Toowoomba with the crop protection systems - entomology unit. Her current research interests include insecticide resistance in whitefly and integrated pest management in grain crops.

Managing Helicoverpa larvae in chickpea crops close to dessication and harvest.

2008-11-05T12:28:00.013+10:00

Over the last week or so we have received a number of enquiries about how best to manage new egg-lays, and populations of small larvae, in chickpea crops that are close to dessication and senescing.

Of most concern are crops that still have reasonable areas of green crop in them, and what the likelihood of damage is if the weather is cool and moist rather than hot and dry.

Hot, dry weather will rapidly advance a chickpea crop which means that very small and small larvae are unlikely to survive on leaves of rapidly deteriorating quality. As the pods dry they also become more resistant to damage by small to medium larvae. In summary, this means that the major source of damage in a scenesing crop is late medium and large larvae.

Therefore, the recommended approach to managing Helicoverpa populations in the later stages of a chickpea crop is to continue to monitor both number and size of larvae. If the population of medium and large larvae exceeds the economic threshold, AND the crop is still susceptible then treatment may be warranted.

The table below gives an indication of how rapidly Helicoverpa larvae will develop at this time of year.

Predicted development times for Helicoverpa larvae (Oct-Nov 2008) - Dalby
Up to 3 November the prediction uses 2008 temperatures for Dalby. Beyond 3 November, the predictions use long term average temperatures (long term averages are generally cooler and development slower).

The predictions indicate that larvae are developing from very small to medium in around 7 days and from small to medium in 3 days.

At this stage of the crop, a wait and see approach (continue checking the crop 1-2 times a week) to is recommended principally because it is difficult to predict a week or two ahead how fast a crop will dry down, and what the Helicoverpa population will be whilst the crop is still susceptible. The alternative approach is to treat above threshold populations of small larvae when they are detected. This approach is likely to result in treatment of fields that subsequently would not have been at risk of damage, particularly if the crop dries faster, or larval mortality is higher than expected.

The options available for the treatment of Helicoverpa infestations late are limited because of withholding periods (WHP). Methomyl has a 1 day WHP while thiodicarb has a 21 day WHP. Indoxacarb (StewardTM) has a 21 day WHP, but no more than one application is permitted per crop growth cycle, and the cut-off for indoxacarb use has now passed in all regions (15 Sep in CQ, 15 Oct in warm areas, 30 Oct in cool areas). Check with others in your local area on their experience with the efficacy of options when making a choice.

Armyworm in wheat

2008-10-20T10:59:00.011+10:00

Over the past couple of weeks there have been numerous reports of armyworm in both barley and wheat. The appearance of armyworm in wheat raises a number of questions:
1) Do they behave the same way in wheat as in barley in relation to the type of damage they cause
2) what is their damage potential and is there an economic threshold?
3) What sort of strategy can be used to monitor and manage populations?

For information on armyworm identification see previous Beatsheet postings on armyworms.
http://thebeatsheet-ipmnews.blogspot.com/2007/10/can-you-confidently-identify-armyworm.html

http://thebeatsheet-ipmnews.blogspot.com/2007/09/watch-for-armyworms-in-barley-and-oats.html

There is no reason to expect armyworm to behave differently in wheat to barley. This means you can expect to see feeding on leaves whilst the crop is still green, and then on stems as the crop dries down further.

Characteristic armyworm damage in winter cereals
During the vegetative growth phase, plants can tolerate considerable leaf feeding. Leaves may look tattered from the eaten-out leaf margins. Faecal pellets around the base of plants are another indication of armyworm infestation. Armyworm generally do not require control during the vegetative stage.

Ragged flag and other leaves on a maturing barley crop

The most serious armyworm damage in cereal crops occurs when larvae feed on the upper flag leaf and stem node as the crop matures. Larvae target the stem node as the leaves become dry and unpalatable, and the stem is often the last part of the plant to dry. Head cutting begins at this time.

One large larva can sever up to seven heads of barley a day. One larva a square metre can cause a loss of 70 kg/ha grain per day. A larva takes around 8-10 days to develop through the final, most damaging instars, so the crop is susceptible to maximum damage for this period.

Head cutting in barley caused by armyworm

Calculating an economic threshold
The following table shows the value of yield loss incurred by 1 larva/square m per day, based on approximate current values for wheat and an estimated loss of 70 kg/ha per larva.

Based on these figures, and the relatively low cost of controlling armyworm, populations in ripening crops in excess of 1 large larva per square metre will warrant spraying.

Monitoring and management strategy
For insecticide application to be economic, check or scout the crop and assess the problem before head cutting starts. Check for larvae on the plant and in the soil litter under the plant. Late in the day, when the larvae are becoming active, use a sweep net (or swing a bucket through the crop) to make a quick assessment of whether armyworm larvae are present in the crop. Infestations are often patchy, so check a number of sites across the field.

Some judgements will need to be made about how quickly the larvae will reach damaging size and when this will occur in relation to the crop's development.For example, if the crop is nearing full maturity/harvest, and the grubs are still small, then there is most likely no need to spray. Small larvae take 8-10 days to reach a size capable of head lopping.The other extreme would be a late crop that is still very green and at early seed fill. In this case, any small larvae present will most likely reach their most damaging size in time to significantly reduce crop yield, and so a spray is more likely to be required.

I you are unable to monitor the crop on a regular (daily) schedule during the critical period of drying down, and armyworm are present, it may be better to spray just in case. This is not the preferred option, but provides peace of mind in a year like this where armyworm are abundant.

Early recognition
It is essential to recognise the problem early and be prepared to spray when economic damage is imminent. A cereal crop can be almost destroyed by armyworm in just a few days. Whilst large larvae do the head lopping, controlling smaller larvae that are still leaf feeding may be more achievable.

Control
Many chemicals will control armyworms. However their effectiveness is often dependent on good penetration into the crop to get contact with the caterpillars. Control may be more difficult in high-yielding thick canopy crops, particularly when larvae are resting under leaf litter at the base of plants. As larvae are most active at night, spraying in the afternoon or evening may produce the best results.

If applying sprays close to harvest, be aware of relevant Withholding Periods. Always read the label.

Biological control agents may be important in some years. These include parasitic flies and wasps, predatory beetles and diseases.

Helicoverpa management in chickpea – a refresher

2008-10-08T14:14:00.019+10:00

A comprehensive overview of Helicoverpa management in chickpea can be found in the DPI&F brochure Helicoverpa management in chickpea (2007). You can read or download a copy of the brochure at the DPI & F website at www.dpi.qld.gov.au/fieldcrops. Click on the link to Helicoverpa management in chickpea where you will find the brochure.

Key management decisions
The following is an excerpt from the Helicoverpa management in chickpea brochure, and deals specificially with determining whether an infestation of helicoverpa warrants control – based on the economics of potential yield loss vs cost of control.

If control is warranted, which product?
There is a range of products registered for helicoverpa control in chickpea. However, the use of synthetic pyrethroids is really only an option in regions where H. punctigera dominates, or where the population is predominantly made up of larvae smaller than 5 mm in length. The use of SPs against a predominantly H. armigera population is likely to deliver a poor result in terms of control.

NPV (VivusMax) and Bt (e.g. Dipel) are two options which are effective against both species of Helicoverpa. They are most efficacious when deployed to control populations of small larvae (less than 7 mm in length), and lower pressure infestations.

Thiodicarb (Larvin) is another option, particularly where efficacy of this product in the local area is known to be high. Methomyl (Marlin®) could be considered whare large larvae are present close to harvest as it has a 1 day withholding period

Spinosad (Tracer II ™) and indoxacarb (Steward ™) are both effective against both H. armigera and H. punctigera. Remember that Steward has a cut-off for use in chickpea (15 September in CQ, 15 October in warm areas, 30 October in cool areas).

One strategy for the management of mixed age populations of helicoverpa is to use Steward™ first, if prior to cut-off, and then one of the other products if the crop needs to be sprayed again.

What are those grubs in winter cereals?

2008-10-01T14:45:00.005+10:00

Grubs in winter cereals are not unusual at this time of year, and already there have been reports of high numbers (up to 20/m2) in Central Queensland wheat (Figure 1). More grubs can be expected in southern districts as the season warms up.

The two most likely larvae (grubs) found in winter cereals are the corn earworm, Helicoverpa armigera, and the common armyworm, Leucania convecta. See previous blog postings for more information on these pests.
http://thebeatsheet-ipmnews.blogspot.com/2007/10/can-you-confidently-identify-armyworm.html

Figure 1. Large corn earworm larva on a wheat head. (Photo: R. Lloyd)

All Helicoverpa larvae found feeding in wheat, barley or triticale crops will be corn earworm. The native budworm, H. punctigera, is not normally found on monocots (grasses). This is important to know, because the corn earworm has developed resistance to pyrethroids, and unless the larvae are small, a pyrethroid spray is unlikely to control them.

If large larvae are present, identification becomes a somewhat academic issue. However, large H. armigera larvae can be identified by the white hairs behind the head (Figure 2). In contrast, the hairs on large H. punctigera larvae are black. These compare with armyworm larvae which have three pale stripes just behind the head, and smooth skin, without any hairs or bumps.

Figure 2. White hairs behind the head of corn earworm larva. (Photo: R. Lloyd)

If corn earworm infestations are detected early and larvae are small, preferably less than 7 mm in length, Helicoverpa nucleopolyhedrovirus (NPV) sold as Vivus Max could be considered as it will not harm beneficials (predators and parasites) in the crop. Some caution is needed as NPV will not kill corn earworm larvae greater than 13 mm in length, and will have no effect on armyworms.

Invariably when larvae are found on cereal crops, they are medium or large (>13 mm in length) and a more robust option is needed to control them. Both corn earworm and common armyworm are usually present in winter cereals, and control measures will be influenced by the relative abundance of each.

Follow the link below for more information related to thresholds and control options.
http://thebeatsheet-ipmnews.blogspot.com/2007/10/are-corn-earworm-problem-in-winter.html

Cereal Aphids in wheat and barley Spring 2008

2008-09-25T13:25:00.012+10:00

Cereal aphid numbers have increased rapidly over the past 3 weeks as the temperatures have increased. Whilst low numbers of aphids have been present in many crops (wheat and barley) for some time, it was not until a couple of weeks ago that numbers reached levels of concern to agronomists and growers.

Until the last few seasons, cereal aphids have not been considered a major pest in winter cereals. However, higher grain prices mean that the value of any yield loss is higher than it was and control may be economic at the densities we are experiencing.

Which species are in crops this season?

There are several species of aphid that occur in winter cereals (oats, wheat and barley). The most abundant, and the species that has been present in low numbers through winter are the oat aphid (Rhopalosiphum padi – it sounds like Row-pal-o-si-fum pad-i). This species tends to colonise the lower portion of the plant, mature adults are a dark green and rounded. Juveniles are paler and smaller.

