Organised by commodities – such as cereals, sugar and tropical pasture legumes – it examines all the pest species for a particular commodity across Australia. Identification, distribution, damage, host range, biology, risk period and monitoring techniques are described for each entry, accompanied by useful illustrations. The book also describes introduced biological control agents that effectively control crop pests. Pests of Field Crops and Pastures will be a useful tool in crop management for progressive farmers, agronomists, agricultural consultants and academics alike.
PESTS OF FIELD CROPS AND PASTURES
This comprehensive handbook on economic entomology for Australian field crops and pastures is the first of its kind. It encompasses pest and beneficial insects as well as allied forms of importance in Australian agriculture.
PESTS OF FIELD CROPS ANDPASTURES IDENTIFICATION AND CONTROL
EDITOR: P.T. BAILEY
EDITOR: P.T. BAILEY
PESTS OF FIELD CROPS ANDPASTURES IDENTIFICATION AND CONTROL
EDITOR: P.T. BAILEY
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© Peter Bailey 2007 All rights reserved. Except under the conditions described in the Australian Copyright Act 1968 and subsequent amendments, no part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, duplicating or otherwise, without the prior permission of the copyright owner. Contact CSIRO PUBLISHING for all permission requests. National Library of Australia Cataloguing-in-Publication entry Pests of field crops and pastures: identification and control. Bibliography. Includes index. ISBN 9780643067585. 1. Agricultural pests – Australia – Identification. 2. Agricultural pests – Control – Australia. 3. Field crops – Diseases and pests – Australia – Identification. 4. Field crops – Diseases and pests – Control – Australia. 5. Pastures – Diseases and pests – Australia – Identification. 6. Pastures – Diseases and pests – Control – Australia. I. Bailey, Peter. 632.60994 Published by CSIRO PUBLISHING 150 Oxford Street (PO Box 1139) Collingwood VIC 3066 Australia Telephone: +61 3 9662 7666 Local call: 1300 788 000 (Australia only) Fax: +61 3 9662 7555 Email:
[email protected] Web site: www.publish.csiro.au Front cover Left: Larva of etiella moth in a rattlepod (DPI&F Qld: Joe Wessels) Right: Predatory shield bug preying on a nymph of green vegetable bug (DPI&F Qld: Kristen Knight) Back cover Left: Adult orange caterpillar parasite (Keith Power) Right: Adult tachinid fly preparing to lay an egg on a caterpillar (DPI&F Qld) Set in 10/13 Adobe Palatino and Optima Cover design by The Modern Art Production Group Text design by James Kelly Typeset by Desktop Concepts Pty Ltd, Melbourne Printed in Malaysia by Imago
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CONTRIBUTORS
P.G. Allsopp, BSES Limited, Indooroopilly, Qld P.T. Bailey, University of Adelaide, Adelaide, SA G.J. Baker, South Australian Research & Development Institute, Adelaide, SA F.A. Berlandier, formerly Department of Agriculture and Food, Western Australia, Perth, WA H. Brier, Queensland Department of Primary Industries and Fisheries, Kingaroy, Qld D.T. Briese, CSIRO Entomology, Canberra, ACT K.J. Chandler, BSES Limited, Bundaberg, Qld S. De Faveri, Queensland Department of Primary Industries and Fisheries, Mareeba, Qld S. Deutscher, CSIRO Plant Industry, Narrabri, NSW P.B. Edwards, Maleny, Qld R.J. Elder, formerly Queensland Department of Primary Industries, Rockhampton, Qld R.N. Emery, Department of Agriculture and Food, Western Australia, Perth, WA G.P. Fitt, CSIRO Entomology, Brisbane, Qld B. Franzmann, Queensland Department of Primary Industries and Fisheries, Toowoomba, Qld G. Goodyer, Dundas, NSW L. Hill, Tasmanian Department of Primary Industry, Water and Environment, Devonport, Tas. W. Hawthorne, Pulse Australia, Naracoorte, SA D.C. Hopkins, South Australian Research & Development Institute, Adelaide, SA D. Hunter, Australian Plague Locust Commission, Canberra, ACT J. Ireson, Tasmanian Institute of Agricultural Research, New Town, Tas. M. Khan, Queensland Department of Primary Industries and Fisheries, Kingaroy, Qld G. McDonald, Victorian Department of Primary Industries, Parkville, Vic. R.E. McFadyen, CRC for Australian Weed Management, Brisbane, Qld M.B. Malipatil, Victorian Department of Primary Industries, Knoxfield, Vic. R.K. Mensah, New South Wales Department of Primary Industries, Australian Cotton Research Institute, Narrabri, NSW M. Miles, Queensland Department of Primary Industries and Fisheries, Toowoomba, Qld D. Murray, Queensland Department of Primary Industries and Fisheries, Toowoomba, Qld M.K. Nayak, Queensland Department of Primary Industries and Fisheries, Indooroopilly, Qld C. Pavri, formerly CSIRO Entomology, Perth, WA B.A. Pyke, Cotton Research and Development Corporation, Narrabri, NSW J-L. Sagliocco, Victorian Department of Primary Industries M.N. Sallam, BSES Limited, Gordonvale, Qld P.R. Samson, BSES Limited, Mackay, Qld M.J. Smyth, CSIRO Entomology, Canberra, ACT M.M. Stevens, New South Wales Department of Primary Industries, Yanco, NSW A.E. Swirepik, CSIRO Entomology, Canberra, ACT P. Walker, Tasmanian Institute of Agricultural Research, University of Tasmania, Hobart, Tas. L.J. Wilson, CSIRO Plant Industry, Narrabri, NSW P.B. Yeoh, CSIRO Entomology, Wembley, WA C.J. Young, Tasmanian Department of Primary Industry, Water and Environment, New Town, Tas.
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ACKNOWLEDGEMENTS
The contribution of state governments, the Commonwealth government, and statutory industry corporations in funding salaries and institutional support for entomologists contributing to this volume, is gratefully acknowledged.
Acknowledgements, text 1. Introduction: ABS data used with permission from the Australian Bureau of Statistics. 2. Cereals: G. Goodyer provided a large amount of unpublished information on NSW cereal pest management. 5. Oilseeds: G. Goodyer and L. Hill for information on NSW and Tas. crops respectively. 6. Poppies: P.J. Cotterill (GlaxoSmithKline), P. Horne (IPM Technologies), R. Kile (GlaxoSmithKline), J. Miller (Tasmanian Alkaloids ) provided advice on the manuscript. 7. Pulses—summer: J. Wessels, QDPI&F Kingaroy, took most of the excellent photographs in this chapter and reared many of the insects illustrated to obtain images of their life stages. Margaret Schneider, University of Queensland, provided copies of photographs taken by Merle Shepard and previously published in the book Insects on Grain Legumes in Northern Australia by Merle Shepard, Bob Lawn and Margaret Schneider. For advice on the manuscript: Pat Collins. DPI&F Qld, Indooroopilly, Ian Crosthwaite, Bean Growers Australia, Paul DeBarro, CSIRO Brisbane, John Donaldson and John Duff, QDPI&F Indooroopilly, Alan Garside, Pat Harden, the Peanut Company of Australia, Megan Hoskings, John Hughes, DPI&F Qld, Mackay, Moazzem Khan, DPI&F Qld Kingaroy, Kristen Knight, formerly QDPI&F, Melina Miles, QDPI&F Toowoomba, Dave Murray, Nat Parker and Jamie Hopkinson, DPI&F Qld Toowoomba, Richard Sequeira, DPI&F Qld Emerald, Brad Scholz, formerly DPI&F Qld Toowoomba. 10. Sorghum: G. Goodyer provided information on NSW crops. 12. Tobacco: G. Baxter, VDPI Myrtleford and P. Tonello DPI&F Qld, respectively, provided information on Victorian and Queensland agronomy. 13. Pastures—summer rainfall: Entomology staff from the Queensland Department of Primary Industries and Fisheries, the Australian Plague Locust Commission and BSES Limited provided advice on a number of the pasture insects described. 15. Pastures—Winter: G. Goodyer, R.J. Hardy, K Henry, J. Matthiessen S. Micic and J. Ridsdill-Smith contributed to the draft chapter.
Acknowledgements, illustrations Illustrations are reproduced by permission of: Agrisearch Limited, New Zealand Andrew Austin, South Australia Australian Bureau of Meteorology Australian Bureau of Statistics Hugh Brier BSES Limited, Queensland Robin Coles, South Australia
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ACKNOWLEDGEMENTS
Commonwealth Bureau of Meteorology CSIRO Division of Entomology CSIRO Division of Plant Industry Department of Agriculture, Western Australia Department of Natural Resources & Mines, Queensland Department of Primary Industries and Fisheries, New South Wales Department of Primary Industries and Fisheries, Queensland Department of Primary Industries, Victoria Department of Primary Industries and Water, Tasmania W.E. Frost, Agrisearch Services Pty Ltd, Adelaide, South Australia GlaxoSmithKline, Tasmania Graphic Science, Dennis Crawford: www.graphicscience.com.au Megan Hoskins, Darwin John Hughes, Queensland Mike Keller, South Australia Peter and Jan Larsen, Queensland Bob Lewis, Canberra Cheryl Mares, Queensland Natalie Moore, New South Wales Keith Power, Queensland Pacific Seeds, Queensland Primary Industries & Resources South Australia Pulse Australia, South Australia John Randles, South Australia David Rentz, Mareeba, Queensland Merle Shepard, Queensland South Australian Research & Development Institute Winfield Stirling, USA Tasmanian Alkaloids, Tasmania Tasmanian Institute of Agricultural Research Jozef Wessels, Queensland John Wightman, Queensland
v
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ABBREVIATIONS
ACT: Australian Capital Territory AFRS: Alan Fletcher Research Station (Queensland Government) APLC: Australian Plague Locust Commission APVMA: Australian Pesticides and Veterinary Medicines Authority BSES: BSES Limited CSIRO: Commonwealth Scientific and Industrial Research Organisation DAFWA: Department of Agriculture and Food, Western Australia DPI&F Qld: Department of Primary Industries and Fisheries, Queensland DPI&W Tas: Department of Primary Industry, Water and Environment, Tasmania NSW: New South Wales NSWDPI: New South Wales Department of Primary Industries NT: Northern Territory NTDPI: Northern Territory Department of Primary Industries PIRSA: Primary Industries & Resources South Australia Qld: Queensland QNRM: Queensland Department of Natural Resources and Mines SA: South Australia SARDI: South Australian Research & Development Institute Tas.: Tasmania TIAR: Tasmanian Institute of Agricultural Research UnivAd: The University of Adelaide VDPI: Department of Primary Industries, Victoria Vic.: Victoria WA: Western Australia
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CONTENTS
1. 2.
3.
4. 5.
INTRODUCTION
1
CEREALS
5
Cereals—pests and beneficials in the field D.C. Hopkins and G. McDonald Pests of stored grains R.N. Emery and M.K. Nayak
5 40
COTTON
63
Pests L.J. Wilson, G.P. Fitt, S. Deutscher, M. Kahn and B.A. Pyke Beneficials R.K. Mensah and B.A. Pyke
63 102
MAIZE
121
D.A.H. Murray
OILSEEDS
135
Winter oilseeds F.A. Berlandier and G.J. Baker Summer oilseeds B.A. Franzmann
135 155
6.
POPPIES L. Hill
163
7.
PULSES—SUMMER (INCLUDING PEANUTS) H. Brier
169
8.
PULSES—WINTER
259
9.
RICE
M.M. Miles, G.J. Baker and W. Hawthorne
M.M. Stevens
279
10. SORGHUM B.A. Franzmann
297
11. SUGARCANE
305
12. TOBACCO
M.N. Sallam, P.G. Allsopp, K.J. Chandler and P.R. Samson
S.G. De Faveri and M.B. Malipatil
343
13. PASTURES—SUMMER RAINFALL R.J. Elder
355
14. PASTURES—SUMMER RAINFALL: WEED BIOCONTROL AGENTS R.E. McFadyen
371
15. PASTURES (INCLUDING LUCERNE)—WINTER RAINFALL
393
Grass pastures and turf C. Pavri and C.J. Young Legume pastures C. Pavri Lucerne P.T. Bailey and G. Goodyer
16. PASTURES—WINTER RAINFALL: WEED BIOCONTROL AGENTS D.T. Briese,
393 412 425
441
J. Ireson, J-L. Sagliocco, M.J. Smyth, A.E. Swirepik and P.B. Yeoh
17. PASTURES—DUNG BEETLES P.B. Edwards and C. Pavri
471
18. LOCUSTS AND GRASSHOPPERS OF PASTURES AND RANGELANDS
485
P. Walker, D. Hunter and R. Elder
GLOSSARY
505
INDEX
513
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1 INTRODUCTION This book aims to provide a comprehensive presentation of the main pest and beneficial species of insects and allied forms in the main field crops and pastures presently grown in Australia. ‘Allied forms’ are other arthropods and molluscs: invertebrates that are normally managed by similar methods as those of insects. ‘Beneficial species’ included are those predators and parasitoids which are known, or thought, to provide a level of control of pest species in crops and pastures. Also included are insects introduced to Australia for the biological control of weeds, and dung beetles introduced for the dual purposes of reducing bovine dung and the bush flies that breed in it. Each of the main crops and pasture regions in Australia is presented as a separate chapter in which the identification, biology and management of pest and beneficial species are detailed. ‘Field crops’ are those grown under extensive cultivation, as distinct from horticultural crops that require more intensive cultivation methods. To keep this book to a reasonable size, it has been necessary to arbitrarily exclude crops such as potatoes, tomatoes and other vegetable crops which, although often grown in extensive field plantings, are usually regarded as horticultural crops. Included are pests from areas where production of some crops has recently ceased. These crops include tropical rice and tobacco, and summer cotton grown in the Ord irrigation area of WA. Inclusion is justified on the basis that the considerable pest management information developed for these crops may be useful if these crops are again produced in tropical areas. This book aims to present the best available knowledge on the identification, biology and management of crop and pasture pest and beneficial species. The authors are field biologists with experience in particular field crops and
030701•Pests of Field Crops and 1 1
pastures. Some of the information presented in this book has been published in mainstream scientific books and journals, but most can be found in industry or regional publications such as ute guides, fact sheets and newsletters, or in the filing cabinets of government entomologists. As such, the basis of the information presented ranges from well-founded laboratory experiments and field trials to field observations and informed guesses by experienced field biologists. In practice, there may be little basis for preferring one over the other, especially in estimating ‘action levels’, choice of monitoring techniques or pest management techniques. The criterion for inclusion is what works best in the field.