On the Downs, the oat aphid is currently the dominant species, with infestations extending from around the base of plants up on to leaves and stems as the crop starts elongation. Smaller number of the rose-grain aphid and corn aphid are also present.

The rose-grain aphid (Metopolophium dirhodum – sounds like meto-pal-o-fee-um di-road-um) is a large, pale aphid with a dark stripe down the midline of the back. This species tends to colonise leaves higher on the plant, and is often very obvious. Clusters of juveniles are common on upper surfaces of leaves.

The corn aphid (Rhopalosiphum maidis – sounds like Row-pal-o-si-fum may-dis) is rectangular in shape rather than round. Legs and antennae are typically dark, the body green-blue, and they may look waxy.

(line drawings from “Insectopedia” Agriculture Victoria, 2000)

In northern NSW, the corn aphid is abundant higher in the canopy, particularly in crops that are booting. Corn aphid is reputed to decline in number as the crop comes out into head.

The photo illustrates a typical corn aphid infestation in a crop of barley prior to head emergence.

How much damage can aphids cause?

There has been surprisingly little work done on cereal aphids in Australia to establish the relationship between aphid numbers, the duration/timing of infestation, aphid species, and ultimately the impact on yield.

Direct aphid damage, as a result of feeding, is difficult to detect. In moisture stressed crops you may see yellowing with high aphid populations. Otherwise, there are generally no early signs of how much impact the aphids are having on the crop.Western Australian recommendation are to check crops regularly from late tillering, and consider control if the aphid population exceeds 15 aphids/tiller on 50% of tillers.

The WA research showed yield losses of up to 10%, and reduction in seed size, with aphid infestations (this was without any impact of barley yellow dwarf virus).

http://www.agric.wa.gov.au/content/pw/ph/dis/cer/bydv_aphidfeeding.htm

Queensland DPI&F recommendations have been to:
Check 5 plants at 6 sites within the field. If 27/30 (90%) of plants are covered with aphids, and there are less than 2 natural enemies per plant, then consider treatment.

A 90% infestation level would be indicative of a well established population. Early infestations tend to be patchy, and become more uniform as the population builds up.

Checking a crop for aphids
Sample away from the edge of a field. Aphid numbers tend to be higher around field margins because this is where initial infestations start. The rest of the field will be more representative of the infestation in the majority of the field.

It is simpler to base estimates of infestation on tillers rather than whole plants. It can be difficult to determine where an individual plant starts and stops, and the number of tillers per plant can be variable.

Taking a representative sample of individual tillers from across a field will provide information on the number of aphids, and the proportion of the tillers infested. The lower the infestation the more tillers you will need to sample (e.g. 30 per management unit). The more established the population the more uniform the infestation will be and the number of tillers sampled can be reduced (e.g. 10-20 tillers may be sufficient). Record the number of aphids per tiller and see how consistent numbers are as you go. Lots of zeros means the population is patchy.

If numbers are high, you may want to use a rating system for estimating density rather than actually counting aphid numbers.
For example: 0= no aphids, 1= 1-10 aphids, 2= 10-20 aphids, 3= 20-50 aphids, 4= more than 50 etc. Once you have your eye in, a rating system is quicker than counting aphids.

It may be useful to rate the number of aphids above and below the flag leaf separately. This will be particularly useful for assessing how effective a spray has been, and determining if surviving aphids are those that may have simply not been contacted.

Information from overseas research (Canada, US) suggests:

that significant yield loss occurs when aphids are present from the flag leaf stage through to milky grain – no yield loss occurs with infestations later than milky grain
infestations of aphids on the flag leaf, and upper portions of the crop, including on the heads, cause greater yield loss than infestations lower in the canopy

Other considerations when making a decision about cereal aphids

Corn aphids may disappear by themselves. Corn aphids, the species that colonises the upper canopy, reputedly decline in number when the crop comes into head. This may be because they tend not to survive as well on leaves as they do on the flag leaf or in the whorl.
Natural enemies (lady beetles, hoverflies, parasitic wasps) can have a big impact on aphid populations, reducing them to very low levels in many instances. This is particularly important in managing the resurgence of any aphids that survive a spray.
Dimethoate and synthetic pyrethroids (e.g. Bulldock®) are highly disruptive to natural enemies. The application of these insecticides early may result in a later reinfestation of the crop because small numbers of surviving aphids are no longer controlled by natural enemies. The impact of these products on natural enemies can persist for some days.
Pirimicarb (e.g. Pirimor®) is a soft option for cereal aphid control, but be aware of the with-holding period.
there is no Australian data on resistance to any of the registered insecticides in cereal aphid populations.
Oat aphids, at the base of the plant, can be difficult to contact in a dense crop, and with aerial application.
Rain will reduce aphid populations by knocking/washing individuals of plants, particularly if the rain is high intensity (storm) rain. When washed off, aphids tend not to get back on the plants. Often ground predators, like carabid beetles, ants etc will eat aphids on the ground. It may be worth re-checking numbers if you get a storm between checking and applying a spray.

Farm hygiene important in pest management

2008-04-17T15:10:00.013+10:00

David Murray, Toowoomba

Farm hygiene is an important component of integrated pest management (IPM), particularly when it comes to managing pests such as cotton aphids and Cotton Bunchy Top (CBT) disease.

Recent surveys of cotton-growing areas indicate the presence of aphids and CBT, and wetter conditions through the coming winter could favour the growth of weeds that are hosts for aphids and also increase survival of cotton volunteers that carry CBT to the next season.

The photo (right) shows volunteer cotton with CBT symptoms adjacent to the current season cotton crop (Photo: Lewis Wilson, CSIRO)

CSIRO and Cotton CRC entomologist Dr Lewis Wilson suggests that growers maintain good farm hygiene to reduce the risk of aphid or CBT problems next season.

CBT can stunt the growth of cotton plants and, if plants are infected when young, dramatically reduce yield.

CBT is spread by cotton aphids when they feed. Both cotton aphids and the disease need a host plant for survival through winter.

Cotton is a good host and volunteer or ratoon cotton plants can be found on farms all year. These plants can carry the disease and aphids through winter. Aphids can then move to cotton crops in the following spring and infect plants with the disease.

Photo: Fallow field with a high number of volunteer cotton plants, a potential resevoir for cotton aphid and CBT (Lewis Wilson, CSIRO).

While it is likely that CBT will survive on alternative weed hosts, these relationships have not been studied.

In recent field inspections on the Darling Downs, cotton volunteers were found on virtually every farm.

Many of these volunteers showed clear symptoms of CBT, such as angular leaf mottling and small leaves (see photo below, Photo: Lewis Wilson, CSIRO).

Cotton CRC extension officer Rod Gordon said that regular rainfall through the summer had encouraged growth of cotton volunteers, many of which were glyphosate-tolerant and difficult to control with herbicides. Wet fields have also limited opportunities to control volunteer cotton, with some fallow fields and field margins containing quite high densities.

The importance of controlling volunteer cotton in fallows can not be stressed enough. Minimising host availability is critical to ensure that we do not see a repeat of the 1998 CBT outbreak.

More information on cotton aphid and CBT can be found on the Cotton Catchment Communities CRC website at
http://www.cottoncrc.org.au/content/Industry/Publications/PestsandBeneficials/CottonBunchyTopCBTCharacteristicsandModes.aspx

Accessing the Diapause Tool - An alternative address

2008-03-27T11:48:00.003+10:00

If you are having trouble with the link to the Diapause tool provided in the story below, try the following:

http://tools.cottoncrc.org.au/xl2/diapause/index.aspx

Diapause tool to identify helicoverpa risk

2008-03-20T12:13:00.008+10:00

While there was a hefty presence of corn earworn or cotton bollworm, Helicoverpa armigera, in the early and middle part of the 2007-08 season, pest activity has declined in recent weeks and for the most part they appear to pose no major risk.

What is diapause?
This is the time of the year when a proportion of mature larvae going to ground to pupate enter a hibernation phase termed diapause or overwintering. This dormancy strategy allows the pest to survive the winter months in temperate regions when host plants are scarce and temperatures are generally too low to allow successful development. The triggers to enter diapause are decreasing daylength and temperature as experienced during late summer and autumn.

Picture of helicoverpa pupa in earthen cell.

The proportion of pupae entering diapause increases from low levels in March, to high levels, almost 100%, by late April. The rate of diapause induction varies from season to season, and region to region. Knowing when diapause is induced is useful for identifying ‘high risk’ fields i.e. those fields most likely to have diapausing pupae.

A web tool is available on the Cotton CRC website to help calculate the likely rate of diapause induction for your area, based on local climate data. The tool is also able to compare the results for the current season with the long term average and hotter than average and cooler than average seasons. Follow this link:
http://tools.cotton.crc.org.au/cl2/diapause/index.aspx

How are diapausing pupae controlled?
Overwintering pupae are very important because they contribute to the spring population and may take with them the resistance genes enabling them to tolerate conventional insecticides and the Bt transgenic toxins found in Bollgard II®.

It is for this reason that full soil surface cultivation to 10 cm depth (also known as pupae busting) is so important. When carried out properly, pupae busting can reduce survival of overwintering pupae to less than 5%.

Pupae busting is mandatory for all Bollgard II® fields; it is a requirement of the Bollgard II® licence.

Some relaxation of pupae busting requirements has been introduced for conventional cotton fields. Sprayed conventional cotton crops defoliated after 9 March are more likely to harbour insecticide resistant H. armigera pupae and should be pupae busted as soon as possible after picking and no later than the end of August.

The same holds true for the majority of grain crops on the Downs. Crops that were mature on or before 9 March are unlikely to harbour overwintering pupae. Conservation tillage (zero till or min-till) can be confidently used on these fields. Late season grain crops that could support development of larvae after 9 March are ‘high risk’ of harbouring overwintering pupae, and each field should be judged on its merits. The decision on whether to pupae bust will be influenced by the density of larvae present in the crop, the age of the larvae, and the timing of their presence in the crop.

Parasites will also influence survival of overwintering pupae, with estimates from a 5 year study on the Darling Downs showing 44% of overwintering pupae were parasitised. The two-toned caterpillar parasite, Heteropelma scaposum, was the most abundant parasite species recorded.

What is the current seasonal outlook for diapause in cotton and grain crops?
Output for the current season at Dalby, which is cooler than average, indicates a higher than average proportion of pupae have entered diapause.

All cotton crops, and a large proportion of the grain crops on the Downs, and in most other areas, will be harvested later than mid March. This means it is worth checking and recording whether these crops are hosting larvae that will diapause. Crops determined to be ‘high risk’ will warrant pupae busting.

Whitefly management options

2008-02-29T13:37:00.005+10:00

In the last week, reports from the Downs are suggesting that silverleaf whitefly (SLW) numbers have increased rapidly, and now there are a number of fields that have a population at or exceeding the treatment threshold.

This posting has been compiled from information and discussion with Richard Sequeira (Principal Entomologist, Emerald) and Paul Grundy (Senior Entomologist, Ayr) who have considerable experience in managing SLW. The aim is to provide information that may help in making decisions about the need for and timing of control, and the appropriate control option.

It appears that there is a larger than predicted population of SLW in cotton this season, especially as the season is average in terms of temperature. Why is this so particularly on the Downs where it is predicted that outbreaks would only occur in hotter than average seasons?
The answer to this is that outbreaks of SLW are driven not only by temperature, but by two factors, the size of the initial population in spring and the summer temperatures. In the 2007-08 season we have experienced temperatures that are close to the long term average, but we would have started the season with a large carryover from the outbreak in 2006-07. The rainfall and weed growth in winter and spring would have provided hosts for the SLW to carryover from last season to this one.

Over the last week or so, SLW populations seem to be increasing with the percentage infestation rising rapidly. In Central Queensland (CQ) it is usual to see a rapid increase in the percentage infestation at the 5th node once the crops cut out. It looks as though the population is increasing in size, but what is actually happening is movement of the population up towards the top of the plant as it stops putting on nodes. In CQ the quality of the leaves lower on the plants also starts to decline around this time and becomes unsuitable for SLW, forcing them higher in the canopy. Paul suggested that there are visual clues to whether the population is increasing in number. Observing the amount of honeydew on the lower leaves can be instructive. Over a week, an increase from a light speckling to larger droplets is indicative of a population increase.

As a SLW population approaches threshold natural enemies will not contain it. It is interesting to note that there has been very little parasitism recorded from samples taken in St George, and on the Downs this season. Parasitism levels in CQ are also low. Richard’s interpretation of this is that wet weather and high humidity has a negative impact on the parasitoids.

As SLW populations approach, or exceed, the treatment threshold, questions arise around the issues of whether to control, when and what product to use. The SLW thresholds are available in the Cotton Pest Management Guide, and on the web at
http://www.dpi.nsw.gov.au/agriculture/field/fibres/cotton/cotton-pest-management-guide Section 2, pages 11-13

To use the threshold recommendations it is necessary to calculate the day degree (DD) accumulations for the season, this will enable you to match the crop stage to the appropriate threshold and management options. The current DD data for a range of cotton-growing regions is presented in the table below.

Weighing up the options
In making a decision about SLW control, it is important to consider not only the level of infestation but also the age of the crop (DD accumulation) , how long it has to go until open cotton or leaf drop, and what is going on in neighbouring crops.

The situation on the Downs currently is that there are crops at SLW threshold that do not yet have open cotton. In this situation, Paul suggests that a second check be made on the population a week after it is first recorded to be at threshold, just to make sure the population really is at that level. If it is, it is necessary to weigh up the options for control.

Richard’s sense was that without open cotton, there is no urgency to control the population. Potentially you can wait a week or two until closer to the first open boll before applying Admiral®. Certainly, the application of Admiral® should be closer to 1600 DD than 1500 DD (first open boll is at 1650 DD). The aim of this strategy to decrease the chance that there could be a resurgence of the population post treatment, still with time to contaminate open cotton. However, on the Downs, the expectation is that the temperatures will be cooling through March and into April, and the risk of a second large population is unlikely.

Another approach is to treat populations at threshold now with Admiral®, particularly knowing that it has 3 week residual, and that SLW will not start to breed in the crop until April. With this timeframe, particularly in cooling conditions, it is unlikely that SLW will build up to threshold before the end of the season.

Pegasus® will knock a population of SLW, and control aphids in the crop. However, given the length of season many crops still have to go, the use of Pegasus ® now may require a re-treatment with Admiral ® later. Whilst Pegasus ® is the less expensive option, it is more likely to require re-treatment if used now than the application of an Admiral ®. Pegasus ® is an option for late crops where SLW populations do not reach threshold until there is open cotton and a quick knockdown is needed late in the season. If using Pegasus® remember that it is a contact product and good coverage is essential for good control.

Whitefly Update

2008-02-22T16:33:00.017+10:00

Reports of increasing numbers of whitefly are filtering in from across the Downs. In some cases, numbers are sufficient to suggest that control of Silverleaf Whitefly (SLW) may be warranted. Correct identification and regular monitoring of populations is essential to determine if intervention is necessary.

Which whiteflies are out there?
Results of whitefly samples identified from Downs cotton crops this week indicated variable incidence of SLW. In all cases except St George, identifications were based on immature (scale) stages used to differentiate Bemisia tabaci (SLW and Native Bemisia) from Greenhouse Whitefly (GHWF). These results infer that all Bemisia were SLW. Very few parasitoids were found in collections.

Anecdotal evidence supports the above findings; that in some cases the whitefly infestations have been predominately GHWF. Where pyrethroids have been applied to control stink bugs (green vegetable bug and cotton stainers), whitefly numbers have been suppressed. While GHWF is susceptible to pyrethroids, SLW is likely to have some level of resistance to pyrethroids and will survive treatments. The risk is that use of pyrethroids and other broad spectrum insecticides used to control bugs may well flare SLW by disrupting natural enemies.

The Downs is considered a marginal area for problems with SLW because in an average season there is not enough heat and generally too few generations to create a problem. There is a perception that the 2007/08 season has been cooler than normal. Day Degree accumulations indicate that 2007/08 has been marginally cooler than average (see Table 1). Surprisingly, it has not been as cool as many have expected.

Management of SLW
Cotton crops are approaching the stage where decisions must be made on whether to control whitefly present in crops. Firstly it is important to know which whitefly is present. Is it SLW?

Given positive SLW identification, are they at levels that will present a problem later on in the crop cycle? To answer this question, careful monitoring of adult numbers is required.

Monitoring SLW infestations
Percentage infestation is based on a fixed sampling protocol, where a leaf is considered infested if 2 or more adult whiteflies are present. Full details of the sampling protocols and threshold values are available in the Cotton Pest Management Guide or the Cotton CRC website http://www.dpi.nsw.gov.au/agriculture/field/fibres/cotton/cotton-pest-management-guide

Control options
The objective of SLW control is to manage the population early to minimise exposure of open cotton to honeydew contamination.

Management options as defined by crop stage, accumulated Day Degrees and corresponding whitefly thresholds – include CONTROL, KNOCKDOWN and SUPPRESSION.
The CONTROL option involves application of an Insect Growth Regulator (IGR) and is appropriate for moderate to high densities after 1300 DD.
The KNOCKDOWN option involves application of conventional chemistry and may be used on low to moderate densities after 1600 DD to minimise contamination of lint from lower bolls.
Low to moderate densities after peak flowering (1300 DD) may also be treated with conventional chemistry for population SUPPRESSION so as to avoid the need for a later application of IGR.

Use of Insect Growth Regulators (IGRs)

Where % infested leaves is consistently equal to or above the corresponding threshold value (> 1 adult/leaf after 1300 DD), use IGR from 1500 DD but no later than 1600 DD (first open boll).
IGR - Pyriproxifen (Admiral®) – is highly effective against SLW, gives excellent control across a broad density range and is the corner stone of effective SLW management in cotton as it is also very selective, allowing survival of predators and parasites.
Ensure only a single application of Admiral® occurs within a season.
Delaying the IGR application beyond 1600 DD and > 2 adults/leaf could result in yield loss, lower efficacy of the IGR, substantial lint contamination or all of the above.

Use of Conventional Chemistry

Adult SLW densities of 0.5-1 adult/5th node leaf beyond 1600 DD (first open boll to leaf drop) are sufficient to contaminate lint from earliest open bolls.
After first open boll (>1600 day degrees) and densities <>
Single applications of Pegasus® or endosulfan (if window permits) used early (1300 – 1450 DD) on SLW densities <>

What contribution might natural enemies make to SLW control?
Natural enemies make an important contribution to the suppression of SLW when populations are low. Natural enemies include ladybeetles that will feed on scale, the parasitoid wasps, and probably a number of predatory bugs. However, their potential impact is not significant enough to be taken into consideration once the SLW population gets to the point that in-crop control (IGR application) decisions are being made.

Whitefly in crops this season

2008-02-08T14:12:00.000+10:00

There is an increasing number of reports of whitefly in cotton crops on the Downs. Typically, whitefly start appearing in crops in mid to late summer, once populations have built up on weed hosts or other host crops (e.g. sunflower) adjacent to cotton crops.

So far this season we have been able to make collections of whitefly adults and immatures (scale) from two sites in the Jimbour – Macalister area. In both cases 75% of the population was Greenhouse Whitefly, and the remainder Silverleaf Whitefly (SLW) (or Bemisia B-type). The native Bemisia was rare in the samples.

We are keen to make further collections to get a picture of the whitefly populations across the region.

Typically, the Downs is at low risk of a SLW outbreak, except in seasons like 2006/07 when conditions are considerably hotter than average. In a relatively cool season like the current one, the expectation is that an outbreak is unlikely. This expectation is supported by the results so far (although limited) that indicate SLW populations to be low.

St George and Emerald are areas where conditions are suitable for a SLW outbreak in average years.

Whitefly as pests – a brief background
Whilst populations of Greenhouse Whitefly can build up in cotton, they are susceptible to a wide range of pesticides, as is the native Bemisia species. As a result Greenhouse and native Bemisia are often controlled incidentally when the crop is sprayed for other pest species.

Both Greenhouse and Silverleaf Whitefly produce honeydew as they feed. However, the honeydew of SLW persists on the lint through to processing, causing sticky cotton. See the photo (right) of open cotton severely contaminated with honeydew, and the black mould that grows on the honeydew. Sticky cotton is difficult to process, and as a result is highly undesirable in the market. Management of SLW is focused on preventing populations outbreaks and limiting the amount of honeydew, rather than limiting numbers to prevent direct damage to the crop from feeding.

The SLW’s pest status is as a result of it being well adapted to warm conditions, having a high reproductive rate, a wide host range, resistance to a wide range of insecticides, and the ability to rapidly develop resistance with exposure.

SLW has resistance to organophosphates, carbamates, synthetic pyrethroids, imidacloprid, endosulfan, bifenthrin, and amitraz. The level of resistance varies from region to region.

In cotton, infestations of SLW require careful management to prevent killing the parasitoids (wasps) that can contribute to the control of low numbers. Spraying to control sucking pests such as stinkbugs, mirids and aphids requires careful consideration in the presence of SLW. Where possible, avoid flaring SLW by choosing options other than broad spectrum insecticides such as synthetic pyrethroids.

Identifying whitefly species
There are three species of whitefly that can infest cotton in Australia, the Greenhouse Whitefly (Trialeurodes vaporariorum), the native Bemisia tabaci, and the Silverleaf Whitefly (Bemisia tabaci B-biotype).

Knowing which species is present in your crops is important, because only one of the three species, the Silverleaf Whitefly (SLW) has the potential to cause significant losses in cotton.