Crop, pasture and rainfall regions The main cropping and pastoral areas of Australia are shown in Figure 1.1 and the main rainfall divisions referred to in this book are shown in Figure 1.2. Indicative crop areas quoted in this book are 5-year averages to 2002/2003 (Australian Bureau of Agricultural and Resource Economics 2005).
Layout of the book In the chapters on insects of annual crops, the order of presentation of the insects is firstly related to the phenological stage of the crop in which they are first likely to cause damage (or in the case of beneficials, to exert control). Secondly, within each growth stage, insects are presented by taxonomic groups (all aphid species, for example, are presented together) in the order that they are presented in The Insects of Australia (CSIRO 1991). In the chapters on stored products insects and pasture insects, order of presentation is by taxonomic group alone. For major pests and those with a widespread distribution, sufficient biological background is provided to enable rational decisions on their
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P E S T S O F F I E L D C RO P S A N D PA S T U R E S
Darwin
NT QLD WA Brisbane
SA NSW
Perth Adelaide
VIC
ACT
Sydney
Canberra
Melbourne
Cattle pastures Cattle and sheep pastures Dairy pastures
TAS
Sugar cane Hobart
Dryland crops Fig. 1.1. Summer and winter rainfall areas of Australia. (Australian Bureau of Statistics)
management. For insects that occur in a number of crops, a full biological background is given under the crop in which the insect causes most economic impact, while crop-specific information is provided in other crops in which it occurs. The format for a full entry is: Names of insects are generally consistent with those of the CSIRO Australian Insect Common Names website.
Minor: Damage may result in some economic loss. Widespread: Occurs as a pest (or beneficial) over most of the range where crop is grown. Restricted: Pest (or beneficial) in only part of the area where crop is grown. Regular: Likely to be a pest (or beneficial) in most years.
Distribution: Origin, present world distribution (by continent) and distribution in Australia.
Irregular: Not likely to be a pest (or beneficial) every year.
Pest (or beneficial) status: (1) Major/moderate/ minor; (2) widespread/restricted; or (3) regular/ irregular.
Identification: Size of insect. Distinctive colouration.
Major: Failure to manage likely to result in significant economic loss.
May be confused with: Any other organism or damage symptom with which the subject may be confused on the particular commodity.
Moderate: Failure to manage likely to result in some economic loss.
Host range: Plants or insects (in the case of beneficials) eaten.
2
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INTRODUCTION
Summer rainfall
Winter rainfall
Arid
Uniform
Fig. 1.2. Major Australian rainfall divisions. (Bureau of Meteorology, redrawn)
Life cycle on commodity: Number of generations each year, and number spent on the commodity. Risk period: Stage of the crop and/or month(s) when the insect causes damage. (Beneficials: when they are effective.)
Chemical control*: Availability (including registration status) of chemicals is indicated, and whether application is cost-effective at the action level indicated. ‘Cost-effectiveness’ is an estimate based on: • value of the crop
Damage: Symptoms and economic impact. (Beneficials: impact on pest.)
• cost of the chemical, including application cost
Monitoring: An indicative guide on techniques to detect significant populations of the species.
• effectiveness of the treatment
Action level: An estimate of the numbers, density, etc., at which it is cost-effective to apply control. Precision of estimates is likely to vary considerably between pest species and crops. The action level also depends on costs and the economic benefits of treatment. Action levels are provided as a guide only.
• ability of the crop to tolerate pest damage and still yield well • prevalence and effectiveness of natural enemies. No specific chemicals or chemical groups are mentioned other than to indicate chemicals to which resistance is recorded. 3
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P E S T S O F F I E L D C RO P S A N D PA S T U R E S
Cultural control: Includes cultural techniques, resistant varieties, and other techniques to avoid damage. May be omitted if no cultural control methods are in general use. Conservation of natural enemies: An attempt has been made to include only those natural enemies that are effective in control on the particular commodity and whose conservation is likely to be of economic benefit. Since there is often a paucity of information to aid this judgement, natural enemies have generally been included where possible. This category may be omitted if little is known of the natural enemies of a particular pest.
*A note on registration and usage of agricultural chemicals in Australia Registration of agricultural chemicals in Australia is the responsibility of the Commonwealth Government, presently through the agency of the Australian Pesticides and Veterinary Medicines Authority (APVMA). To be registered, an agricultural chemical must satisfy public health, food safety, occupational health and safety, environmental, trade and efficacy criteria. Regulation of usage of agricultural chemicals is the responsibility of State and Territory Governments who, through various acts, regulate the use, storage, disposal and compliance with label instructions, including
withholding periods. Pesticides acts between various states and territories are generally similar in objectives and content, but there are some differences. States may place additional restrictions on pesticide usage that further limit their registered use. Pesticide usage may be further regulated by non-statutory means, such as area-wide resistance management agreements between farmers. Residue requirements more restrictive than those allowed for registration may be required by national and international commodity buyers. Farmers producing for organic and integrated pest management (IPM) markets may have restrictive contractual requirements for pesticide usage.
Sources of information used throughout this book Australian Bureau of Agricultural and Resource Economics (2005). Crop Report No. 133, February 2005. Australian Bureau of Statistics (1996). Australian Agriculture and the Environment. Catalogue no. 4606.0. Figure 2.31. p. 40 CSIRO. Australian Insect Common Names.
. CSIRO (1991). The Insects of Australia. Melbourne University Press, Melbourne
4
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2 CEREALS CEREALS—PESTS AND BENEFICIALS IN THE FIELD D.C. Hopkins and G. McDonald N Barley, Hordeum vulgare Poaceae. Origin: Asia, Europe N Oats, Avena sativa Poaceae. Origin: Europe N Rye, Secale cereale Poaceae. Origin: Europe N Triticale, Triticosecale rimpaui Poaceae. Hybrid N Wheat, Triticum aestivum, and durum wheat, T. durum Poaceae. Origin: West Asia
(a) Tillering
(b) Wheat
(c) Barley
(d) Stubble
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P E S T S O F F I E L D C RO P S A N D PA S T U R E S
sowing & germination tillering stem elongation heading grain development harvest
Mar Apr May Jun Jul Aug Sep Oct Nov Dec Jan Feb Phenology of a cereal crop in the south-eastern and southwestern grain belt. Time of sowing depends on opening rains.
GROUP
PEST (major pests in bold)
Pests of establishing cereals (immediately prior to and for about 4 weeks post seeding) helical snails common white snail white Italian snail pointed snails pointed snail small pointed snail slugs black-keeled slug reticulated slug mites blue oat mites redlegged earth mite lucerne flea lucerne flea sandgroper sandgroper aphids corn aphid oat or wheat aphid rose-grain aphid beetles blackheaded pasture cockchafer white curl grubs (cockchafers) spinetailed weevil spotted vegetable weevil wireworms false wireworms caterpillars black cutworm brown cutworm common cutworm pasture webworm Pests of the vegetative growth stage of cereals (4 weeks post seeding through to head emergence) mites cereal rust mite and wheat curl mite brown wheat mite blue oat mite redlegged earth mite lucerne flea lucerne flea aphids corn aphid oat or wheat aphid rose-grain aphid
PAGE 7 9 9 10 11 11 11 12 13 13 14 16 17 17 18 20 21 21 23 23 24 25 26 28 28 11 12 13 14 16 17
6
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CEREALS
GROUP
PEST (major pests in bold)
black cutworm brown cutworm common cutworm grass anthelid Pests of mature cereals (head emergence through to harvest) locusts and grasshoppers Australian plague locust migratory locust small plague grasshopper aphids corn aphid, oat or wheat aphid, rose-grain aphid caterpillars common armyworm northern armyworm sugarcane armyworm southern armyworm inland armyworm native budworm cotton bollworm pests at harvest common white snail, pointed snail, small pointed snail, and white Italian snail (but treated pre-harvest post harvest
PAGE
caterpillars
Introduction Cereals are grown mostly as dryland crops in the rainfall belt of 250 to 600 mm in NSW, southern Qld, SA, Tas., Vic. and WA. Wheat is the predominant cereal crop (approximately 12 M ha in 2004) then barley (3.5 M ha in 2004), oats (0.85 M ha in 2004) and triticale (0.34 M ha in 2004). Approximately 70% of the wheat and barley and 25% of the oats grown are exported, while triticale and rye are used domestically. Many of the pests of cereals are introduced exotic species (e.g. mites, snails and aphids) but some are native species (e.g. soil dwelling scarab, elaterid and tenebrionid beetles) that existed in woodland and grassland habitats prior to European settlement but now still occur in these same areas that have been cleared largely for grain and pasture production. In the Australian grain belt, cereals may be rotated with oilseeds, pulses and, less commonly, pastures.
23 24 25 29 29 30 30 14–17 30 33 35 35 36 37 37 7–10
40
Distribution: European–Mediterranean Basin in origin, now in Australia (Baker 1986), where it occurs in NSW, SA, Tas., Vic. and WA but is most prevalent in SA (Hopkins et al. 2003). Pest status: Major, restricted, regular. Identification: The diameter of the shell of a mature snail ranges from 10 to 15 mm. The coiled white shell has a brown band around the spiral in some individuals while others completely lack this banding. The umbilicus is open and circular (Fig. 2.1) (Hopkins and Miles 1998). Under magnification, regular straight scratches are visible across the shell. May be confused with: The white Italian snail, Theba pisana. They can be separated by the differences in the umbilicus (Figs 2.1 and 2.3) and the scratchings/etches on the shell.
Common white or vineyard snail
Host range: Includes field crops such as wheat, barley, oats, field peas, faba beans, canola and also pastures. It feeds mainly on organic matter on the soil surface but may damage young plants. It is an important pest because it contaminates grain crops at harvest and clogs and damages harvest machinery.
Cernuella virgata (Da Costa) Eupulmonata: Hygromiidae
Life cycle on cereals: This species aestivates over summer by climbing on to crop stubble and
P E S T S O F E S TA B L I S H I N G C E R E A L S
7
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P E S T S O F F I E L D C RO P S A N D PA S T U R E S
Fig. 2.1. The shell of the common white snail (shell diameter 15 mm). Note the completely round hole (umbilicus) in the shell on the right. (SARDI: G.J. Baker)
residues, various weeds and fence posts to avoid high summer temperatures at the soil surface (Fig. 2.2). Autumn rains trigger activity down onto the ground, mating and egg-laying. Most eggs are laid in late autumn or early winter but some egg-laying continues through to early spring. Eggs hatch in about 2 weeks and immature snails grow steadily throughout the winter and spring before they aestivate through the next summer. It can have an annual or biennial life cycle. Risk period: Cereals, particularly barley, and pulses and canola are susceptible to direct feeding damage shortly after crop emergence in the autumn. The risk of machinery clogging and grain contamination occurs at harvest. Damage: Moderate to high densities of snails can cause serious defoliation of emerging barley, pulse and canola crops to the point where re-sowing of the worst-affected areas is necessary. At harvest, snails can clog and damage harvest machinery and cause frustrating delays in the harvest period. They can also contaminate harvested grain samples and lead to the downgrading of grain classification, or total rejection of the contaminated load at the point of delivery. Monitoring: Successful management depends on regular monitoring of snail numbers across the whole farm. Square sampling quadrats (30 × 30 cm) can be placed on the ground, and snails counted and usually converted to numbers of snails per square metre. It is important to count only live snails; the shells of dead snails persist for many years and should be excluded from the count. As snails often move from adjacent roadside verges, it is important to also monitor these areas to assess the risk of snail movement in adjacent paddocks.
Fig. 2.2. Common white snail aestivating on a fence post. (SARDI)
Action level: In cereals, 20 per square metre or more at the time of crop-sowing. Chemical control: A number of baits are registered for snail control. Controls should be applied prior to or immediately after seeding of cereals. The objective is to control adult snails prior to major egg-laying for the season and prevent major increases in numbers that pose a risk at harvest time later in the season. Use the label rate of bait for moderate snail numbers, but if snail densities exceed 80 per square metre this rate should be increased. Cultural control: Stubble management (slashing, rolling or cabling) in January and February is an important control tactic for snails. Snails dislodged from stubble and crop residues onto the soil surface on hot summer days (maximum temperature greater than 35°C) may desiccate and die; snail numbers can be reduced by 50 to 70% using stubble management techniques. Burning stubble residues provides excellent snail control but should only be practiced where the risk of soil erosion is low and when burning is allowed. Burning may reduce snail numbers by up to 99%. Both of these cultural control tactics should be followed with baiting if snail numbers still exceed the established threshold levels.
8
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CEREALS
can cause significant damage to emerging crops and pastures. Like the common white snail, this species also contaminates grain crops at harvest and clogs and damages harvest machinery. Life cycle, risk period, damage, monitoring, action level and control: The same as for the common white snail. Fig. 2.3. The shell of the white Italian snail (shell diameter 17 mm). Note the hole (umbilicus) in the shell on the right is half closed off. (SARDI: G.J. Baker)
Host-plant resistance: No host-plant resistance is recorded for cereals or other host plants. Natural enemies: Apart from some predation by birds and lizards, there are no known natural enemies of the common white snail across southern Australia.
White Italian snail Theba pisana (Müller) Eupulmonata: Helicidae Distribution: An introduced species of European–Mediterranean Basin origin. Recorded from South Africa and Australia (Baker 1986), where it occurs in coastal areas of NSW, SA, Tas., Vic. and WA. Pest status: Minor, restricted, regular. Identification: The diameter of the shell of a mature snail ranges from 10 to 20 mm. The coiled white shell has a broken brown band around the spiral in some individuals, while others completely lack this banding. The umbilicus is semi-circular or partly closed (Fig. 2.3) (Hopkins and Miles 1998). Under magnification cross-hatched scratches/etchings can be seen on the shell.
Natural enemies: Apart from some predation by birds and lizards, there are no known natural enemies of the white Italian snail across southern Australia.
Pointed snail Cochlicella acuta (Müller) Eupulmonata: Hygromiidae Distribution: An introduced species of European origin. Recorded from Australia (Baker 1986), where it occurs in NSW, SA, Vic. and WA, but is most prevalent on Yorke Peninsula in SA. Pest status: Major, restricted, regular. Identification: Fawn, grey or brown in colour with some white markings. The shell is conical in shape (Fig. 2.4). The length of the shell of a mature snail is up to 18 mm (Hopkins and Miles 1998). The ratio of the shell length to its diameter at the base is always greater than two. May be confused with: Immature snails of this species may be confused with the small pointed snail, Cochlicella barbara. They can be separated using the ratio of the shell length to diameter. Host range: This snail feeds on organic matter but has not been reported to damage crops. It is an important contaminant of grain, particularly barley on southern Yorke Peninsula in SA.
May be confused with: The common white or vineyard snail, Cernuella virgata. It can be separated from this species by the different umbilicus (Fig. 2.1) and the scratchings on the shell, and sometimes by the nature of the banding around the spiral of the shell. Host range: The white Italian snail occurs in a broad range of agricultural field crops and pastures including wheat, barley, oats, field peas, faba beans and canola. This species feeds on green plant material and organic matter and
Fig. 2.4. Shell of the pointed snail (shell length 16 mm). (SARDI: G.J. Baker)
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Life cycle on cereals: This species aestivates over summer by climbing onto crop stubble and residues, various weeds and fence posts to avoid high summer temperatures at the soil surface. Autumn rains trigger activity down onto the ground, mating and egg-laying. Most eggs are laid in late autumn or early winter but some egg-laying continues through to early spring. Eggs hatch in about 2 weeks and immature snails grow steadily throughout the winter and spring before they aestivate through the next summer. It can have an annual or biennial life cycle. Risk period: This species causes major contamination of cereal crops, particularly barley, at harvest but should be controlled during January to April, before the crop is sown. Damage: Unlike the common white and white Italian snails, this species does not damage emerging cereal crops and does not cause major clogging of harvesters. However, it passes through harvesting machines and contaminates harvested grain (Hopkins et al. 2003). If limits for snail contamination are exceeded, the load can be downgraded to a lower classification or totally rejected by the grain marketer. Monitoring: Monitor using a square quadrat as for the common white snail. Action level: No thresholds have been established for the pointed snail. Chemical control: Use baits as for the common white snail. Baits are less effective against this snail as compared to the common white or white Italian snails. Cultural control and host-plant resistance: The same as for common white snail. Natural enemies: A parasitic fly, Sarcophaga penicillata, has been released as a biocontrol agent against this species (Fig. 2.5). Its establishment was confirmed in 2003 but its impact on the target species is yet to be assessed (Leyson et al. 2003).
Small pointed snail Prietocella barbara (Linnaeus) Eupulmonata: Hygromiidae
Fig. 2.5. Sarcophaga penicillata (sarcophagid parasite) on the shell of C. acuta. (SARDI: N. Luke)
Distribution: An introduced species of European–Mediterranean Basin origin. Recorded from Australia (Baker 1986), where it occurs in NSW, SA, Tas., Vic. and WA. Pest status: Minor, restricted, regular. Identification: Shell is fawn, grey or brown in colour (Fig. 2.6). The length of the shell of a mature snail is up to 10 mm. The ratio of the shell length to its diameter at the base is always two or less (Hopkins and Miles 1998). May be confused with: Immature pointed snails. They can be separated using the ratio of the shell length to diameter. Host range: Organic matter and green plant material. It has mostly been recorded as a pasture pest in districts with greater than 500 mm annual rainfall. It may feed on cereals, pulses and canola but its pest status is as a contaminant of cereals and canola in higher rainfall districts in SA. Life cycle on cereals: Annual or biennial life cycle. This species aestivates over summer by seeking refuge in clumps of grass or in soil cracks or climbing onto crop stubble and residues to avoid high summer temperatures at the soil surface. Autumn rains trigger activity, mating and egg-laying. Most eggs are laid in late autumn or early winter but some egg-laying continues through to early spring. Eggs hatch in about 2 weeks and immature snails grow steadily throughout the winter and spring before they aestivate through the next summer.
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the cereal zone, particularly on farms where stubble retention is practised. Host range: Includes most crop and pasture plants. Can be significant pests of emerging pastures, pulses and canola and sometimes damages young wheat, oats and barley, particularly those direct-drilled into existing stubble. Fig. 2.6. Shell of the small pointed snail (shell length 8 mm). (SARDI: G.J. Baker)
Risk period: Germination and harvest. Control during January to April, before the crop is sown, avoids contamination of cereal and canola grain at harvest. Damage: May defoliate cereals shortly after crop emergence. Snails pass through harvesting machines and contaminate harvested grain. If limits for snail contamination in grain samples are exceeded, the load can be downgraded to a lower classification or totally rejected by the grain marketers. Monitoring: Monitor using a square quadrat as for the common white snail. Action level: No thresholds have been established for the small pointed snail.
Risk period: Cereals are most susceptible at or soon after crop emergence. Damage is more pronounced when seedling growth is slow due to cool wet weather. Damage: Defoliation of seedling cereals. Monitoring: The presence or absence of slugs can be monitored by providing a series of refuges for slugs, such as carpet, ceramic ties or damp hessian bags randomly placed across a paddock. Action level: None established. Chemical control: Use molluscicide baits in autumn to kill mature slugs before they commence the breeding cycle for the growing season and to protect seedling crops. Where slug densities are high, baiting may not prevent some damage.
Cultural control: Stubble management and burning as for the common white snail.
Cultural control: No cultural techniques have been developed, although European experience suggests that heavy grazing of previous-crop stubbles during summer can reduce oversummering populations. The presence of stubble provides protected habitat, so windrowing of stubbles or cultivation may assist.
Host-plant resistance: No host-plant resistance has been recorded for cereals.
Host-plant resistance: No resistant cereal varieties are available.
Natural enemies: The parasitic fly released against the pointed snail also attacks this species.
Natural enemies: Predatory ground beetles (Family Carabidae) prey on slugs.
Chemical control: Bait as for the common white snail. Baits are less effective against this snail as compared to the common white or white Italian snails.
Slugs Eupulmonata: Limacidae Black-keeled slug, Milax gagates Draparnaud and reticulated slug, Deroceras reticulatum (Müller) N (Main entry in Chapter 5 Oilseeds.)
Pest status: Minor, restricted, irregular. Mainly pests in the high rainfall (> 500 mm p.a.) areas of
Blue oat mites Acarina: Penthaleidae Penthaleus major (Dugès), P. falcatus (Qin and Halliday) and P. tectus Halliday Distribution: Cosmopolitan distribution, unknown origin. Presently found in Europe, the Americas, Asia, South Africa, New Zealand and 11
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Australia. In Australia, P. major has a wider distribution than the redlegged earth mite, occurring across many areas of southern Australia including southern NSW and Qld, SA, Vic., Tas. and WA. The range of P. falcatus appears to overlap with that of P. major, but P. tectus appears to have a discontinuous and restricted distribution in parts of NSW and Vic. (Robinson and Hoffmann 2001). Pest status on cereals: Major, widespread, except for P. tectus, which has a restricted distribution, irregular. Identification: Adult mites are about 1 mm long with a blue–black body, red legs and a red mark on the back (Fig. 15.35). The three species are difficult to distinguish in the field but may be distinguished under a microscope by the length and position of the hairs on their back. P. major has comparatively long hairs running in four or five longitudinal rows down the mite’s back. P. falcatus has short hairs covering the back of its body. P. tectus has short hairs on the anterior of its body and longer hairs running in longitudinal rows on the posterior of its body. May be confused with: Blue oat mites appear similar in size and colour to the redlegged earth mite, but blue oat mites may be distinguished by a red dot on their backs (Fig. 15.35). Blue oat mites are generally seen feeding singularly or in small groups, whereas redlegged earth mites generally feed in larger groups. Blue oat mites prefer grasses and cereals, whereas redlegged earth mites prefer the legume and capeweed content of pastures (Robinson and Hoffmann 2001; Umina et al. 2004). Hosts: Mainly a pest of cereals and grass pastures, but will also feed on pasture legumes and many weeds, including capeweed. Life cycle on winter rainfall cereals and grassy pastures: Blue oat mites are active in the cool wet months from May to November. The first generation develops from oversummering eggs after the onset of favourable conditions of moisture and temperature, usually around April–May. They hatch into six-legged larvae that develop through two nymphal stages into adults. Both nymphs and adults have eight legs.
During the winter, the mites pass through two generations on average, each lasting about 11–12 weeks. When conditions are favourable their populations can increase rapidly, with peaks in autumn and/or spring. Females of both generations lay winter eggs, usually on leaves, stems or roots of food plants. Unlike the redlegged earth mite, both generations of blue oat mite are able to produce diapause (oversummering) eggs, beginning soon after emergence in autumn. However, most of the diapause eggs are produced in spring. The eggs are not retained inside the body of the mite as with redlegged earth mite but are laid on food plants. In late spring, when temperatures increase and the pasture begins to senesce, the mites die. The oversummering eggs remain on the vegetation throughout the summer months (Narayan 1962). Risk period: Autumn, particularly shortly after crop emergence. Damage: Silvering of leaves is symptomatic of damage. Monitoring: At crop emergence look for the presence of mites, being careful to distinguish the blue oat mite from the redlegged earth mite. Accurate identification is important for the selection of appropriate control strategies (Umina and Hoffmann 2004). Action level: No action thresholds have been established for cereals but any evidence of mite activity indicates potential damage. Control: Foliar applications of insecticides may be cost-effective, particularly if applied within 2–3 weeks of emergence in the autumn. Penthaleus spp., particularly P. falcatus, are often more tolerant of insecticides than redlegged earth mite (Robinson and Hoffmann 2001). In addition, the use of control tactics solely in spring will not prevent the carry-over of all eggs into the following autumn, unlike with the redlegged earth mites.
Redlegged earth mite Halotydeus destructor (Tucker) Acarina: Penthalaeidae N (Main entry in Chapter 15 Pastures—winter rainfall.)
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Pest status: Major, widespread and irregular pests of cereals. Pest status is greater if cereals are grown in rotation with legume pastures. Damage: The damage of this species and blue oat mite species is similar. Separate these two mite genera by using the presence (blue oat mites) or absence (redlegged earth mites) of a small red marking in the middle of the back (Fig. 15.35). Host range: Will damage all cereals but cannot survive or reproduce on them (McDonald et al. 1995). Risk period: Autumn to spring, but especially at crop emergence. Monitoring: Close monitoring after crop emergence is required to achieve well-timed treatments. Estimate the numbers of mites in 10 cm × 10 cm squares around the base of plants. Repeat at five to 10 sites across the paddock. Avoid sampling in sunny conditions; sample on cloudy days, early morning or late afternoon. Mite damage is typically greater around the edge of paddocks. Action level: For cereals, treatment is usually warranted at the seedling stage if there are 50 or more mites per 100 cm2 (10 × 10 cm). Chemical control: Bare earth treatments for this pest in autumn are not warranted for cereals. Foliar sprays are cost-effective. In pasture-crop rotations, control in the spring in the pasture phase may remove the need for control in the following cereal crop. The TIMERITE® model is designed to determine the best time for spring treatments. Cultural control: Pest pressure in autumn may be reduced using crop rotations that include lentils or lupins prior to cereals (McDonald et al. 1995), or alternatively heavy spring grazing, cultivation, clean fallowing and destruction of weeds prior to cereals. Natural enemies: See Chapter 15.
Lucerne flea Sminthurus viridis (Linnaeus) Collembolla: Sminthuridae N (Main entry in Chapter 15 Pastures—winter
rainfall.)
Pest status on cereals: Major, widespread and irregular. Host range: Includes all cereals. Life cycle on cereals: Similar to that on winter rainfall pastures (Chapter 15). Risk period: Autumn to spring, but especially at crop emergence. Pest risk is greatest if cereals are grown in rotation with legume pastures. Bare earth pyrethroid treatments for redlegged earth mite in the preceding canola or pulse crop may increase the pressure of this pest in the current cereal crop. Damage: Lucerne flea damage appears as thin, transparent windows on the leaf and is sometimes confused with that of the redlegged earth mite that appears as a silvering of the leaf surface. See more detail under Chapter 15. Monitoring: Monitor for 3–4 weeks after crop emergence. Action level: No thresholds have been developed for cereals. Chemical control: Foliar insecticides are costeffective. Cultural control: No cultural control is available in cereal crops. Host-plant resistance: No resistant varieties of cereals are available. Natural enemies: See Chapter 15.