The adults of the Greenhouse and Bemisia species can be readily distinguised by eye. The Greenhouse Whitefly is about twice the size of the Bemisia species, and has overlapping wings, held flat over the body. The SLW and native Bemisia hold their wings tent like over the body and there is a visible split between the wings. See pictures left (Greenhouse) and right (SLW).

Juvenile stages of the Greenhouse Whitefly (the scale and pupae) can be distinguished easily from those of the Bemisia species with a good (x20) hand lens or microscope. Greenhouse Whitefly scales are white and a fringe of long hairs protruding from them. The Bemisia species pupae are yellowish in colour and are hairless (see pictures left and right above).

Eggs are laid haphazardly on the undersides of leaves, and are brown shortly before the scales hatch from them. When there are large numbers of eggs the undersurface of the leaf can look like sandpaper. See photo (right) of high density egg lay on the underside of a leaf.

The SLW and native Bemisia are indistinguishable from each other, and can only be separated by enzyme or DNA-based tests in the laboratory. This is the technique used to identify two collections made on the Downs this week.

More information on Silverleaf Whitefly and its management can be found on the website of the Cotton Catchment Communities CRC.

http://www.cotton.crc.org.au/content/Industry/Publications/PestsandBeneficials/Whitefly.aspx

Monitoring insecticide resistance in SLW populations
The management of SLW is highly dependant on a limited number of insecticides, particularly the insect growth regulator pyriproxifen (Admiral®). Together with the history that SLW has for rapidly developing resistance when exposed repeatedly to insecticides, insecticide resistance is a real threat. For this reason it is important to monitor populations of SLW from different regions to determine the levels of resistance they may have to key insecticide groups, and use this information to provide advice on appropriate resistance management strategies for SLW.

The DPI&F Entomology group in Toowoomba is doing this resistance monitoring for the cotton industry, and will be screening collections of whitefly from the field during the season.

We need your help to identify populations of SLW that can be included in the screening.
Contact Richard Lloyd at DPI&F in Toowoomba by phone on 46881315, or email: richard.lloyd@dpi.qld.gov.au.

Will Rutherglen bug damage sorghum post grain fill?

2008-01-16T15:04:00.000+10:00

As many of the early sorghum crops reach physiological maturity, and approach harvest, questions are being asked as to whether it is necessary to control large populations of nymphs in these crops. More specifically, whether these RGB will cause any damage to the maturing grain between physiological maturity and harvest.

Photo: Dave Murray (BigBug) out looking at a sorghum crop at Glen Ogden's property on the Darling Downs.

In summary, the best available information suggests that sorghum is will not suffer yield loss as a result of RGB feeding from physiological maturity (hard dough) through to harvest.

DPI&F trials to examine the impact of RGB on grain as it matured, did not provide a conclusive answer to this question. In 2008 we are undertaking field trials to try and answer this question.

However, if we look at what is happening with the sorghum plant from physiological maturity on, we can draw some conclusions about the potential for RGB to cause damage to maturing grain. Importantly, once grain reaches physiological maturity it has reached its full potential weight, and from then on starts to lose moisture as it matures. This means that even if large numbers of RGB continue to feed on the sorghum plant (on stems and leaves) their feeding will not impact on the development of filling of the grain at this stage.

Photo (left): Physiological maturity is reached when a black layer appears at the base of the seed, near to where the seed is attached to the stem

Another question that we do not have a definative answer to is whether RGB continue to feed directly on the maturing seed, or if they feed only on the sorghum plant once the grain reaches hard dough. In trials where plants at hard dough were exposed to RGB we did not see the evidence of feeding damage to grain that we saw when heads were infested at earlier stages of grain development (see earlier posting on RGB damage).

Crops that have had early infestations of RGB, even some of those that have been treated, now have moderate to large populations of nymphs and adults in them. The adults are likely to be newly emerged, having developed from nymphs in the crop. The nymphs will have emerged from eggs that were laid by an earlier infestation.

Controlling RGB to prevent the problems associated with infestations at harvest remains an issue. The inclusion of an insecticide along with the herbicide when the crop is being sprayed out prior to harvest is a practical approach. Be alert to the witholding period of any insecticide used. Treatment with insecticide prior to harvest is not a guarantee that the crop will be free of RGB at harvest. There remains the possibility of reinvasion by adults at any time.

Watch out for midge this season

2008-01-09T15:09:00.000+10:00

Adam Hardy

Senior Entomologist, Queensland Department of Primary Industries and Fisheries, Toowoomba.

Photo: Adam Hardy (right) and Bernie Franzmann inspecting midge rating trials.

Over the last decade sorghum midge have not caused many headaches for sorghum growers.

However, staggered sorghum plantings mean that sorghum midge are likely to be found building up in numbers in later planted sorghum crops with the potential to cause significant economic loss.

Even if midge are not in high numbers in a crop, the high grain price combined with the low cost of the insecticide of choice (synthetic pyrethroids) means that it is likely that spraying for midge will be a simple economic decision.

In the past the best way to avoid midge damage was to plant as high a midge rated variety as possible as early as possible in the season, and to plant crops well outside overlapping flowering windows of 2-3 weeks within districts. This has not been possible for many growers this season as rain came in a late and staggered fashion. This means that many crops planted over progressive flowering windows are likely to be at risk of midge population build up.

The only way to avoid economic damage is to monitor very closely for midge numbers EVERY DAY during head emergence and flowering and know what your spray threshold for midge is, based on crop value and cost of control, before the midge turn up. This insect pest requires very careful monitoring at exactly the right time of day. By the time you see the adult midge, they are already causing damage.

HOW TO COUNT FOR MIDGE

Generally, peak midge activity occurs between 9-11am, and this is the best time to look. However changes in weather can bring midge into a field from surrounding areas at any time of day. Midge numbers can differ widely both within a crop and between plants, even between those right next to each other.

Photo: Female midge laying into a sorghum floret.

Sorghum heads are most attractive to midge at mid flower. It is not uncommon to see double or triple the number of midge per panicle on panicles at early-mid flower vs pancles at the end of flowering. At lower midge densities, adult flies will move around and lay almost exclusively on the flowering portion of the panicle.

Midge flies are only 1-2 mm long, so if your eyesight is not what it used to be, then make sure you get someone with good eyes to carefully check each head. It is very easy to underestimate midge numbers if you are not careful.
The easiest way to 'get your eye in' is to look at the top half of mid flowering panicles and look for MOVEMENT of the small red flies against a still sorghum panicle looking from side on and slightly above side on one section of the sorghum panicle at a time. Keep your eyes focused over a couple of branches of florets for several seconds at a time to detect female midge walking around the branch or bobbing up and down probing their ovipositor into each floret. Female midge generally walk around quickly when they are ‘on the job’ spending as much time walking around as they do laying eggs.

On windy days you may have to hold each head still and shelter the panicle with your body or a sheet of cardboard (or a large jacket or umbrella?) for 10-20 seconds before scanning each panicle to allow you to more accurately see midge movement. If you can avoid the wind by getting out into the paddock slightly earlier in the morning then your job will be much easier.

Growers should monitor for midge over 10 metres of row in at least 4 different locations in your crop. It may be necessary to spray only one section of crop at a time, or the whole crop accordingly.

A common experience is that the first midge seen are caught in spiders webs in the field - a sign that midge are active in that particuar field.

As the season progresses, you may also start to see the midge parasitoid, Eupelmus spp (Photo: C. Freebairn). Whilst it can be present in reasonably large numbers, this parasitoid does not occur early enough to prevent midge from causing damage.

MANAGING MIDGE IN SORGHUM

Adult female midge lay their eggs inside sorghum florets where chemicals cannot reach. Insecticides only target the adult midge as they move about the crop and do not kill the eggs or hatched larvae that are already present inside the sorghum florets. While these midge adults only live for one day, they do most of their egg laying (and subsequent damage to the crop) in the morning.

It is possible to calculate theoretical yield loss estimates for a particular crop senario (see table). These yield loss estimates are based on extensive field trials by DPI&F that determined the average yield losses per midge per day on different rated midge hybrids

The yield loss estimates in the table assume that spraying results in a 100% kill and that there is no midge damage prior to chemical application. It also assumes that you will receive the same average midge pressures over 4-5 days. In reality research has shown that one well timed insecticide for midge (put on from panicle emergence and before midge even enter the crop) will still only prevent 70-80% damage protection in lower rated sorghum hybrids. In 8 rated hybrids, yield losses can be reduced by over 90% with this spray timing.

If the total cost of applying a synthetic pyrethroid by plane is around $20/ha, we can see that at a grain price of $250-300/t, it is economic to spray low to mid rated hybrids at even 1 midge per panicle and 8+ hybrids at 3 midge per panicle.

Many growers may choose to include a spray for midge with a synthetic pyrethriod in with a virus application for helicoverpa, or may look to clean up midge and rutherglen bug at the same time with one or two well timed pyrethriod applications. In all cases spraying pyrethoids will devastate beneficial insect populations and the implications of this should be included in the decision to spray. Pyrethroid applications on their own are likely to flare helicoverpa and aphid populations.

Rutherglen Bug in sorghum

2007-12-21T17:03:00.000+10:00

Rutherglen bug infestations of grain sorghum
This season we are seeing large infestations of Rutherglen bug (Nysius vinitor) (RGB) in sorghum crops from Central Queensland to the Downs, and further south. The large infestations are most likely a result of storm activity and the growth of weed hosts on which they breed up. RGB are very mobile, with large numbers of bugs appearing in (or disappearing from) crops before or after storms. There doesn’t seem to be particular stage of crop development at which RGB start to infest sorghum. The timing of infestation is determined by storm activity, or by hot dry weather that causes weed hosts to dry off, forcing bugs into nearby crop.

What we are seeing in the field this season
Where RGB are active, early sorghum crops, or early heads in a crop, are most affected. Damaged grain is red where bugs have fed on the portion of the seed not covered by the glumes. Dark spots are also visible on the grain. Grain is small and shrivelled, and has not continued to fill beyond the point at which it was damage. Cutting open the damaged grains, there is fungal and bacterial infection of the seed, making it black and mushy. This grain will not continue to fill, and is likely to be lost at harvest.

Sampling for RGB
Because RGB populations can start to build up at any stage of crop maturity, and our data show that RGB can significantly impact on seed set during flowering and early seed development, monitoring for RGB should start at mid flowering. Practically, RGB can be sampled for along with Helicoverpa; beating individual heads into a bucket. The distribution of RGB is typically patch across the field. A few heads with high numbers, lots of heads with lower numbers. This means that more, rather than fewer, heads need to be sampled to get an idea of the overall level of infestation in a field.

RGB and their damage potential
….. in sunflower
There has been little research on RGB in any crops other than sunflower. In sunflower they are known to be a major pest, feeding on the seed as it sets and matures, resulting in reduced yields and oil quality.
…….in sorghum
In the USA there is a related species, the false chinch bug (Nysius raphanus), which causes yield loss and reduced seed viability in sorghum. Yield loss is caused not only by the direct feeding of the bugs, but also through allowing a fungus to infect the seed through the feeding wounds, causing further deterioration and discolouration of the grain. Threshold trials showed that chinch bug caused a reduction in seed set when present on grain from flowering through soft dough stages. Where seed set was reduced, overall grain weight increased. In other words, the plant compensated with fewer, heavier grain in damaged heads.

DPI&F research on RGB in sorghum - what we know

RGB feeding post flowering will reduce grain set
Preliminary research on RGB has been done in sorghum and has shown that adult RGB will reduce seed set by around 20% at densities of 50-100 bugs/head. If bugs infest after grain is set, adult RGB will feed on the seed. Affected seed looks spotty and red externally (see photo), and hollowed out internally. Fungal infection also occurs resulting in further deterioration of the developing grain and ultimately seed that is small, shrivelled and light – and likely to be lost at harvest.

Early instar nymphs do not damage grain
No evidence of grain feeding was found for first, second and third instar nymphs. Older nymphs did feed on developing grain as do adults.

There are no soft-options for RGB control
Fipronil, indoxacarb and dimethoate were compared with alphacypermethrin in a replicated field trial on the Downs. None of the products provided control of nymphs statistically equivalent to the commercial comparison, alpha-cypermethrin. Compared to the unsprayed, alpha-cypermethrin was highly disruptive to predatory invertebrates.

What we don’t know

Can RGB damage grain from hard dough through to harvest?
The data for the impact of RGB on maturing sorghum does not provide a clear picture of whether grain continues to be damaged. We know that large populations of nymphs can be present in crops right up to harvest, but whether they are feeding on the plant, or on the grain are unknown.

RGB economic threshold for sorghum
Further work is needed to determine the density-damage relationship. This includes work to understand compensation when seed set is reduced early, the direct damage to grain during filling and maturation, and potentially indirect damage caused by RGB feeding on the sorghum plant which may impact on grain fill.

Soft options for RGB
Biopesticides, in this case the fungal disease Metarhizium is an option for RGB. It is currently being evaluated by DPI&F, and if effective will be an important addition to sorghum IPM.

Based on what we know, the following strategies can be suggested for minimising RGB damage to sorghum:

Start checking for RGB from flowering
Developing and filling seed are susceptible to damage from RGB adults, so protect the crop during these stages.

A threshold of 20-50 bugs/head is suggested. Treating populations of adults does not guarantee there will not be further infestation. RGB can reinfest a field overnight if the conditions are right.

RGB will breed in sorghum, so infestations of adults will result in increasing numbers of nymphs. Large populations of RGB can cause problems with machinery at harvest. Continue to monitor populations through to harvest as a decision to treat will need to take into account withholding periods for harvest.

Getting the most from NPV sprays on grain sorghum

2007-12-14T11:12:00.000+10:00

Some issues have recently been raised on the Darling Downs regarding the use of Helicoverpa nucleopolyhedrovirus (NPV) against corn earworm on grain sorghum. These issues involve the delay in time to kill and the level of control not necessarily meeting growers’ expectations.

Seasonal conditions
The last couple of weeks on the Downs have been cooler than normal. The average daily screen temperature for Dalby for the week ending 11 December was 25.5°C, while for the week ending 4 December it was just 22°C. Daily minimums were as low as 14°C. These lower temperatures will influence the time to death of NPV-infected caterpillars.

Temperature affects the development rates of caterpillars, their feeding, and the rate at which they die from NPV once they are infected. The lower the temperature, the slower caterpillars develop and the longer it takes for NPV-infected caterpillars to firstly stop feeding, and then to die.

Studies by Chris Monsour (formerly DPI&F) investigated these relationships. At 30°C, an NPV-infected 6-day old caterpillar (late second instar) will feed normally for 2 days before feeding is greatly reduced. At 20°C, caterpillars feed normally for 5 days before feeding is reduced. Importantly, the total amount of food consumed by these NPV-infected caterpillars is similar, whether at 30°C or 20°C (see Figure). Also remember that most caterpillars feed on anthers until late fourth instar (21 mm in length) and this feeding will not affect yield.

Time to death from NPV

At 30°C, it takes on average 4.5 days after infection for a 6-day old caterpillar to die from NPV. This compares with 6.2 days at 25°C and 7.5 days at 20°C. Under recent field conditions, many larvae are taking more than 8 days to die from NPV after spray application.

Figure. Consumption of artificial diet by healthy and NPV-infected 6-day old caterpillars and time to death from NPV at three temperatures (Data from Chris Monsour).

High grub infestations
If we accept that NPV will kill 90% of caterpillars, a starting infestation of 10 caterpillars per head results in 1 caterpillar per head surviving. This would not normally be an issue, but with the current high value of grain sorghum, this number of survivors may be above the economic threshold. You need to ask, is any other product going to do a better job? The answer is ‘No’.

Beneficial safety
NPV is safe on beneficials (parasites and predators) and these are important in helping mop up any survivors, as well as ensuring aphids are not flared. The end result is generally better than a 90% job.

What do we need to do to make sure of good results with NPV?

Timing
Consider applying NPV sprays earlier i.e. before 50% brown anthers, particularly if the spread of flowering is large. In this way most early flowering heads will be fully protected and secondary infection will control most caterpillars on the late flowering heads. It is best to target caterpillars less than 7 mm in length when using NPV, and this is the size of caterpillars on heads that have just finished flowering.

Water quality
Water used in spray mixes should have a pH of 7. Alkaline water will seriously reduce the performance of NPV, so buffer water with Li700 or equivalent to neutralise pH.

Water volumes
For high-volume, water-based sprays, a minimum of 30 L water/ha is recommended for aerial application, and 100 L water/ha for ground rig application.

Coverage
NPV must be ingested to be effective, so the challenge is to achieve good coverage of the target. This means paying particular attention to water volumes, nozzles, operating pressure, weather conditions, etc. You want to spread NPV over as much of the head as possible to ensure caterpillars have a high chance of picking up a lethal dose as they feed on the head.

Additives

NPV historically performs very well on grain sorghum, usually achieving greater than 90% control when used alone. For this reason, additives such as AminoFeed, etc. are not recommended when NPV is applied to grain sorghum.

However, for all crops other than sorghum, the use of a molasses-based additive containing the reducing sugars glucose and fructose (such as AminoFeed) is recommended when applying NPV.

Friendly fighter conquers foe

2007-11-30T09:33:00.000+10:00

Microplitis demolitor is just one of many friendly fighters that battle to contain numbers of one of our most important pests, the corn earworm, Helicoverpa armigera.

Corn earworm on grain sorghum is making its presence felt and many crops are being sprayed with Helicoverpa nucleopolyhedrovirus (NPV) to control above-threshold infestations of caterpillars.

The current high value of grain sorghum (over $250 per tonne) means that it is economic to control caterpillars at lower numbers (density) than growers may have sprayed previously when grain value was lower.

Low caterpillar numbers is a perfect situation for Microplitis to chip in a helping hand. It is not uncommon to find 30-40% of small caterpillars on grain sorghum parasitised by Microplitis. In many cases, this level of parasitism may be sufficient to sway a decision to not spray.

What is Microplitis?
Microplitis is a small native wasp that lays it eggs in (parasitises) small helicoverpa caterpillars. The life cycle from egg to adult takes about 12 days. This is made up of 7 days from egg laying to forming a pupa, and then 5 days for pupal development.

Adult Microplitis are small black-brown wasps. They are often seen flying slowly above the crop canopy in search of caterpillars (hosts). A female wasp will parasitise as many as 70 helicoverpa caterpillars. The parasite develops inside the host caterpillar. When fully developed, the Microplitis larva chews a hole in the side of the caterpillar and spins a fawn-coloured cocoon around itself and pupates. The caterpillar that was parasitised may still be alive, but it will soon die.

Clues to identify Microplitis activity include

Adult wasps foraging on sorghum heads
Split test of caterpillars to reveal internal parasites
Distinctive fawn cocoons next to dead or dying caterpillars

Identifying parasitised caterpillars
In the field, parasitised caterpillars can be identified by performing a simple split test. Parasitised caterpillars will only grow to about 15 mm in length, so caterpillars smaller than this are potentially Microplitis hosts. Hold a caterpillar across a forefinger with one thumb and forefinger on the rear end of the caterpillar, and with the other thumb on the head. Gently stretch the caterpillar until the skin ruptures. A Microplitis larva developing within the caterpillar looks like a white maggot up to 4 mm long.

Interactions between Microplitis and NPV
Caterpillars infected with NPV within 3 days of parasitisation by Microplitis will die from NPV. The immature Microplitis will also die because of the death of its host.

When NPV is applied to control corn earworm, it is not unusual for some parasitised caterpillars to survive the treatment. Caterpillars parasitised more than 3 days prior to the NPV treatment will produce healthy Microplitis. Parasitised caterpillars feed less and may not ingest NPV.

In shaking sorghum heads to make post-treatment assessments, parasitised larvae may be dislodged free of the pupal cocoon attached to them. Careful inspection of these caterpillars may reveal a hole in the side of some of these caterpillars, indicating prior parasitisation. These larvae will eventually die.

Microplitis is an important natural enemy of the corn earworm and they need to be considered when making decisions about when to manage corn earworm.

For more information on Microplitis, follow the link to the brochure ‘Microplitis demolitor and ascovirus: Important natural enemies of helicoverpa’ http://www2.dpi.qld.gov.au/fieldcrops/17576.html

Corn earworm chews into sorghum profits

2007-11-16T10:01:00.000+10:00

Sorghum growers across the Darling Downs can expect to see an influx of the corn earworm, Helicoverpa armigera, in their flowering sorghum crops over the next few weeks. Growers are well equipped to deal with the problem in an environmentally friendly way.

Moths are active and wanting to lay eggs on susceptible crops, and sorghum crops putting up heads are highly attractive - just what this insect pest loves. The majority of eggs are laid in a narrow window, between emergence of the head from the boot leaf and the commencement of flowering (yellow anthers). This results in highly synchronous development of larvae in a crop – larvae of uniform age in the crop.

Yellow anthers
during flowering
(too early to spray).

Eggs and newly hatched
larvae on sorghum.
Photo: D. Ironside

It is important that growers check their crops because in many cases feeding by corn earworm is likely to cause economic loss. One larva is estimated to consume 2.4 g of sorghum. Larvae up to 13 mm in length feed mostly on anthers and do not affect yield.

The table below provides examples of crop loss for different larval densities.
Table: The value of crop loss caused by corn earworm larvae in grain sorghum, for a range of larval densities and grain prices and based on 10 heads/metre of row on 1 metre row spacing.
*Based on estimated consumption of 2.4 g per larva.

The current high value of grain sorghum (over $300 per tonne) means that it is economic to control larvae at lower numbers (density) than growers may have sprayed previously when grain value was lower.