Sandgroper Cylindracheta psammophila Orthoptera: Cylindrachetidae Distribution: Native to WA. Pest status: Minor, restricted mainly to the sandy coastal areas north of Perth, irregular (Woods and Michael 1987). Identification: Sandgropers remain under the soil surface and are only seen when soil is worked or dug. The front section of the body is hard and orange–brown, whereas the remainder of the body is soft and creamcoloured (Fig. 2.7). The body is cylindrical in
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Damage: Sandgropers attack the underground portion of the stem, and to a lesser extent the roots, giving them a characteristic shredded appearance. Damaged plants turn yellow, wither and die, giving rise to bare or thinned patches in the crop (Fig. 2.8). Monitoring: Look for yellow or wilting plants. Digging in soil may lead to detection of this pest. Action level: None available. Fig. 2.7. Sandgroper (body length 75 mm), an occasional pest of WA cereals. (DAFWA)
shape and up to 75 mm long. The front legs are flat and strong and adapted for digging. Young sandgropers are similar to the adult except they are smaller and paler. May be confused with: Mole crickets, from which they are distinguished by the orange– brown front part of the body. Host range: Includes wheat, barley and sweet lupins. Life cycle on cereals: Adults probably survive the summer deep in the soil. Eggs are probably laid in autumn as immatures are often seen during winter. Risk period: Autumn and winter.
Chemical control: No insecticidal control measures have been developed. Cultural control: Autumn fallowing for several weeks can reduce numbers but may be insufficient to prevent damage. Host-plant resistance: If monitoring detects a heavy infestation is present in a paddock, it may be advisable to sow oats which are less susceptible to attack. Natural enemies: None recorded.
Corn aphid Rhopalosiphum maidis (Fitch) Hemiptera: Aphididae Distribution: An introduced species, probably Asiatic in origin, found in all states of Australia.
Fig. 2.8. Part of a wheat crop thinned by sandgroper feeding on underground parts of plants. (DAFWA)
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Fig. 2.9. A colony of corn aphids on a maize leaf. Each aphid has two siphunculi protruding from its abdomen. Each siphunculus has a purple area at its base, characteristic of corn aphids. The largest, adult, aphid is 2 mm. White skins are moulted casts. (SARDI: G.J. Baker)
Pest status: Minor, widespread, irregular. Identification: Colour varies from light green to dark olive green with two dark-coloured areas near the base of the cornicles at the tip of the abdomen (Fig. 2.9). Adults are approximately 2 mm long. May be confused with: Oat aphid but can be separated using an antennal feature (Fig. 2.10). The terminal process of the sixth antennal segment of the corn aphid is about twice as long as the basal portion, whereas in the oat aphid it is about four to five times as long (Hopkins 1987). Host range: Grasses and cereals including wheat, oats, barley, millet, corn and sorghum. Common on barley across southern Australia. Life cycle on cereals: A parthenogenetic species that undergoes many generations through the growing season. Both alate (winged) and apterous (non-winged) forms occur. Risk period: Most prevalent on cereals in late winter to early spring. High numbers often occur in years when an early break to the season and mild weather in autumn and early winter provide favourable conditions for colonisation and multiplication. Damage: Aphids feed directly on stems, leaves and heads, and in high densities cause yield losses. However, this type of damage is uncommon throughout the cereal belt.
Fig. 2.10. Adult corn and oat aphids and detail of antennae. The antennal segment VI of the corn and oat aphids have different ratios of the length of the terminal process to the length of the basal portion; about 2:1 for corn aphid and 4 or 5:1 for oat aphid.
Corn aphids are vectors of barley yellow dwarf virus (BYDV), a disease of wheat, oats and barley (Fig. 2.11). Yield losses due to BYDV can be severe, particularly in oats and to a lesser degree in wheat. Losses due to BYDV are most severe in the wetter parts of the cereal belt (rainfall > 500 mm p.a.). Monitoring: Assess the potential for directfeeding damage in late winter. Estimate the number of aphids per tiller. In areas of high risk for BYDV, prediction models may assist in deciding whether to use insecticidal seed dressings or early season insecticide applications to reduce or prevent yield losses by BYDV.
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a
b
Fig. 2.11. (a) Depressed patch in an oat crop with reddened leaves, typical of the secondary spread of BYDV (left) (UnivAd: J. Randles) , and (b) the wheat plant on the right is stunted by BYDV compared with a healthy plant on the left (CSIRO: N. Grylls) .
Action level: Aphids are unlikely to cause economic damage to cereal crops expected to yield less than three tonne per ha. To avoid direct-feeding damage, consider treatment if there are 10 to 20 or more aphids on 50% of the tillers. There is no threshold for aphid transfer of BYDV.
parasitism by wasps (Fig. 2.12b) can suppress aphid populations, but this does not happen in every season. Heavy rain may cause significant mortality of aphid populations.
Chemical control: Apply a foliar insecticide in late winter or spring to avoid direct damage to tillers and heads. To prevent major losses from BYDV in virus-prone areas, control aphids early in the cropping year. Use a seed dressing and apply foliar treatments at 4 and 8 weeks after sowing. Foliar treatments with registered pyrethroids provide extended protection through an anti-feedant effect.
Rhopalosiphum padi (Linnaeus) Hemiptera: Aphididae
Cultural control: There are no known effective cultural control methods for this aphid. Host-plant resistance: In virus-prone areas, use resistant plant varieties to minimise losses due to BYDV. Natural enemies: Predation by hoverflies (Fig. 2.12c), lacewings and ladybirds and
Oat or wheat aphid
Distribution: Probably Palaearctic in origin and now cosmopolitan. An introduced species found in all states of Australia. Pest status: Minor, widespread, irregular. Identification: The oat aphid varies in colour from olive-green to black and has a rusty red area around the tip of the abdomen (Fig. 2.12a). Adults are approximately 2 mm long. May be confused with: The corn aphid, from which it can be readily separated using an antennal feature (Fig. 2.10). The terminal process of the sixth antennal segment of the oat aphid is about four to five times as long as the basal
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a
b
Fig. 2.13. Rose-grain aphid (body length 3 mm). (SARDI: G.J. Baker)
c
Distribution: Recorded from NSW, Qld, SA, Tas. and Vic. It is an introduced species that was first recorded in the country in 1984 (Carver 1984). Elsewhere in the world it is found in Europe, Central and West Asia, North and South America and New Zealand. Pest status: Minor, restricted, irregular. Identification: A green or yellowish-green aphid. Adults are about 1.6–3.3 mm long (Fig. 2.13). May be confused with: Unlikely to be confused with other aphid species on cereals because of its distinctive colour.
Fig. 2.12. (a) A wingless adult oat aphid (body length 2 mm) (SARDI: G.J. Baker) , (b) a parasitised aphid (mummy) to the right of which is an unparasitised aphid (VDPI) , and (c) larva of a hoverfly in a colony of cereal aphids (SARDI: G.J. Baker) .
portion, whereas in the corn aphid it is about twice as long.
Host range: Many species of grasses and cereals including wheat, oats and barley. Damage: The rose-grain aphid rarely damages cereals but can transmit BYDV. Life cycle, risk period, monitoring, action level and control: Similar to the corn aphid.
Host range: Includes all major cereal and pasture grasses including wheat, oats, barley, sweetcorn and phalaris. Occurs mainly on oats and wheat in Australia.
Blackheaded pasture cockchafer
Life cycle, risk period, damage, monitoring, action level and control: Similar to the corn aphid.
Distribution: A native species found in NSW, SA, Tas. and Vic.
Rose-grain aphid Metopolophium dirhodum (Walker) Hemiptera: Aphididae
Acrossidius tasmaniae Hope (=Aphodius tasmaniae) Coleoptera: Scarabaeidae
Pest status: Minor, restricted, irregular. Identification: The larvae are white C-shaped grubs with a black or dark brown head capsule. Mature larvae are about 20 mm long (Fig. 15.8). The beetles are black and about 10 mm long (Fig. 15.9).
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May be confused with: This is the only cockchafer species that is likely to damage cereals and whose larva has a black head capsule. It is also the only surface-feeding scarab larva found in cereals. Host range: May damage cereals in higher rainfall cropping districts (approximately 450 mm annual rainfall or greater). More important as a pest of pastures. Life cycle on cereals: Adult beetles are active in late January to March after rain (Allen 1986a). Oviposition occurs during this period and young larvae feed on surface organic matter at night. By the time the cereal crop is sown in autumn, second-instar larvae tunnel in the soil and will surface to defoliate young seedlings. The larvae feed right through the growing season and pupate in December. The next generation of beetles emerges early the following year (January–March). For more details on life cycle, see Chapter 15.
White curl grubs (scarab grubs) Coleoptera: Scarabaeidae Yellowheaded cockchafer, Sericesthis harti (Sharp) Wheat root scarab, S. consanguinea (Blackburn) Black soil scarab, Othnonius batesii Olliff Cockchafer, Heteronyx obesus Burmeister Distribution: S. harti is the main species affecting SA cereal crops (Allen 1986c), S. consanguinea and O. batesii in NSW (Goodyer 1993) and H. obesus in WA (Emery and Szito 1999). Pest status: Minor, restricted, irregular. Identification: The larvae have yellow head capsules and grow up to 40 mm long (Fig. 2.14). The body of the larva is whitish grey when it is feeding and changes to white before pupation. a
Risk period: Seedling stage. Damage: Larval movement to the soil surface at night is triggered by rainfall. They defoliate the young cereal seedlings and take the severed foliage into their soil tunnels before eating. Monitoring: Monitor for the larval stage by taking soil cores to a depth of about 10 cm. Take five to 10 soil cores to estimate the grub density. Action level: Apply chemical control after crop emergence if grub densities exceed 30 per square metre.
b
Chemical control: Some pyrethroid insecticides are registered for control of blackheaded pasture cockchafer larvae. The insecticide must be applied to the foliage of cereal seedlings to be effective. Treatments prior to crop emergence are not likely to be effective. Cultural control: Good pasture cover during summer may deter blackheaded pasture cockchafer beetles from laying their eggs. Host-plant resistance: No resistance cultivars are known. Natural enemies: See Chapter 15 Pastures— winter rainfall.
Fig. 2.14. (a) Larva of yellowheaded cockchafer, Sericesthis harti (body length 25 mm) (SARDI: G.J. Baker) , and (b) larvae of Heteronyx obesus (body length 20 mm) (DAFWA) .
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a
b
cycles. Adults of H. obesus begin to emerge in November and swarm around, and feed on, eucalypt trees for about 2 hours after sunset during warm evenings between December and March. Beetles mate and lay eggs in adjoining paddocks. Eggs hatch in May and survive as small grubs, causing little damage, until the following year when they move to the soil surface to feed on roots of germinating cereals (Emery and Szito 1999). S. harti probably has a 2-year life cycle similar to that of H. obesus. Risk conditions: Crops following several years of pasture are most susceptible. Third-instar larvae feeding during winter and early spring cause the most damage. Damage: Larvae prune the roots of young cereal plants by actively feeding on the roots or damaging them while foraging for soil organic matter. Damaged plants initially grow normally but wither and die at tillering resulting in bare patches in the crop. Monitoring: Monitor for larvae in the soil prior to sowing. White curl grubs are usually in the top 50 mm of moist soil, and birds following the tractor in pre-seeding cultivations are often a sign of the presence of cockchafers. Take soil samples using a soil auger or a spade to a depth of 10 cm and determine the number of larvae per m2.
Fig. 2.15. (a) Beetle of yellowheaded cockchafer (body length 11 mm) (SARDI: G.S. Dearman) , and (b) beetles of Heteronyx obesus (DAFWA) .
Larvae curl into a C-shape when disturbed. Adults are light to dark brown, chunky beetles up to 11 mm long (Fig. 2.15). May be confused with: Separation of pest species (both adults and larvae) is difficult. In addition, large numbers of non-pest species of curl grubs (e.g. H. elongatus may occur with H. obesus) may be found in cereals. Identifications should be confirmed by a specialist. Host range: Larvae feed on the roots of a broad range of crop and pasture plants that include all cereals. Life cycle on cereals: These species may have 1- or 2-year cycles. S. consanguinea has a 1-year life cycle. O. batesii and H. obesus have 2-year life
Action level: Treatment is warranted when there are five or more Sericesthis larvae per square metre or two or more Othnonius per square metre. Twenty H. obesus grubs per square metre can cause visible damage, while more than 50 grubs per square metre will destroy crops (Emery and Szito 1999). Chemical control: Can only be achieved by incorporating insecticides at sowing. If the pest is not detected before seeding and treated, it may be necessary to re-sow damaged areas using an insecticide. Cultural control: Rotations that include intensive cropping or short pasture/crop phases can be used to avoid damage. Current trends to continuous cropping have reduced the pest status of this pest group. Host-plant resistance: No resistant cultivars are available.
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Natural enemies: There are no known natural enemies that are effective against white curl grubs.