The economic threshold (i.e. the number of larvae per head where the cost of control is equal to the value of the grain saved) can be calculated using the formula:

No. larvae/head = (C x R) ÷ (V x N x 2.4)

where
C = cost of control ($/ha)
R = row spacing (cm)
V = value of crop ($/tonne)
N = number of heads/metre of row
2.4 = weight of sorghum (grams) lost per larva.

How to sample sorghum heads for corn earworm
Count the number of larvae dislodged from 30 heads to arrive at a control decision. Obtain 5 consecutive heads at the brown anther stage from at least 6 locations in a field, each location preferably more than 50 m apart. Use the palms of your hands to spin each of the heads into the bucket. Pour the contents of the bucket onto a beat sheet or tray and count the number of larvae in each size class
very small (VS=less than 3 mm in length)
small (S=3–7 mm)
small-medium (SM=7-13 mm)
medium-large (ML=13-21 mm)
large (L=greater than 21 mm).

Control
Effective larval control can be achieved with the use of commercially available nucleopolyhedrovirus or NPV sprays, sold as either Vivus Max® (succeeding Vivus Gold®) or Gemstar®.

NPV is dynamite against corn earworm larvae in sorghum and has the bonus of being a natural disease of the pest, so that spraying only kills the pest and not other beneficial insects and spiders in the crop.

Gemstar® and Vivus Gold® have both been registered for use on sorghum at 375 mL/ha. Lower rates (250-300 mL/ha) have been used successfully by many growers.

Please be aware that Vivus Max® now replaces Vivus Gold®. It is a more concentrated product (2.5 x) and has a registered rate of 150 mL/ha in sorghum (equivalent to 375 mL/ha of Vivus Gold®).

Research into the use of NPV sprays has shown several key points that growers and consultants should remember when using NPV.

First, checking is easy and important – it not only tells you whether you have the pest in enough numbers to justify spraying, but it also gives you information on when to time an NPV spray, since it works best when targeted against the very youngest larvae.

At the end of flowering (heads with brown anthers to base), most larvae will be first or second instar (less than 7 mm in length), and ideal to target with NPV. The best spray timing is when 50% of heads in the field have brown anthers to their base. A further delay of 3 days will help conserve the important larval parasite, Microplitis demolitor.

In crops where there is a large spread of flowering, it is better to spray before 50% of the heads are at the brown anther stage. In these cases, experience has shown secondary infection by NPV can kill a high proportion of the caterpillars that hatch after the NPV application.

NPV should not be used against larvae greater than 13 mm in length.

Good coverage over the plant and especially the sorghum head is critical, since a larva has to actually feed on an NPV particle to become infected with the virus. Sprays should be put on at the time of day that is best suited to getting good coverage, and this will often be in the morning.

Ultra low volume (ULV) sprays of NPV applied by a plane are highly effective in sorghum and allow for large areas to be treated in a relatively short time – this is good news when the pressure is on to treat large areas.

For ULV application, NPV is combined with approved spray oils such as D-C-Tron, Canopy or Biopest oil, to make a minimum spray volume of 3 L/ha.

Corn earworm larvae killed by NPV.

Further information on the use of NPV can be found in the brochure 'Using NPV to manage helicoverpa in field crops' by following this link

http://www2.dpi.qld.gov.au/fieldcrops/17677.html

Rutherglen bugs are everywhere!

2007-11-02T11:00:00.000+10:00

Rutherglen bug (Nysius vinitor)(RGB) is one of the insect species that arrives in crops in spring in large numbers, usually in association with storm activity. You may also have seen them on your windows and screens (and around the lights) at home in recent days. It is likely that the bugs are moving around in the environment, perhaps even transported from some distance away on storm fronts. You will probably find RGB in all crops and weeds at the moment, not just the winter cereals.

Because RGB are moving so much in the environment, and probably migrating in on storm fronts, we are likely to see ongoing infestations of crops for some time. This means that any decision to control RGB needs to be done with full knowledge that the treated crop may be infested again in a short period of time.

(Photos: RGB adult (top) and RGB late instar nymph (bottom). Keith Power)

Rutherglen bugs in wheat and barley
At this stage in the season, RGB is not going to have any impact on yield in winter cereals. Grain that is hard will not be damaged by this sucking bug.

The main issue with RGB around harvest time is contamination of harvested grain. When RGB are in very large numbers they can cause a number of issues at harvest:

Live bugs in the sample can result in rejection of a load at the delivery point
Large numbers of bugs (and bits of bugs) in the grain can elevate grain moisture. This problem is probably worst when RGB are breeding in the crop, and there are large numbers of nymphs - this is unlikely to eventuate in wheat.
There are no insecticides registered for RGB in winter cereals. If you are controlling armyworm, or helicoverpa (except with NPV) then some of these options will control RGB to some extent. But be very aware of the WHP of any insecticide used this late in the season.

Treating a crop now for RGB is no guarantee that there will not be a reinfestation before harvest

What can you do if you have large numbers of RGB at harvest?

The best approach is to try and limit the number of RGB that end up in the harvested grain. Some of the suggestions for doing this include:

Harvesting at night
Fitting screens to the header
Leave the tarp off the load for as long as possible to allow RGB to escape post harvest

If you are storing grain, RGB will not cause damage to grain in storage, and there is no need to treat grain to kill them. Large numbers of crushed RGB in harvested grain have been identified as tainting the grain with the oily exudates from their scent glands. It is unclear over what period this tainting persists.

What about RGB in other crops?

Seedling and vegetative crops
RGB is primarily a seed-feeding species, and have the capacity to damage crops during grain filling – but we know very little about how much damage in any crop other than sunflower.

In very large numbers, RGB can damage seedling crops purely by weight of numbers feeding on seedlings. In more advanced vegetative crops they will not cause any impact as long as the crop has adequate moisture and is growing actively. Be alert to RGB in sunflower at budding and flowering, and in sorghum from flowering through to soft grain; these infestations may warrant treatment.

RGB in sunflower
Whilst RGB numbers may be high in vegetative sunflower now, it is important to weigh up the decision to spray with the following:
Actively growing plants, with adequate moisture, will not be greatly impacted on by RGB feeding
Reinfestation is a real possibility, and if treating during the vegetative stage, it is likely that insect control in the crop will run to 3 sprays (vegetative, budding and flowering) if RGB pressure remains high and helicoverpa infest the crop as well.

In sunflower there are two critical periods during which RGB can cause significant crop damage:
Budding: bugs congregrate on the upper stem and bud. Bug feeding on the stem behind the head may cause the stem to wither and the bud droop.
Flowering: eggs are laid in the head and nymphs emerge in about a week and start feeding on developing seeds. Adult numbers are often minor in comparison with the size of the population once nymphs start to emerge.
Feeding on developing seeds causes yield loss, and a loss of oil content and quality of grain.

Thresholds for sunflower:

Budding: 10 bugs per head
Flowering: 20-40 bugs per head
If it is necessary to treat at flowering, do so before the heads turn down, otherwise it is difficult to get good contact with the bugs in the flower.

Control considerations in sunflower

Synthetic pyrethroids (SP) are the most effective option for controlling RGB

If RGB are in large numbers at budding and flowering, but there are a few helicoverpa present, consider an SP/NPV mix. Steward™ (for use in sunflower under permit to control helicoverpa and RGB) will provide, at best, suppression of RGB and will not provide adequate control of a large population.
The impact of insecticides on bees is an important issue in sunflower, particularly if there are hives nearby. Spraying later in the day, when bees are less active, will reduce the impact on them.

No concern for tell-tale holes

2007-11-02T09:08:00.000+10:00

Corn earworm larvae on vegetative sorghum crops produce characteristic holes in the leaves after feeding in the throat of the plant. These tell-tale signs are of no great concern as this type of feeding will not affect crop yield.

Caption: BEB alias Austin McLennan showing a characteristic holey sorghum leaf.

The recent presence of high numbers of corn earworm, Helicoverpa armigera, on winter cereals and chickpea could herald the beginning of a busy time ahead for grain sorghum in southern Queensland.

As larvae on cereal crops mature, they climb down the plant and burrow into the soil to pupate. Moths emerge from these pupae 2 to 3 weeks later, and start the next generation by laying eggs on suitable host plants.

Vegetative sorghum is attractive to egglaying moths, and larvae hatch from newly laid eggs in 3 to 4 days. Survival of larvae on vegetative crops may not be high, but vegetative sorghum can be an important intermediate host that bridges the gap between winter and summer.

Armyworm larvae may also be present in vegetative sorghum. Armyworm larvae cause sorghum plants to look ‘ragged’, but again this leaf feeding does not result in any yield loss in advanced and actively growing seedling crops.

Control of larvae on vegetative sorghum is generally not recommended as the damage is cosmetic and unlikely to affect yield.

While corn earworm larvae are advertising their presence in southern Queensland grain sorghum crops, of greatest importance are larval infestations after flowering and during grain fill.

Egglaying by corn earworm moths and larval management on sorghum heads will be the subject of a future posting.

Can you confidently identify armyworm and helicoverpa larvae in winter cereals?

2007-10-17T11:27:00.001+10:00

Both Helicoverpa armigera and armyworm larvae are occurring together in wheat and barley. It is important to be able to separate the helicoverpa larvae from the armyworm larvae in order to determine whether the numbers are above or below threshold, and, if needed, to make the most appropriate decision about control options.

Armyworm larvae

have three white stripes on the collar, behind the head. These stripes may or may not persist down the body so concentrate on the collar (see the image above at right).

skin is smooth without obvious hairs and bumps.

larger larvae tend to curl up when disturbed.

Helicoverpa larvae

skin is lumpy with obvious hairs.

may be considerable variation in colour.

may or may not have a 'saddle'.

Control considerations for Helicoverpa in chickpea

2007-10-11T15:52:00.000+10:00

The recently released brochure outlining how to calculate an economic threshold for helicoverpa in chickpea is available on the DPI & F website at www.dpi.qld.gov.au/fieldcrops. Click on the link to Helicoverpa management in chickpea where you can view or download the brochure.

Control considerations – which product when?
There is a range of products registered for helicoverpa control in chickpea. However, the use of synthetic pyrethroids is really only an option in regions where H. punctigera dominates, or where the population is predominantly made up of larvae smaller than 5 mm in length. The use of SPs against a predominantly H. armigera population is likely to deliver a poor result in terms of control. Larvin (thiodicarb) is another option, particularly where efficacy of this product in the local area is known to be high.Spinosad (Tracer II ™) and indoxacarb (Steward ™) are both effective against both H. armigera and H. punctigera. Remember that Steward has a cut-off for use in chickpea (15 September in hot areas (with an extension to 30 Sep this season); 15 October in warm areas, 30 October in cool areas). One strategy for the management of mixed age populations of helicoverpa is to use Steward™ first, if prior to cut-off, and then one of the other products if the crop needs to be sprayed again.

Are corn earworm a problem in winter cereals?