Spinetailed weevil or cereal curculio Desiantha caudata Pascoe Coleoptera: Curculionidae Distribution: A native weevil recorded from NSW, SA and Vic. Pest status: Minor, restricted, irregular. Became a pest when tillage practices changed during the 1950s from long-term spring-prepared fallows to short-term autumn-prepared fallows allowing adults to survive the shorter fallow period (Hopkins 1996). Further changes to rotations in the 1990s to continuous cropping have seen this species decline in pest status. Identification: The soil-dwelling larvae are white legless grubs up to 8 mm long with golden head capsule (Fig. 2.16). Adults are greyish-black beetles up to 7 mm long with a definite weevil snout. The female has two spines on the tail end of the body (Fig. 2.17). May be confused with: The larvae of the spinetailed and spotted vegetable weevils can only be distinguished by microscopic examination of the spiracles (May 1977). Host range: Pasture grasses and all cereals and some weeds. Life cycle on cereals: One generation per year. Sexually immature adults emerge from the soil
Fig. 2.17. Female (left) and male (right) beetles of the spinetailed weevil (body length 7 mm). (SARDI: G.S. Dearman)
in November, feed on early summer annual weeds and then shelter under stones or clods of soil during summer. Autumn rain stimulates the adults to resume feeding, to mate and to begin laying eggs. The adults are flightless and lay eggs close to where they emerge. The larvae hatch in 2–3 weeks and feed on germinating grass (or cereal) seed or seedlings. Larvae can also feed on soil organic matter. The larvae feed during winter and spring. Larvae pass through five instars and pupate in the soil during October or early November. Generally, this species needs 3 or more years of consecutive grass dominant pastures to build up to damaging numbers prior to a cereal crop (Allen 1973). Risk period: Following sowing. Damage: The larvae can attack cereal plants at three different stages: they may eat out swelling seeds soon after sowing; they may bore onto the underground part of the stem of seedlings causing them to wither and die; or they may bore into tillers causing them to wither and die. Seed or plant deaths result in thinned or bare patches in the crop. Monitoring: Monitor for larvae using soil sampling prior to seeding. Count the number of larvae by examining soil in a 30 × 30 cm quadrat to a depth of 10 cm. Repeat at five to 10 random sites throughout the paddock. Action level: Treatment is warranted if there are 10 or more larvae per square metre.
Fig. 2.16. Larva of spinetailed weevil (body length 8 mm). (SARDI: G.S. Dearman)
Chemical control: Control of larvae can only be achieved with a seed dressing. As significant
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larval feeding can occur before the seed dressing causes larval mortality, higher than normal seeding rates may be required for highdensity infestations. If there are 100 or more larvae per square metre, use a seeding rate of about 95 kg/ha to compensate for the damage caused before the larvae are killed. Cultural control: Short pasture phases of 1–2 years in rotations with cereals or continuous cropping of cereals, pulses and oilseeds helps avoid damage. Host-plant resistance: No resistant varieties are available. Natural enemies: Soil dwelling pathogens such as the fungus Beauvaria may kill some larvae.
Fig. 2.18. Beetle of spotted vegetable weevil (body length 7 mm). (VDPI)
Arachnodima bribbarensis (Calder)
Spotted vegetable weevil or maculate weevil Desiantha diversipes (Pascoe) Coleoptera: Curculionidae Distribution: A native weevil recorded from NSW, SA, Tas., Vic. and WA. Also recorded in New Zealand. Pest status: Minor, restricted, irregular. Unlike the spinetailed weevil, this species can cause damage to successive crops (Hopkins 1996). Identification: Adults are distinctly mottled black–grey beetles up to 7 mm with a definite weevil snout (Emery et al. 2005). The female has no spines on the tail end of the body as for the spinetailed weevil (Fig. 2.18). The larvae are very similar to those of the spinetailed weevil.
A. xenikon (Calder) A. opaca Candèze A. ourapilla (Calder) Distribution: Wireworm pests of Australian cereals are native species recorded damaging wheat and barley in NSW and SA (Calder 1996). Pest status: Minor, restricted, irregular. Identification: Wireworm larvae have soft semiflattened, smooth creamy white or pale bodies with darker wedge-shaped heads and forked smooth or tooth-edged tails (Fig. 2.19). Arachnodima has paired tail processes, each with two spurs (Fig. 2.19). Agrypnus has a flattened tail plate with spurs around the perimeter (Fig. 2.20). When fully grown, larvae are 25 mm (Agrypnus) and 15 mm (Arachnodima) long.
May be confused with: The larvae of this species and the spinetailed weevil require microscopic examination of spiracles to separate the species. Host range, life cycle, risk period, damage, monitoring, action level and controls: Similar to the spinetailed weevil.
Wireworms Coleoptera: Elateridae. Agrypnus sp.
Fig. 2.19. Larvae of the wireworm Arachnodima sp. (preserved and not true colour) (body length 12 mm). The body is soft and semi-flattened with a wedgeshaped head and six legs. Arrows point to paired tail processes. (SARDI: G.J. Baker)
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Wireworms: true and false Wireworms are named for the supposed wirelike appearance of their larvae. False wireworms belong to the Family Tenebrionidae, the adults of which are dark beetles and may be elongate to oval (‘pie dish’) in shape. True wireworms belong to the insect Family Elateridae. The adults are elongate beetles that jump and click when disturbed. There are numerous species (named and unnamed) of both true and false wireworms, but only a few are recorded as crop pests.
Beetles are brown or black, torpedo-shaped and 15 mm (Agrypnus) and 10 mm (Arachnodima) long. They also have enlarged pronotums with sharp posterior corners. When laid on their backs they can flex their bodies and flip onto their feet. This action is often accompanied by a clicking sound, hence the common name ‘click beetles’. May be confused with: Wireworm larvae can be confused with false wireworm larvae. a
Wireworm larvae have flattened heads while false wireworms have round heads in crosssection. Wireworms may also be confused with the larvae of predatory carabid beetles. However, carabid larvae have enlarged mandibles and a prominent fleshy process on the tail, whereas wireworm larvae have smaller mandibles and a flattened process with short spines, or just forked processes (e.g. Fig. 2.20). Host range: Includes wheat, barley, triticale and oats. Life cycle on cereals: Probably one generation per year; life cycles are not well understood. Larval stages are usually present through autumn and winter. Risk period: At seeding or shortly after crop emergence. Damage: Wireworm larvae feed on the seed and bore into the underground stem of the cereal plants causing them to wither and die (Goodyer 1993). Damage causes thinned crops or bare patches in the crop. Damage usually occurs following long pasture phases of 4–5 years. Monitoring: Monitor for larvae during preseeding cultivations or use a spade to take soil samples to a depth of 10 cm to inspect for larvae. Action level: 40 or more larvae per square metre warrant treatment. Chemical control: Wireworms can only be controlled if they are detected before sowing. Crops can be protected by sowing treated seed or mixing insecticide with fertiliser at seeding.
b
Cultural control: Wireworm numbers can be substantially reduced by clean cultivating in summer and autumn. This discourages egglaying by the adults and kills the young larvae by desiccation and starvation. Rotations including continuous cropping or short pasture phases often reduce wireworm damage. Host-plant resistance: No resistant cereal cultivars.
Fig. 2.20. Tail end of Agrypnus sp. (a) viewed from above and (b) from the side, showing flattened tail plate and spurs. (SARDI: G. Caon)
Natural enemies: Birds feeding on wireworm larvae after cultivation which brings them to the surface. The presence of birds on newly worked ground can often indicate approaching problems
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with false wireworms or wireworms. Soildwelling larval stages of predatory carabid beetles prey on false wireworms and wireworms.
False wireworms Coleoptera: Tenebrionidae N (Main entry in Chapter 5 Oilseeds.)
The main species of false wireworms recorded as pests in the cereal belt include: eastern false wireworm, Pterohelaeus darlingensis; pie-dish beetle, Helea tuberculata; southern false wireworm, Gonocephalum macleayi and Gonocephalum spp.; striate false wireworm, Pterohelaeus alternatus; grey false wireworm, Isopteron punctatissimus; and bronzed field beetle, Adelium brevicorne and Saragus sp. Pest status in cereals: Minor, restricted, irregular. Identification: See Chapter 5. Life cycle on cereals: Larval stages present from autumn to spring and adults aestivate over summer. Risk period: At seeding or shortly after crop emergence. Damage: False wireworm larvae feed on the seed and bore into the underground stem of the cereal plants causing them to wither and die. They may also damage roots, causing thinned crops or bare patches in the crop. This usually occurs following long pasture phases of 4–5 years. Some false wireworms, Isopteron and Pterohelaeus, survive through continuous cropping by feeding on stubble mulch. Monitoring: Monitor for larvae during preseeding cultivations or use a spade to take soil samples to a depth of 10 cm to inspect for larvae. In Qld and Vic., the use of germinating seed baits is recommended (see Glossary). Action level: Treatment is only warranted if larvae are present in large numbers. Treat if there are 10 or more Pterohelaeus, Helea or Saragus or 50 or more Isopteron larvae per square metre. These thresholds do not apply to Gonocephalum sp. where threshold numbers are probably higher. If using germinating seed baits,
treatment is recommended if there are more than 25 southern false wireworm larvae per 20 baits (Elder et al. 1992). Chemical control: False wireworms can only be controlled if they are detected before sowing. Use insecticides mixed with fertiliser at seeding or insecticides sprayed on the soil surface and worked into the soil prior to seeding. Cultural control: Rotations including continuous cropping or short pasture phases may reduce and often eliminate false wireworm damage of some species. Eliminating food and shelter for beetles over summer (various stubble management techniques) can reduce false wireworm problems. Use of press wheels or rolling at planting is recommended in Qld and Vic. High seeding rates can also offset the effects of moderate infestations. Host-plant resistance: No resistant cereal cultivars. Natural enemies: Birds feed on false wireworm larvae after cultivation brings them to the surface. The presence of birds on newly worked ground can often indicate approaching problems with soil-dwelling beetles such as false wireworms or wireworms. Soil-dwelling larval stages of predatory carabid beetles are recorded to prey on false wireworms and wireworms.
Black cutworm Agrotis ipsilon (Hufnagel) Lepidoptera: Noctuidae Distribution: A. ipsilon is a cosmopolitan species. The subspecies A. ipsilon aneituma (Walker) occurs in Papua New Guinea, Australia, New Zealand and some Pacific islands (Common 1993). In Australia, the subspecies is recorded from NSW, Tas., Vic. and WA. Pest status: Minor, restricted, irregular. Identification: Larvae are black, green–brown or grey without distinct hairs or markings and have a greasy appearance. Body is faintly striped or without stripes (Fig. 2.21). Larvae grow up to 40 mm long. The forewing of the moth is pale purplish-brown, brown or grey–black, always
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Damage: Recorded as damaging wheat, oats and barley. The damage symptoms are similar to those of common cutworm. Also damages turf. Monitoring: Larvae hide under clods of soil or bury themselves in soil and may be difficult to find. Examine plants, soil litter and soil surface in a 0.5 m row in at least 10 sites spread evenly through the crop.
Fig. 2.21. Black cutworm larvae (20–30 mm extended). (DPI&F Qld: J. Wessels)
with a pale brown patch towards the tip and a blackish, wedge-shaped mark extending from the outer edge of the large, dark kidney-shaped spot near the centre (Goodyer 1985) (Fig. 2.22). The wingspan ranges from 30 to 50 mm. May be confused with: The larvae may be confused with those of the common cutworm. The granules on the skin of black cutworm are slightly raised whereas those on common cutworm are flattened.
Action level: Treatment of cereals is warranted if there are two or more larvae per 0.5 metre of row. Chemical control: Sprays of pyrethroid insecticides are effective against this cutworm, particularly during early- to mid-instars. Host-plant resistance: No cereal varieties are known to be resistant to cutworm damage. Natural enemies: Parasitic braconid and ichneumonid wasps have been reared from the larval stages of this species.
Brown or pink cutworm
Host range: Cereal, pulse and pasture grasses and a number of weed species.
Agrotis munda Walker Lepidoptera: Noctuidae
Life cycle on cereals: Several overlapping generations per year. Eggs are laid in cracks in the ground and also on vegetation (Williamson and Potter 1997). Eggs laid on the ground may survive tillage, and survival is increased in low or minimum-tillage culture. There is evidence of long-range migration of adults.
Distribution: A native species, recorded from NSW, SA, Tas., Vic. and WA.
Risk period: Black cutworm is most damaging during spring and autumn.
Pest status: Minor, restricted, irregular. Outbreaks are usually patchy. In SA, major outbreaks in cereals occur about 1 in 10 years, affecting tens of thousands of hectares (Hopkins 1982). Identification: Mature larvae grow up to 40 mm long, are greyish-green to brown and often with a reddish tinge, and generally have a greasy appearance (Fig. 2.23a). Moths have light to dark brown forewing with a light grey hind wing (Fig. 2.23b). Wing veins on the hind wing often appear as dark lines. Wingspan ranges from 30–40 mm. May be confused with: Common and black cutworm, which generally have much darker larval stages (Goodyer 1985).
Fig. 2.22. Adult black cutworm (wingspan 50 mm). (DPI&F Qld: J. Wessels)
Host range: An important pest of lucerne, pulses and summer fodder crops. Also damages late-sown clover and medic seed crops and late-sown cereals.
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a
b
Host-plant resistance: No cereal varieties are known to be resistant to cutworm damage. Natural enemies: In outbreaks of this cutworm, predatory carabid larvae and sphecid wasps have been observed preying on cutworm larvae. Insect pathogens (diseases) have also been recorded affecting larvae during large brown cutworm outbreaks, particularly in late autumn or early winter.