2007-10-01T08:59:00.000+10:00

Corn earworm, Helicoverpa armigera, are frequently found in winter cereals but usually numbers are too low to warrant control. Occasionally, however, corn earworm numbers may be sufficient to cause economic damage. The high value of today’s grain is further reason to carefully check for grub infestations.

It is not unusual to find both corn earworm and armyworm in cereal crops. Correct identification of the species present is very important as it influences damage potential and control measures.

Virtually all of the helicoverpa in cereals (barley, wheat, triticale, oats, canary) are the corn earworm, H. armigera. This is very relevant, because this species has developed resistance to many of the older insecticide groups that have been used to control it.

Armyworm larvae (such as common armyworm Leucania convecta) are distinguished from corn earworm larvae by the presence of three pale stripes just behind the head, and by their smooth skin, without any hairs or bumps. While corn earworm will be active on the crop day and night, most armyworm will shelter on the ground during the day and feed at night.

Corn earworm (top) and common armyworm (bottom)

Life history
Corn earworm development on winter cereals is very much like on sorghum, where moths lay eggs singly on the preflowering heads, soon after emergence from the boot leaf. This results in relatively synchronous development stages in the crop, depending on moth flights, weather conditions and the spread of flowering in the crop. Larvae hatch from the eggs in 3 to 5 days and develop through the grain fill stage over 14 to 28 days, depending on temperature. Larvae tend to graze on the exposed tips of developing grains. Rather than totally consuming a low number or whole grains, they graze on a larger number of grains, thus increasing the potential losses. Most of the feeding will be during the final two instars. When mature, larvae drop to the ground and pupate in earthen cells. Moths will emerge 2 to 3 weeks later and start the cycle again.

Corn earworm do not cause the typical head-cutting of armyworms as seen in ripening barley crops.

Damage
How much damage do larvae cause to cereals? There are currently no data from cereals on which to base this decision, but in the past extrapolation from the old sorghum damage value (1.5 g grain loss per larvae) has been used as a guide.

To put this corn earworm damage value into perspective, there are some comparative data for other crops. The damage value for sorghum was recently revised upwards from 1.5 to 2.4 g per larva. The value for chickpea is 2.0 g per larvae, and the latest mungbean value is 3.5 g per larva. Using the old sorghum value (1.5 g per larva) is not unreasonable and may be on the conservative (low) side, but it provides sound guidance for decision-making.

Based on the preceding information, we can make an estimate of what represents a problem. One larvae per m2 potentially causes 15 kg grain loss/ha (based on the figure of 1.5 g/larva eaten). At $350/tonne, this loss equals $5.25/ha. For this example, the break even point where cost of control ($28.50/ha) equals potential lost grain is 5.4 larvae/m2 (28.50÷5.25). This is based on the cost of control being $28.50/ha (methomyl at 1.5 L/ha ($16.50) plus aerial application ($12.00/ha)).

Any of these parameters can be varied to suit individual costs, and can incorporate a working benefit:cost ratio. A benefit:cost ratio of 1.5 is common and means that the projected economic benefit of the spray will be 1.5 times the cost of that spray. Spraying at the break even point (benefit:cost ratio of 1) is not recommended.

It should also be remembered that larval damage is irrespective of yield potential of the crop i.e. each larva will eat its fill whether it is 1 tonne/ha crop or a 3 tonne/ha crop.

Management
In many cases corn earworm larvae are not identified in cereals until they are medium or large in size i.e. >7 mm in length. This has implications for their management. Because corn earworm have historically had high resistance to pyrethroids, a pyrethroid is unlikely to provide satisfactory control, particularly if larvae are greater than 3 to 5 mm in length. While resistance levels to pyrethroids may have declined in recent years, control of medium-large corn earworm larvae using pyrethroids is not recommended.

Methomyl is another registered option at 1.5 to 2.0 L/ha, with the higher rate against larvae greater than 20 mm in length. It has contact action only (no residual), but that is not a problem because reinfestation is most unlikely. Resistance to carbamates (methomyl) has been a problem in the past, so any decision to use this product should be based on its recent performance against pest infestations.

In situations where both corn earworm and armyworm are present, carefully assess the relative abundance of each. Head-cutting activities of large armyworm larvae in ripening barley crops can be very serious and require prompt action. While registered pyrethroids may be a preferred option for armyworm, they are unlikely to have much impact on corn earworm larvae. Methomyl is also registered for both pests in wheat and barley. As armyworm mostly feed at night, spraying at dusk is recommended for best results.

Always read the label and abide by relevant withholding periods and export grazing/slaughter intervals where feeding to stock may be involved.

Many natural enemies (predators, parasites and pathogens) attack corn earworm larvae. Where winter cereals have been previously treated with broad spectrum insecticides to control aphids, fewer natural enemies may be present and survival of caterpillar pests could be greater than normal.

Further information
Understanding Helicoverpa ecology and biology in southern Queensland: Know the enemy to manage it better. http://www2.dpi.qld.gov.au/fieldcrops/17696.html

WATCH FOR ARMYWORMS IN BARLEY AND OATS

2007-09-21T08:38:00.000+10:00

With the current high value of barley, growers should closely monitor armyworm infestations as crops approach maturity. Armyworms are important pests in southern Queensland where they attack winter cereals, particularly barley and oats, in September and October. Larvae appear in plague proportions in some years, and are patchy in others. Head cutting by large larvae can lead to serious losses in barley.

Description and Life History

The common armyworm moths are light brown-reddish. Wings are speckled with black and a have a white spot in the centre. Moths have a wingspan of 30 to 40 mm. Moths become active in spring, sometimes moving long distances on suitable winds from inland areas where they breed on grasses, to more easterly cropping areas. Female moths lay small, white spherical eggs in irregularly-shaped masses in leaf litter, on dead leaves at the base of the plants, in folded blades or under the sheaths of the upper leaves. Eggs hatch in as little as 3 to 4 days depending on temperature.

Larvae can grow up to 35 mm in length with conspicuous white, pink and brown stripes running the whole length of the body. Larvae are distinguished from similar larvae (helicoverpa, cutworm) by the presence of three pale stripes just behind the head, and by their smooth skin, without any hairs or bumps. Larvae mostly feed at night and shelter on the ground during the day. Larvae consume about 90% of their total food intake in the last larval stage (20-35 mm). Mature larvae leave the plant and burrow below the soil surface, where they transform into pupae in earthen cells. Moths subsequently emerge from these pupae.

Damage
During the vegetative growth phase, plants can tolerate considerable leaf feeding. Leaves may look tattered from the eaten-out leaf margins. Faecal pellets around the base of plants are another indication of armyworm infestation. Armyworm generally do not require control during the vegetative stage.

The most serious armyworm damage in cereal crops occurs when larvae feed on the upper flag leaf and stem node as the crop matures. Larvae target the stem node as the leaves become dry and unpalatable, and the stem is often the last part of the plant to dry. Head cutting begins at this time. One large larva can sever up to seven heads of barley a day. One larva a square metre can cause a loss of 70 kg/ha grain per day. A larva takes around 8-10 days to develop through the final, most damaging instars, so the crop is susceptible to maximum damage for this period. The current high value of barley (over $400/tonne) would suggest keeping a close watch on armyworm infestations in maturing crops.

Armyworm populations in ripening crops in excess of 1 large larva per square metre will usually warrant spraying. For insecticide application to be economic, check or scout the crop and assess the problem before head cutting starts. Check for larvae on the plant and in the soil litter under the plant. Late in the day, when the larvae are becoming active, use a sweep net (or swing a bucket through the crop) to make a quick assessment of whether armyworm larvae are present in the crop. Infestations are often patchy, so check a number of sites across the field.

Early recognition
It is essential to recognise the problem early and be prepared to spray when economic damage is imminent. A barley crop can be almost destroyed by armyworm in just a few days. Whilst large larvae do the head lopping, controlling smaller larvae that are still leaf feeding may be more achievable.

Control
Many chemicals will control armyworms. However their effectiveness is often dependent on good penetration into the crop to get contact with the caterpillars. Control may be more difficult in high-yielding thick canopy crops, particularly when larvae are resting under leaf litter at the base of plants. As larvae are most active at night, spraying in the afternoon or evening may produce the best results. If applying sprays close to harvest, be aware of relevant Withholding Periods. Always read the label.

Biological control agents may be important in some years. These include parasitic flies and wasps, predatory beetles and diseases.

Further information:
Common, northern and sugar-cane armyworms in pasture, and winter cereals, maize and sorghum. http://www2.dpi.qld.gov.au/beef/3250.html

Cereal Aphid Update

2007-09-04T16:42:00.000+10:00

Making decisions about control of Cereal Aphids

This post is an update on cereal aphid management following a number of enquiries from growers and agronomists over the last week or so.

Which species in crops?
There are two species of aphid you are most likely to encounter in winter cereals (oats, wheat and barley). They are the oat aphid (Rhopalosiphum padi) and the corn aphid (R. maidis). The oat aphid is found around the base of the tillers, and the corn aphid in the whorl and under leaves higher on the plant. Both aphid species are greenish to black with rusty red-purple areas on the rear end around the ‘tail’.

(Line drawings from DPI Victoria's Insectopedia)
See the Northern Region Ute guide for more detailed descriptions and pictures – or see August 23 posting on the Beatsheet Blog

How much damage can aphids cause?
The key question about aphids is “will the population of aphids in my crop cause damage to the crop, and yield loss?”

Direct aphid damage, as a result of feeding, is difficult to detect. In moisture stressed crops you may see yellowing with high aphid populations. Otherwise, there are generally no early signs of how much impact the aphids are having on the crop.

West Australian research showed yield losses of up to 10%, and reduction in seed size, with aphid infestations (this was without any impact of barley yellow dwarf virus).

Overseas research (Canada, US) suggests that significant yield loss occurs when aphids are present from flowering through to milky grain. The data also suggests that yield loss does not occur when infestations are present earlier or later than this period.

Making control decisions

Corn aphids may disappear by themselves
Corn aphids, the species that lives in the whorl, generally disappears when the crop comes into head. This is because their preferred site is no longer there, and they tend not to survive as well on leaves. If you have the corn aphid in crops, consider delaying a control decision until the crop starts to head.

Control thresholds
Qld and WA threshold recommendations are comparable, with the WA recommendations based on the most recent research that has been undertaken in Australia.