Common cutworm or Bogong moth Agrotis infusa (Boisduval) Lepidoptera: Noctuidae Distribution: A native species, recorded from NSW, SA, Tas., Vic. and WA. Fig. 2.23. (a) Brown cutworm larva (length 40 mm) (SARDI: G.J. Baker) , and (b) moth (wingspan 40 mm) (VDPI) .
Life cycle on cereals: Moths prefer to lay their eggs in soil in lightly vegetated or bare areas. Early autumn egg-laying results in most damage to young cereals. Larvae hatch and feed on host plants right through to maturity. Mature larvae pupate in the soil. Risk period: Damage may occur between late autumn and early winter but is more common in late-sown cereal crops in spring. Damage: Larvae 20–40 mm long cause the most damage when they ringbark or cut off seedlings at ground level. Monitoring: Larvae hide under clods of soil or bury themselves in soil and may be difficult to find. Examine plants, soil litter and soil surface in a 0.5 m row in at least 10 sites spread evenly through the crop. Action level: Treatment of cereals is warranted if there are two or more larvae per 0.5 m of row. Chemical control: Pyrethroid insecticides are effective against this cutworm, particularly in the early to mid-larval stages. Cultural control: Avoid late sowing (i.e. in late winter) of cereals.
Pest status: Minor, restricted, irregular. Identification: Eggs are about 0.8 mm in diameter, flattened with radiating ridges (Fig. 2.24). Larvae are black, green–brown or grey without distinct hairs or markings and have a greasy appearance (Fig. 2.24). Body is faintly striped or without stripes. Larvae grow up to 40 mm long. Adults are often observed in houses and buildings in late spring and early summer seeking shelter. Moths have a dark brown or grey–black forewing with black or grey markings near the centre (Fig. 2.25). The wingspan ranges from 30–50 mm. May be confused with: Common cutworm larvae of this species may be confused with those of the black cutworm. For differences, see black cutworm. Host range: Cereal, pulse and pasture plants. Recorded as damaging wheat, oats and barley. Life cycle on cereals: Moths aestivate during summer. They are carried large distances by wind in September–November to cluster in caves and crevices in mountainous areas of south-eastern Australia (Common 1954). The moths become active in autumn and these return migrations are continuous throughout the autumn and early winter. Egg-laying is not synchronised so larvae hatch throughout this period.
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a
b
Fig. 2.26. A cereal plant severed just above ground level by night-feeding cutworm larvae. The name ‘cutworm’ is derived from these damage symptoms. (VDPI)
Natural enemies: A range of braconid and ichneumonid wasps have been reared from the larval stages of this species.
Pasture webworm Hednota spp. Lepidoptera: Pyralidae. There are probably up to five different species in the complex (Woods and Michael 1987). Fig. 2.24. (a) Common cutworm eggs, each 0.8 mm diameter, and (b) pupa and fully-grown larva (length 40 mm) of the common cutworm, Agrotis infusa. (Graphic Science: © Dennis Crawford)
Risk period: Most damaging in late autumn and winter as larvae approach maturity. Damage: Common cutworm damages wheat, oats and barley by ringbarking or completely defoliating the plants (Fig. 2.26). Monitoring, action level and control: Similar to brown cutworm.
Distribution: A native group of species recorded in NSW, SA, Tas., Vic. and WA. In NSW, the sod webworm Ptochostola microphaella, has similar behaviour to Hednota spp. (Goodyer 1993). Pest status: Minor, restricted, irregular. Pest status has declined in recent years as pastures have been phased out of cereal rotations. Important in marginal cereal growing areas of SA, Vic. and WA. Identification: Larvae are smooth-bodied with dark shiny heads and grow up to 18 mm long. Older larvae have darker raised patches on each segment (Fig. 2.27). Larvae live in silken-lined tunnels in the ground. Adult moths are about 12 mm long when wings are folded over the body. They are grey–brown with protruding beak-like mouthparts. May be confused with: Unlikely to be confused with other species. Host range: A range of grasses. Damage is mainly to wheat and barley (Allen 1986b).
Fig. 2.25. Moth of the common cutworm (wingspan 40 mm), showing grey markings on the forewing. (VDPI)
Life cycle on cereals: One generation per year. Moths fly during March, April and May and lay their eggs on the ground. During the day,
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Sometimes severed leaves can be seen protruding from these tunnels. In heavily infested areas bare patches appear in the crop (Fig. 2.28), and with lower numbers, extensive thinning of the crop occurs. Monitoring: Inspect crops for damage every few days after crop emergence. Continue monitoring for 2–3 weeks as most damage is done during this period. When monitoring, look for severed leaves protruding from silken-lined tunnels in the soil.
Fig. 2.27. Full-sized larvae of pasture webworm, Hednota sp. (body length 18 mm). (VDPI)
the moths shelter in standing dry grass. The eggs hatch in 1–3 weeks and the young larvae construct silken tunnels in the soil. They feed on the foliage of grasses and cereals by cutting off leaves at ground level, usually during night, and carrying them into their tunnels. The early-hatched larvae finish feeding in June, but later-hatched larvae may feed for another 3 months. Mature larvae seal their tunnels and remain dormant during summer. They pupate in autumn and emerge as moths shortly after.
Action level: Control if 10 or more plants per square metre are damaged. Even heavily infested crops will recover if treatments are timed correctly. Chemical control: Most pyrethroid insecticides are effective as foliar sprays.
Risk period: At crop emergence and the 2–3 weeks following. Expect damage in cereals that are grown after 3–5 years of pasture.
Cultural control: Pastures that are heavily grazed before March are not attractive to moths and are not favourable for the survival of larvae. This grazing management can be used to reduce the incidence of this pest in paddocks that are to be sown to cereals following minimum cultivation. After an initial full cultivation or use of a knockdown herbicide, the risk is slight if the time of sowing the crop is delayed for at least 4 weeks. The young larvae cannot survive starvation for more than 3 weeks. Delayed sowing may not be advisable in marginal areas where early sowing is essential to maximise the benefits of early moisture. In these areas, crops should be sown as early as possible and treated for webworm if they occur.
Damage: Larvae sever the leaves of young cereal plants and drag them into their tunnels.
Host-plant resistance: No resistant varieties are available.
Fig. 2.28. Bare patches in a young cereal crop caused by pasture webworm. (SARDI)
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Natural enemies: Little information is available on natural enemies.
P E S T S O F T H E V E G E TAT I V E G R OW T H S TAG E O F C E R E A L S Pests that are risks during seedling growth may continue to be risks to the vegetative stage. These include blue oat mite, redlegged earth mite, brown wheat mite, lucerne flea, corn aphid, oat aphid, rose-grain aphid, black cutworm, brown cutworm, common cutworms and pasture webworm (see entries above).
Wheat curl mite and cereal rust mite Acarina: Eriophyidae
often appear as aggregations of white debris on the edges and tips of the leaves and flower awns. Host range: Wheat curl mite can be found on cereals and on a number of introduced grasses including annual and perennial ryegrass, barley grass, brome grasses, wild oats Avena fatua and bearded oats A. barbata. The cereal rust mite is common on perennial and annual ryegrass in southern Australian pastures. Life cycle: As the grasses mature, the mites often enter a migratory phase and move to the apex of the leaves and flower stems, where they are dispersed by the wind. Damage: Eriophyid mites cause no direct physical damage to their host plants, but may cause indirect crop losses through the transmission of a number of viral diseases (potyviruses).
Pest status (both species): Minor, widespread, regular.
Wheat curl mite is very widely distributed on wheat and barley throughout southern Australia and is the vector of wheat streak mosaic virus (WSMV). Little is known of the distribution and economic impact of WSMV in Australia. However, it is likely to have been present in Australia for some years, and has probably fully extended over its potential geographic range. Grain losses due to WSMV in Australia are probably small.
Identification: These eriophyid mites are semimicroscopic, elongate, phytophagous mites which, although extremely common, are seldom noticed (Fig. 2.29). On grass and cereal hosts the mites appear as very small white specks within the grooves of the upper surfaces of the leaves. During the migratory phase, eriophyid mites
The cereal rust mite is common on perennial and annual ryegrass in southern Australian pastures. It is the vector of ryegrass mosaic virus (RMV), a widely distributed disease of grasses. While RMV has been shown to decrease pasture yields under some circumstances, its economic impact in Australia is likely to be insignificant.
Aceria tosichella (Keifer) and Abacarus hystrix (Nelapa) Distribution: Wheat curl mite is widely distributed on wheat and barley throughout southern Australia. The cereal rust mite is common on perennial and annual ryegrass in southern Australian pastures.
Control: Control is not recommended.
Brown wheat mite Petrobia latens (Müller) Acarina: Tetranychidae Distribution: Recorded from NSW and Qld. Pest status: A major pest in southern Qld and NSW, irregular.
Fig. 2.29. Wheat curl mite (0.2 mm long) among leaf hairs on a cereal leaf. Mites in the Family Eriophyidae have only four legs. (W.E. Frost)
Identification: Adult brown wheat mites are oval, about 0.6 mm long and greenish-brown to nearly black. Immature mites are smaller and orange–red. Feeding causes fine mottling on leaves.
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May be confused with: A very small mite that is unlikely to be confused with the redlegged earth mite or blue oat mite. Host range: Probably most cereals and grasses. Life cycle on cereals: Brown wheat mite oversummers as eggs laid in the soil under clods, stones or plant debris. The eggs hatch after heavy rain in autumn or winter, but infestations only develop if the following weather is warm and dry. Risk period: Crops are at risk during warm, dry periods.
Fig. 2.30. Larva of a grass anthelid, Pterolocera sp. (body length 50 mm). (Graphic Science: © Dennis Crawford)
Damage: Damaged leaves look bronzed or yellowish. When viewed from a distance, crops have a bronzed or yellowish appearance (Elder et al. 1992).
Life cycle on cereals: One generation per year. Caterpillars increase in numbers during winter in grassland or pastures and often invade the edges of cereal crops.
Monitoring: Monitor cereals for the presence of mites in autumn or spring, particularly if dry conditions are prevailing.
Risk period: Late winter and early spring.
Action level: Spray if bronze–yellow patches appear throughout the crop and conditions are dry without the prospect of rain (Elder et al. 1992). Chemical control: Foliar treatments may sometimes be cost-effective.
Damage: Defoliation of cereal plants in late winter or early spring. Monitoring: Monitor crop edges adjacent to grasslands or pastures. Action level: No thresholds developed. Chemical control: Pyrethroid insecticides provide effective control.
Cultural control: None available. Host-plant resistance: No resistant cultivars. Natural enemies: No natural enemies recorded.
PE S T S O F M AT U R E C E R E A L S
Australian plague locust Grass anthelid Pterolocera amplicornis Walker Lepidoptera: Anthelidae Distribution: A native grass-feeding anthelid recorded from NSW, SA, Tas., Vic. and WA. Pest status: Minor, restricted, irregular. Identification: A hairy caterpillar that can grow to 50 mm in length (Fig. 2.30). The male moth is fawn with feathered antennae and a wingspan of about 40 mm. May be confused with: Unlikely to be confused with other caterpillars damaging cereals. Host range: A grass-feeding insect that will damage all cereals.
Chortoicetes terminifera (Walker) Orthoptera: Acrididae N (Main entry in Chapter 18 Locusts and grasshoppers of
pastures and rangelands.)
Pest status in cereals: Major in all mainland states, widespread, irregular. Host range: Includes all cereals. Life cycle on cereals: It is usually the spring generation that affects cereal crops. Hopper bands may march and adult swarms may fly into cereal crops late in the growing season as the crop approaches maturity. Adult swarms may also fly into cereal cropping districts in late autumn/early winter and damage newly emerged cereals. 29
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Risk period: There are two main risk periods for cereals: mid to late spring when cereals are approaching maturity, and in autumn when newly sown crops at risk. Damage: Both nymphs and adults can defoliate crops. Damage in maturing crops can be particularly severe if head-lopping occurs. Monitoring: Paddock transects should be used to detect locust activity. Nymphal bands are often very discreet and it is important to map these accurately so that control measures can target the infested areas. Swarms of adults are easily detected as they are flushed from the crop by the movement of a vehicle or person. As swarms (and to a lesser degree bands of nymphs) are very mobile, monitoring should be conducted regularly during times of risk. Action level: In cereals, when nymphs aggregate to form bands, they warrant treatment. A dense nymphal band may have up to 1000 individuals per square metre. Swarms should be treated when there are four or more adults per square metre. Chemical control: A number of insecticides are registered for the control of plague locust. A formulation of Metarhizium (a pathogenic fungus) is also available for use on properties accredited for organic production and sensitive environmental areas. Control of nymphal bands can be achieved with ground or aerial spraying but swarm control is best achieved by aerial spraying. The Australian Plague Locust Commission and various state agencies usually conduct coordinated area-wide control programs in plague years. Cultural control: No cultural control is available for nymph or adult control. If egg beds occur in arable areas, cultivation of these egg beds is known to cause significant mortality. Host-plant resistance: No host-plant resistance in cereals or other hosts. Conservation of natural enemies: Scelionid egg parasites control plague locusts in some years but there are no known ways to enhance their impact.
Migratory locust Locusta migratoria (Linnaeus) Orthoptera: Acrididae
Pest status: Major in Qld, irregular. Can cause damage to wheat, oats, barley and triticale in Qld in outbreak years. For further details see Chapter 18 Rangeland grasshoppers and locusts.
Small plague grasshopper Austroicetes cruciata (Saussure) Orthoptera: Acrididae N (Main entry in Chapter 18 Locusts and grasshoppers of
pastures and rangelands.)