Recommendation are to check crops regularly from late tillering, and consider control if the aphid population exceeds 15 aphids/tiller on 50% of tillers.

http://www.agric.wa.gov.au/content/pw/ph/dis/cer/bydv_aphidfeeding.htm

Other considerations when making a decision about cereal aphids

Natural enemies (lady beetles, hoverflies, parasitic wasps) can have a big impact on aphid populations, reducing them to very low levels in many instances.
Dimethoate and synthetic pyrethroids (registered for cereal aphid control) are highly disruptive to natural enemies. The application of these insecticides early (e.g. during the vegetative and early tillering stages) may result in a later reinfestation of the crop because small numbers of surviving aphids are no longer controlled by natural enemies. The impact of these products on natural enemies can persist for some days.
Pirimicarb (Pirimor®) is a soft option for cereal aphid control, compare its price with that of dimethoate when making a decision – it may be competitive. Pirimicarb has some systemic activity.
Oat aphids, at the base of the plant, can be difficult to contact in a dense crop. Dimethoate will kill aphids by contact, but its systemic activity is by translocation upwards, so its efficacy against oat aphid is unclear.
The systemic activity of both pirimicarb and dimethoate may be reduced in moisture stressed crops.
Rain will reduce aphid populations by knocking/washing individuals of plants, and the aphids tend not to get back on the plants. Often ground predators, like carabid beetles, ants etc will eat aphids on the ground. Check populations of aphids again if you get more than 20 mm rain.

Are aphids sucking away cereal profits?

2007-08-23T12:04:00.000+10:00

Aphid control decisions tend to be more problematic in moisture stressed winter cereal crops, since in a well supplied crop the level of moisture extracted from the crop by aphids is of little concern. However, in dry times every drop seems precious.

Had we not received the recent rain over the last few days throughout southern Queensland and northern NSW grain growing areas, we could have expected enquiries about aphids in stressed winter cereal crops. But even so, the crops aren’t finished yet, and so it’s timely to be reminded of what the different aphid pests are in winter cereals and the principles for managing them.

Aphid species
Four different species of aphid commonly attack barley and wheat in Queensland. They all prefer barley more than wheat. Aphids suck sap from the plants. Under heavy infestations plants may turn yellow, be stunted and appear generally unthrifty.

Oat or Wheat Aphid (Rhopalosiphum padi)
The oat aphid is brown to muddy green with rusty red patches at the base of the tubes at the rear end of the body. It normally occupies the base and lower portions of the plant. This is generally the most common aphid attacking winter cereals.

Corn Aphid (Rhopalosiphum maidis)
The corn aphid is green to dark olive-green with a purplish area at the base of the tubes at the rear end of the body. It normally lives on the tops of the plants particularly within the rolled up terminal leaf.

Rose-grain Aphid (Metopolophium dirhodum)
The rose-grain aphid is pale green with a darker green stripe along the middle of its back. It normally occupies the undersides of the leaves. It colonizes the lower leaves and moves up the plant as leaves senesce.

Rice Root Aphid (Rhopalosiphum rufiabdominalis)
The rice root aphid is a honey-brown colour with a rusty red area at the base of the tubes at the rear end of the body. It normally occupies the roots of the plants under the soil surface.

Making decisions
The decision as to whether controlling aphids on winter cereals will provide an economic return is often complex, and is not just dependent on the size of the aphid population.

Several other factors could influence the control decision. Aphids are more readily controlled in seedling and pre-tillering crops which are less bulky than post tillering crops. Aphids tend to disappear as crops come into head.

Prolonged infestation of moisture stressed crops can exacerbate the effect of moisture stress. Yield potential, value of grain and cost of control are important considerations, but anecdotal evidence suggests that direct feeding by very large numbers of aphids is needed to impact on yield.

Natural enemies (ladybird beetles, ladybird larvae, hover fly larvae, lacewing larvae or parasitic wasps) can exert effective control on small to moderate aphid infestations.

Photo: Larva of hover fly in an aphid colony.

A general recommendation is to check for aphids by choosing six widely-spaced positions in the crop and at each position examine five consecutive plants in a row. If 27 out of 30 plants are covered with aphids and if there are less than two natural enemies per plant on each of the infested plants, then consider treatment. Delay any planned chemical control if rain is forecast and check again after rain.

Dimethoate at 500 mL/ha of 400 g/L product is the recommended chemical control. It has a withholding period of 28 days for harvest and one day for grazing.

Dimethoate has a contact action but is also a systemic insecticide taken up through the leaf and then translocated through the upper portion of the plant. Aphids are subsequently controlled when they feed on the plant.

The rice root aphid feeds below ground and can’t be effectively controlled by spraying.

Dimethoate may be tank-mixed with certain broadleaf herbicides. Check the label before use. Also check water quality as high pH can affect performance of dimethoate.

Dimethoate will kill natural enemies, increasing the possibility of subsequent aphid infestations later in the season.

St George growers meet to discuss area-wide SLW management

2007-08-16T16:31:00.000+10:00

On Friday August 10, St George growers and agronomists met to discuss strategies to manage the Silverleaf whitefly population in the irrigation area.
Over the past two seasons, a number of cotton fields in St George have been treated for SLW. Last year more fields were treated earlier, and required a second treatment. It is clear that SLW is now part of the insect pest complex in St George, and it is important that St George growers take the same area-wide approach to managing SLW as has the Emerald irrigation area.
Richard Sequeira (DPI&F entomologist, Emerald) discussed the role of cotton as the main driver of the local SLW population. Tactics discussed that can contribute to minimising SLW outbreaks include narrowing the planting window as much as possible, avoiding the use of broadspectrum insecticides early in the season for other pests (OPs and SPs for mirids and helicoverpa) and the correct timing of SLW control.
Melina Miles (DPI&F entomologist, Toowoomba) discussed the survey data from 06-07 that shows the rate of population growth in St George is very similar to that in Emerald. In contrast, data from the Downs shows that growth is significantly slower and the risk of outbreak on the Downs is low in most seasons. Overall parasitism of SLW was very low in 06-07.
The development and implementation of tactics to manage SLW will be an ongoing process for growers and agronomists in this area. For more details contact Dallas King (Regional Extension Officer, St George 0427 635 621) or Richard Sequeira (Emerald, 49837410).

A better way to make decisions about Helicoverpa control in chickpea

2007-08-16T14:09:00.000+10:00

A new guide to calculating economic thresholds available now
Growers and agronomists are now able to calculate how much yield they will lose if they do not control an infestation of helicoverpa in chickpea. They can calculate the point at which it is economic to control an infestation based on the potential yield loss, their costs of control, the expected grain price and their own preference for benefit:cost.

Being able to calculate the economic thresholds based on an individual grower’s situation is a significant advance on the old recommendations which were fixed at 1-4 larvae per square metre, and did not allow for adjustment when there were changes in grain price or costs of control.

The calculation is now possible following several seasons of research by Melina Miles and Richard Lloyd in Toowoomba, and Paul Grundy and Sherree Short in Biloela. The DPI&F entomologists have worked out that one helicoverpa larva, surviving from hatchling to pupa, will consume 2 grams of chickpea grain. This information is the basis on which potential crop loss ($/ha) is calculated.

The research also examined the impact of spray timing on yield and found that no yield loss occurred when larvae were allowed to feed in vegetative and flowering crops. Yield loss only occurred when larvae fed on developing, filling and maturing Many hours observing larval feeding in the field supported this result. Observations found that larvae of all sizes fed readily on leaves, large larvae did the majority of feeding on pods, and that neither small nor large larvae fed on buds and flowers. However, there are seasons when larval pressure in vegetative and flowering crops may necessitate control.

Grain quality was assessed in both threshold and spray timing trials, and no loss in quality (increase in splits or damaged grain) was seen within the range of densities that it was economic to spray.
In summary, the recommendations for helicoverpa management in chickpea are:

Use a beatsheet to sample helicoverpa in chickpea.
Exclude very small larvae from threshold calculations – they are difficult to assess accurately in the field – they will be counted as smalls at the next count.

Adjust the number of small larvae for natural loss from disease, predation etc (30% loss)

Calculate potential crop loss ($/ha) using the equation.
Unless pressure is extreme, delay any control until crops are setting, filling or maturing pods. It may be necessary to apply control during flowering if targeting small larvae that will be medium-large by pod set.

Details of the research, and calculations of economic thresholds are in the new brochure – get yourself a copy!

This research was supported by GRDC and DPI&F.

Get a copy of new Helicoverpa Management in Chickpea brochure from the DPI&F Call Centre, phone 13 25 23. Will soon be downloadable from the web.

IPM Researchers Forum 2007

2007-08-02T11:46:00.000+10:00

About 50 devoted enthusiasts of Integrated Pest Management (IPM) gathered in Toowoomba on 24-25 July 2007 for the annual Northern Farming Systems IPM Researchers Forum.

Participants in the forum were from industry, research, extension and stakeholders. The aims of the IPM Forum were to exchange the latest information about research and projects and to maintain linkages between IPM projects and people across industries.

The Forum mostly focused on R,D & E being conducted in NSW and Qld, in the broadacre farming system. The broad range of topics covered in presentations included some key pests – green mirids, stink bugs, helicoverpa and silverleaf whitefly.

Of particular interest were presentations by a number of postgraduates from The University of Queensland.

James Hereward - Dispersal and host plant interactions of green mirids (or ‘Where do mirids come from – are we dealing with locals or immigrants from far inland?’)

Corinna Lange - The population genetic structure of native budworm, Helicoverpa punctigera, in the Australian landscape (or ‘My trip to pick grubs off wildlowers in the middle of Australia’)

Jamie Hopkinson - Host acceptance behaviour of an aphid parasitoid (or ‘How I learnt to dissect tiny aphids and find even tinier wasp eggs inside them’).

It was interesting that many IPM Forum participants were noticeably envious of Corinna’s work – but less so of Jamie’s!

Other welcome participants included the Grains Research and Development Corporation Crop Protection Program Team who seized the opportunity to meet with project leaders and discuss research results.

If you have any questions about the 2007 IPM Forum, leave a post below.

Until next time, Big Bug.

Photo: The GRDC Crop Protection Program Team includes (L to R) John Sandow, GRDC Crop Protection Manager, Ralph Burnett, Western Panel member, Allan Mayfield, Southern Panel member, Rohan Rainbow, GRDC Project Manager, and Bill Yates, Northern Panel member. Photo provided by GRDC.

The Beat Sheet is now a blog

2007-07-11T14:55:00.000+10:00

Hi, and welcome to the first online edition of The Beat Sheet - a blog focussed on integrated pest management news relevant to Australia's northern grain region.

Until now, The Beat Sheet has simply been a newsletter circulated via email and snail mail (going back further, its orginal name was the Heliothis Hotline). While the newsletter format will continue for the time being, we are excited about the potential for this move into the world of blogs and blogging.

So who are we? The team looking after this blog is made up of the the following Queensland Department of Primary Industries and Fisheries entomology staff.

Big Eyed Bug (a.k.a. Austin McLennan, Development Extension Officer - IPM)
Big Bug (a.k.a. Dr Dave Murray, Principal Entomologist)
Lady Bug (a.k.a. Dr Melina Miles)

To all our longstanding readers of the newsletter format, we're confident you'll find The Beat Sheet blog is a much more convenient and timely way to receive updates on insect pest management issues relevant to the grain cropping regions of Queensland and northern NSW.

To both new and old readers, if you have any ideas about how to improve The Beat Sheet, please let us know by posting your comments to this blog.

Welcome and Enjoy!