Pest status on cereals: Minor in SA, restricted, irregular. Outbreaks in 1998 and 1999 around Orroroo and Peterborough in SA. Risk period: Spring to harvest. Damage: Consumption of leaves and stems results in many cereal heads falling to the ground. Monitoring: Monitor for bands of nymphs or swarms of adults. Action level: If bands are present, treatment is warranted. For adults, treat if there are four or more adults per square metre. Chemical control: There are governmentmanaged control programs in SA. For further information see Chapter 18.
Common armyworm Leucania convecta (Walker) Lepidoptera: Noctuidae Distribution: Native Australian species, recorded in NSW, Qld, SA, Tas., Vic. and WA. Pest status: Major, widespread, irregular. Identification: First-instar larvae are about 1 mm long. From the second instar, stripes develop along the top and sides of the larva and become more distinct as the larva grows. Crowded larvae are usually darker than those uncrowded. The mature larva grows up to 40 mm in length and has three characteristic pale stripes on the head, collar (segment behind the head) and tail segment. They are smoothbodied with no distinct hairs. The body of the larva also has lateral stripes (Fig. 2.31).
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Armyworms (Lepidoptera: Noctuidae) Armyworm larvae may be distinguished from budworms and cutworms by three prominent stripes on the first segment (or ‘collar’) behind the head. Armyworms owe their name to the observation that larvae sometimes ‘march’ in large numbers away from crowded sites and areas depleted of food. Armyworms of importance in
Australian agriculture belong to the genera Leucania (previously Mythimna), Persectania and Spodoptera (see also Chapter 7 Pulses—summer, and Chapter 13 Pastures—summer rainfall). Larger larvae have stripes along their bodies. Larvae of Leucania and Persectania are generally night-feeders, except during plagues, while larvae of
Spodoptera are mainly dayfeeders. In spite of their name, armyworm larvae are usually solitary feeders in cereal crops. Armyworms feed on grasses, including graminaceous crops, and sedges. Adults of a number of armyworm species are known to undertake long-distance migrations at night.
To separate common and northern armyworm larvae, use the following information. In mature common armyworm larvae, the horizontal width of the centre of the spiracle on abdominal segment 8 is about seven times the width of its rim, whereas in northern armyworm this ratio is only about three and half times (Goodyer undated). Host range: Damages barley, oats, wheat, native pasture grasses and perennial grass seed crops.
Fig. 2.31. Armyworm larva curled up on soil (body length 30 mm). Droppings (to left of larva) are an indication of armyworm activity. (SARDI: G.J. Baker)
Life cycle on cereals: Common armyworms have three generations per year. The winter and spring generations damage cereals. Moths fly into cereal crops and lay their eggs in the folds of dried or drying leaves on grasses or cereals. Females lay up to 1000 eggs in irregularly-shaped
The forewings of the moth have a wingspan of about 40 mm and are fawn or buff coloured (Fig. 2.32). May be confused with: The adults may be confused with those of the northern armyworm. Genitalia dissections by a specialist are required to separate these two species. The larval stages of the four armyworm species likely to be encountered in cereals (common, inland, northern and southern) are all similar in appearance. On mature larvae of common and northern armyworms, the hair above the spiracles on abdominal segments 3, 4, 5 and 6 (Fig. 2.33) is on a small dark plate near the middle of the dark stripe (Fig. 2.34a). On the southern and inland armyworm, the hair above the spiracles on abdominal segments 3, 4, 5 and 6 is on a large dark plate near the upper edge of the dark stripe (Fig. 2.34b) (Goodyer undated).
Fig. 2.32. Male (upper) and female (lower) moths of the common armyworm (wingspan 40 mm). (SARDI: G.J. Baker)
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Fig. 2.33. Position of thoracic and abdominal segments of a caterpillar. (NSWDPI: A.Westcott)
masses, cemented in tight folds of foliage. Eggs hatch as little as 3–4 days after laying and young larvae, with the assistance of wind, disperse through the crop on fine silken threads (McDonald 1991a). The larvae feed on leaves and stems. Larvae usually develop through six instars but sometimes seven. Indicative a
b
Fig. 2.34. Distinguishing mature larvae of Leucania spp. from Persectania spp. (a) Abdominal segments 3, 4 and 5 of a Leucania larva, showing the hair above the spiracles on a small plate (arrowed) near the middle of the dark stripe (SARDI: G. Caon) , (b) segments 3, 4 and 5 of a Persectania larva, showing the hair above the spiracles on a large dark plate (arrowed) near the upper edge of the dark stripe (SARDI: G. Caon) .
development times at constant temperatures are: egg-laying to hatch, 7 days at 20°C and 2.5 days at 30°C; larval stages (including pre-pupal stage) 34.2 days at 20°C and 17.2 days at 30°C. Larvae pupate in the soil. The pupal stage lasts 20.1 days at 20°C and 10.1 days at 30°C. Development time from neonate egg to adult emergence is 61 days at 20°C and 41 days at 30°C (Smith 1984). Risk period: The greatest risk to cereals is spring. Moth flights often occur in September and October, and the later-stage larvae damage cereals often in the weeks prior to harvest. The mature larval stages of the winter generation will sometimes march in cereal crops in late winter and cause serious damage to crops, particularly on the edges of paddocks. Crops directly seeded into standing stubbles are susceptible to severe defoliation during the vegetative stage as the winter generation matures. Damage: There are two distinct periods for economic damage. The first, defoliation during early vegetative development, is less common than the second during ripening. Ripening barley is most susceptible to armyworm damage because the last part of the plant to dry off is the green tissue of the stem just below the head. Mature larvae feed on this area and thereby sever the stem causing the head of the cereal to fall to the ground (Fig. 2.35a). One larva can lop many heads very quickly causing large grain losses. Oats are also damaged but the less compact seed head means less damage. In northern Australia, wheat can also be damaged, but in the south the wheat head stays green later and armyworms feed along the heads and damage grain rather than excise the whole head (Fig. 2.35b). Monitoring: Large numbers of armyworm moths are attracted to farm lights on warm nights in September and October. This provides the first warning of potential problems in cereals. Armyworm larvae are difficult to find in cereal crops as they hide at the base of plants or under clods of soil during the day. Search at the base of plants and under clods of soil to estimate the number of larvae per square metre. The presence of green–yellow pellet-shaped droppings of the larvae on the ground are usually a reliable sign of larvae (Fig. 2.31). Monitor for larvae at dusk with a sweep net; sweep netting during the day can be unreliable.
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a
b
Fig. 2.35. (a) Cut barley stems from which heads of grain have been lopped by night-feeding armyworms (VDPI) , and (b) grain damaged earlier by armyworm feeding (Graphic Science: © Denis Crawford) .
Action level: Two larvae per square metre for barley. Other cereals are likely to tolerate slightly higher numbers. Chemical control: A range of insecticides is registered for armyworm control in cereals. Insecticides should target larvae 10 to 20 mm long. Larvae larger than 20mm long can be difficult to kill and may require higher rates of insecticide. If possible, spray late in the day as larvae are active at night. Cultural control: Windrowed or swathed crops dry out rapidly rendering them unattractive to the feeding of armyworm larvae. They are also less susceptible to wind damage (head shattering).
Host-plant resistance: No cereal varieties are known to be resistant to common armyworm damage. Natural enemies: Armyworm larvae are attacked by a number of parasitoids that may be important in reducing the intensity of outbreaks. However, when armyworms are in crops and in numbers likely to cause damage, parasitoids are unlikely to give timely control. Parasitoids bred from armyworms collected in south-eastern Australia by McDonald and Smith (1986) include several wasps: Hymenoptera, Braconidae, (including one sample 32% parasitised by Apanteles sp., Fig. 2.36), Eulophidae, Ichneumonidae (including the most common parasitoid, Campoletis sp. which parasitised up to 59% of armyworms in one sample) and Encyrtidae. In WA cereal areas, Woods et al. (1980) recorded some samples with up to 60% parasitism by Litomastix sp. (Encyrtidae) (Figs 7.152, 7.153). Tachinid (Diptera) parasitoids appear of minor importance in eastern Australia but Woods et al. recorded up to 81% of one WA sample parasitised by Palexorista sp. (Diptera: Tachinidae). In south-east Qld, Broadley (1986) recorded total average parasitism by Hymenoptera and Tachinidae of 18%, with one sample recording 52% parasitism by Cotesia ruficrus (Haliday). Predators include green carab beetles, Calosoma schayeri Erichson (Coleoptera: Carabidae) (Fig. 3.42), populations of which increase dramatically in inland Australia in response to abundant noctuid larvae induced by favourable seasons. Other predators include the predatory shield bugs, Cermatulus nasalis (Westwood) (Fig. 3.38), and Oechalia schellenbergii (GuérinMéneville) (Fig. 3.39) (Hemiptera: Pentatomidae), and perhaps common brown earwigs, Labidura truncata Kirby (Dermaptera: Labiduridae) (Horne and Edward 1995). Fungal diseases, including Nomuraea rileyi, are recorded as causing mortality of armyworm larvae (Broadley 1979).
Northern armyworm Leucania separata (Walker) Lepidoptera: Noctuidae Distribution: Distributed throughout SouthEast Asia, New Zealand and in Australia, where it occurs in all states except Tas. 33
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a
Fig. 2.37. Northern armyworm (35 mm), Ord River, WA. (M. Shepard)
Pest status: Minor pest of cereals in Qld, restricted, irregular. Identification: Larvae (Fig. 2.37) and adults (Fig. 2.38) are very similar to the common armyworm. May be confused with: Common armyworm. Genitalia dissections are required to separate the moths of common and northern armyworm. See common armyworm (page 32) for details on how to separate the larval stages of northern armyworm from other armyworm species. Host range: In Qld, recorded as damaging sorghum, maize, barley, wheat and rice. In New Zealand, the species damages maize and pastures (Chapman 1984). b
Fig. 2.36. (a) Armyworm larva with cocoons of Apanteles sp., and (b) emerged adult. (DPI&F Qld: J. Wessels)
Fig. 2.38. Northern armyworm (38 mm wingspan). (M. Shepard)
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Fig. 2.39. Sugarcane armyworm larva (30 mm long). (DPI&F Qld: J. Wessels)
Life cycle, damage, monitoring, action level and control: Similar to the common armyworm. Risk period: In Qld, early summer. Natural enemies: As for common armyworm. A wasp parasitoid, Cotesia ruficrus, introduced to New Zealand provides some control of this species in maize crops (Burgess 1987; Hill 1988).
Sugarcane armyworm and nightfeeding sugarcane armyworm Lepidoptera: Noctuidae Leucania stenographa Lower and L. loreyi (Duponchel) Distribution: L. stenographa is recorded from Asia and Australia, where it occurs throughout drier parts of the mainland and as an occasional vagrant to Tas. L. loreyi is recorded from Europe, Asia, some Pacific Islands and Australia, where it is recorded from NT and Qld as far south as Brisbane (Edwards 1992).
Fig. 2.40. Adult sugarcane armyworm (35 mm wingspan). (DPI&F Qld: J. Wessels)
Distribution: It is a native species that only occurs in Australia. Found across southern Australia; NSW, SA, Tas., Vic. and WA. Pest status: Major, restricted, irregular. Identification: The mature larva grows up to 40 mm in length and has three characteristic pale stripes on the head, collar (segment behind the head) and tail segment (Fig. 2.41). The body of the larva also has lateral stripes. The moth is grey with a wingspan of up to 45 mm. The forewing has white streaking and a characteristic white fish-shaped marking near the leading edge of the wing (Fig. 2.42). May be confused with: Adults of the inland armyworm. However, the inland armyworm moth lacks the fish-shaped marking on the forewing. To separate the larval stages of
Pest status in cereals: Larvae (Fig. 2.39) of L. stenographa are recorded as a minor pests, occasionally damaging some WA grain crops. Identification: Adult L. stenographa (Fig. 2.40) has a lighter forewing than L. loreyi, which has more prominent lines on its forewing than has L. stenographa. The species can only be separated with certainty by dissection of adult genitalia (Edwards 1992).
Southern armyworm Persectania ewingii (Westwood) Lepidoptera: Noctuidae
Fig. 2.41. An armyworm larva showing the longitudinal stripes characteristic of all species of armyworms. (VDPI)
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a
b
Fig. 2.42. (a) Moth of the southern armyworm (wingspan 40 mm). Note the white fish-shaped marking (arrowed) near the leading edge of the wing (Vic DPI) . (b) Moth at rest (Graphic Science: © Denis Crawford) .
Persectania spp. from Leucania spp., see common armyworm and Fig. 2.36 on page 34. Rearing the larvae of Persectania spp. through to moths is the most reliable way to differentiate the larvae of southern and inland armyworms. Host range: Barley, oats, wheat and perennial grass seed crops may be damaged by southern armyworm (Hopkins 1999). However, vegetative oats and seedling barley suppress larval growth and survival. Many native and introduced grasses are hosts (McDonald 1991b). Life cycle on cereals: Southern armyworms can have three generations per year. It is the winter and spring generations that damage cereals. Moths fly into cereal crops and lay their eggs on dried grasses or cereals. Eggs hatch soon after and the larvae disperse and feed on the cereal foliage, causing more damage as they grow. Risk period: Spring. Moth flights often occur in September and October and large larvae cause most damage just prior to harvest. In addition, the mature larval stages of the winter generation will sometimes march in cereal crops during late winter and seriously damage crops, particularly on the edges of paddocks.
Damage: Barley is the most susceptible cereal to armyworm damage because the last part of the plant to dry off is the green tissue of the stem just below the head. Mature larvae focus their feeding in this area and thereby sever the stem causing the head of the cereal to fall to the ground. One larva can lop many heads very quickly causing large grain losses. Oats are also attacked but the less-compact seed head results in less damage. The wheat head remains green to late in the development of the crop and armyworms tend to feed along the heads and damage grain. Monitoring: Large numbers of armyworm moths swarming around farm lights on warm nights in September and October provide the first warning of potential damage to cereals. Armyworm larvae are difficult to find in cereal crops during the day as they hide at the base of plants or under clods of soil. Search at the base of plants and under clods of soil to estimate the number of larvae per square metre. The presence of green–yellow pellet-shaped droppings of the larvae on the ground are usually a reliable sign that larvae are present (Fig. 2.31). Monitoring for larvae at dusk using a sweep net or plastic bucket through the canopy can provide a rapid assessment; sweep netting during the day is unreliable. Action level: Barley is two larvae per square metre. Other cereals may tolerate slightly higher numbers. Chemical control: There is a broad range of insecticides registered for armyworm control in cereals. Insecticides should target larvae 10– 20 mm long. Larvae greater than 20 mm long can be difficult to kill with most insecticides, and even higher rates sometimes give variable control. If possible, spray late in the day as larvae are active at night. Cultural control: Windrowing barley may be an alternative to spraying. Natural enemies: Armyworm larvae are parasitised by a range of braconid and ichneumonid wasps and tachinid flies but their presence rarely reduces densities below the action threshold.
Inland armyworm Persectania dyscrita Common Lepidoptera: Noctuidae
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directly on the grain in the heads of wheat, barley and triticale.
Cotton bollworm Helicoverpa armigera (Hübner) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Science: © Denis Crawford)
Pest status: Minor pest of cereals in northern NSW and a major pest in Qld (Elder et al. 1992), where it is an irregular pest. The host range includes wheat and barley.
Distribution: A native species that only occurs in Australia. Found across southern Australia, NSW, Qld, SA, Tas., Vic. and WA.
Risk period: On cereals, larvae feed during September to November after an early spring moth flight. The main risk period is when the heads are green.
Fig. 2.43. Moth of the inland armyworm at rest. Note the two distinct white markings on the forewing. (Graphic
Pest status: Minor, restricted, irregular. Incidence of severe damage appears to be greater in SA, WA and some parts of NSW than in Vic. and Tas. Identification: A grey moth with a wingspan of about 45 mm (Fig. 2.43). The larval stages are as described for the southern armyworm. May be confused with: Southern armyworm. The inland armyworm lacks the clear fishshaped marking present on the forewing of the southern armyworm. The hind wing of the inland armyworm is generally lighter in colour than that of the southern armyworm. Rearing the larvae of Persectania spp. to moths is the most reliable way to differentiate the larvae of southern and inland armyworms.
Damage: Damage is caused by feeding of mature larvae on the heads of wheat and barley. Cotton bollworm can be monitored with a sweep net between flowering and harvest or when larvae are observed in the heads of the cereal crop (Fig. 2.44). Sample 2 m lengths of row at six positions widely spaced in a paddock. Action level in Qld is one or more larvae per metre of row (Elder et al. 1992).
Life cycle, risk period, damage, monitoring, action level and control: Similar to the common armyworm.
Native budworm Helicoverpa punctigera (Wallengren) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Pest status: A minor and irregular pest of cereals in all states. In SA, damage in cereals has only been recorded in 1 or 2 years over a 20-year period. Damage: Native budworms only damage cereals in years where there are very high numbers present prior to harvest. Mature larvae feed
Fig. 2.44. Cotton bollworm larva feeding on a cereal plant during daytime. (Graphic Science: © Denis Crawford)
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Migrations of cereal pests in south-eastern Australia The sudden appearance of many pest species in cropping areas of southern Australia suggests strongly that these insects have migrated from elsewhere, generally assisted by favourable weather conditions. Although some insect migration occurs annually, exceptionally large flights often follow droughts, subsequently causing severe damage to crops. A number of endemic insects are well equipped to exploit variable rainfall across the continent by migratory behaviour associated with rain-bearing weather. One well-understood moth migrant is the common armyworm. Adult moths migrate southwards from winter grassy refuges in southern Qld and northern NSW (Fig. 2.45). The moths can migrate many hundreds of kilometres on northerly and north-westerly airstreams that typically precede east-moving weather fronts across the continent in spring (Fig. 2.45). These migrant populations (and often ‘plagues’) can be concentrated as outflows from intense prefrontal storms. These migrants mate, lay eggs and develop on cereal crops and grassy pastures during spring and early summer. As crops and pastures dry during summer, densities of armyworms decline, except in some coastal areas where grass habitats remain green (Fig. 2.45). It is likely that the relatively small populations that survive in areas of summer rainfall, together perhaps with return migrants from southern Australia, form the basis of the autumn and winter populations
in southern Qld and northern NSW. These long-distance migrations are probably an adaptive response to seasonal or uncertain supplies of food, extreme temperatures (common armyworms do not breed in southern winter or northern summer temperatures) and prevailing winds. In contrast, the southern armyworm is restricted
to southern Australia but there is also strong evidence that this species migrates between regions of favourable habitat (McDonald 1995). Many other pests use related survival strategies. For example, it is most likely that brown cutworm populations, found across most of Australia’s cropping and pasture regions in
N
Spring
Autumn / Winter
Summer
Insect densities Abundant Low to moderate
Fig. 2.45. Likely sources of spring invasions of common armyworm into cropping, pasture and grassland areas of eastern Australia, and possible seasonal shifts in distribution. The size of the autumn/winter refuges may vary with season, with a core area (dark stippling) expanded to a larger area after good rainfall (light stippling). Arrows indicate general direction of migration of adult moths during the early stages of each season.
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winter, migrate southwards in spring. These migrations are likely to be in response to similar seasonal cues to those of the common armyworm, although they are not as temperaturelimited. In contrast, the better known Bogong (or common cutworm) moths move eastwards to mountain refuges during spring and summer, where they can be found densely lining cave
walls. These moths return westwards to the emerging pastures and crops of autumn to produce a single annual generation. Subsequent cutworm (caterpillar) activity in young crops is most evident in early winter. In spring, massive Bogong moth migrations are commonly reported in cities and towns across south-eastern Australia, suggesting that
oversummering sites are not restricted to the eastern Alps, and that migrations are as much north to south as east–west events. Other noctuid moths, including the native budworm (Chapter 3 Cotton) use similar migratory patterns, although the winter distributions and migratory strategies vary in each case.
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PESTS OF STORED GRAINS R.N. Emery and M.K. Nayak
On-farm equipment and seed grain storage (left) can be reservoirs of stored grain insects, and (right) silos near to production areas store grain up to several months before transport to export silos or to local processors. (DAFWA) GROUP Arthropods in stored grain mites book lice beetles Anobiidae Bostrichidae Chrysomelidae
Curculionidae
Dermestidae
Laemophloeidae Lathridiidae Nitidulidae Silvanidae Mycetophagidae Tenebrionidae
moths
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PEST (major pests in bold)
PAGE
flour mite mould mite psocids
42 43 43
cigarette beetle drugstore beetle lesser grain borer cowpea weevil (see Chapter 7) southern cowpea weevil (see Chapter 7) bean weevil (see Chapter 7) pea weevil (see Chapter 8) rice weevil granary weevil maize weevil warehouse beetle khapra beetle (not present in Australia) furniture carpet beetle flat grain beetle minute mould beetles driedfruit beetle sawtoothed grain beetle foreign grain beetle hairy fungus beetle rust-red flour beetle confused flour beetle longheaded flour beetle angoumois grain moth Indian meal moth tropical warehouse moth Mediterranean flour moth
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Damage Harvested grain is at its highest quality when it is first loaded into storage. It can never improve but if the storage environment is not managed correctly the quality will steadily deteriorate. Stored grain insects damage grain by feeding, tainting with their excrement and by increasing temperature and moisture. They can also encourage other species of insects and microflora. Worldwide, grain losses attributed to pests in storage are estimated at 10–30+ per cent. In Australia, dry grain and careful grain storage management has enabled regulations for ‘niltolerance’ of stored grain pests to be established. Management techniques include hygiene, fumigation and insecticides.
a
b
Hygiene The contamination cycle is initiated and continued by insects completing their life cycles in grain that is allowed to remain in and around sheds, headers (particularly), augers, field bins, trucks, animal feed troughs and silos (Fig. 2.46). Good hygiene commences before harvest, when equipment and storage containers must be cleaned of the previous season’s grain, after which contact insecticides may be applied to storage surfaces and equipment to kill insects walking on those surfaces. An inert dust of amorphous silica can also be applied, as a dust or slurry, as a storage and equipment protectant. Careful cleaning of all storage and handling equipment can achieve a ‘clean pipeline’ from the paddock to the silo (Table 2.1). Reduction of temperature or moisture content using cooling aeration or drying are important in slowing the development of stored pests in freshly-harvested grain.
Fumigation When stored grain insects are detected in storage, fumigation with phosphine is the most common and effective treatment used on farms in Australia and is best done in gas-tight silos (Table 2.1). The insects must be exposed to the fumigant for at least 7 days to kill all stages. Fumigation in ordinary, unsealed storages will kill adults but most eggs and pupae will survive to continue the breeding cycle. Phosphine tablets should be applied according to the volume of the silo, whether it is full or not. To delay
Fig. 2.46. (a) Insects breeding in grain residues that have not been cleaned from harvesting machinery, and (b) improperly stored grain may initiate infestation through the chain of grain storage and handling. (ABB Limited)
development of phosphine resistance, there are now national programs to monitor for phosphine resistance and to encourage proper silo sealing. Holding high moisture grain without fumigation in a gas-tight silo can lead to mould spoilage.
Grain-protectant insecticides Where storages are not suitable for use of fumigation, insecticides may be registered in some states for spraying onto cereal grains either to protect the grain against later infestation (contact insecticides; each individual grain must be covered with insecticide) or to kill insects already present in the grain (vapour insecticides). Chemicals are registered only for cereal grains, not oilseeds or pulses. Although some chemicals may be registered for use in some states of Australia, some national and international buyers may not accept grains with any chemical residues (Table 2.1). 41
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Table 2.1. Pest control options for various grains and markets (QDPIF) Cereal grains* for:
on-farm use
markets accepting residual treatments
markets not accepting residual treatments
Pulses** or oilseeds***
Any grain for organic markets
Hygiene
Aeration
Drying
Controlled atmosphere
Phosphine fumigation
Dichlorvos
Treatment
Treatment of storages and equipment
Amorphous silica
?
Residual chemicals
Mixture with grain
Amorphous silica
?
Residual chemicals
*Cereals include: barley, maize, millet, oats, rice, sorghum, triticale, wheat. **Pulses include: faba bean, chickpea, cowpea, field pea, mung bean, navy bean, soybean, pigeon pea. ***Oilseeds include: canola, linseed, safflower, sunflower. can be used. cannot be used. ? some organic markets are reported to accept this treatment, others do not. contact insecticides (malathion excepted) are not permitted on farm-stored grain in Western Australia.
The use of chemicals results in the development of resistance, which occurs rapidly in grain insects, perhaps because most of the population is exposed and there is little immigration of susceptible insects. Resistance may be delayed by using chemicals in accordance with the insecticide label. Sudden treatment failures are anticipated by resistance monitoring.
A R T H RO P O D S I N S TO R E D G R A I N
Flour mite Acarus siro Linnaeus Acarina: Acaridae Distribution: Cosmopolitan, but prevalent in the temperate regions. Pest status: Minor, widespread, irregular. Identification: Adults have eight legs, are less than 0.5 mm long and almost colourless. The first two pairs of legs are widely separated from the hind two pairs and two pairs of long hairs trail at the end of the body. May be confused with: Psocids, which are insects and have six legs.
Host range: This mite infests a range of cereal products including grain and flour. It can also attack cheese and hay. Life cycle: The development of this mite is extremely sensitive to humidity levels outside the range of 75–85%. Optimum conditions for this mite are 28–32°C and 80–85% relative humidity, under which it can multiply sevenfold in 1 week. A female lives up to 60 days and lays 200–250 eggs during this period. Eggs develop through a larval and two nymphal stages (protonymph and deutonymph) to reach adult. The second nymphal stage may be replaced by a special stage known as ‘hypopus’. This stage is highly tolerant of unfavourable conditions, insecticides and fumigation, and may exist for several months without feeding. The hypopi are transported from place to place by clinging to small insects, and when they encounter favourable conditions, resume their normal growth and development to reach adult stage. A complete life cycle from egg to adult takes 9–12 days. Heavy infestation of other stored product pests often creates favourable conditions to mite outbreaks. Risk period: All year.
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Damage: On cereal grain, it attacks the germ and consumes it completely, reducing both germination and nutritive quality. Heavy infestations of this mite emit a ‘minty’ odour and a brownish ‘mite dust’ may appear on open shelving, around the base of flour sacks, on the surface of cheese or in other foods. Such piles consist of dead and living mites, cast skins and faeces. Action level: As required. Chemical control: Fumigation in sealed storage. Cultural control: General storage hygiene can minimise infestations. Lowering the humidity of the storage below 50% and increasing the temperature above 35°C will drastically reduce the mite infestations.
Mould mite or lemon-scented mite Tyrophagus putrescentiae (Schrank) Acarina: Acaridae Distribution: Cosmopolitan, but dominant in stored food in the humid tropics. Pest status: Minor, widespread, irregular. Identification: Adults are pale, greyish white, smooth, wingless, soft-bodied creatures without any antennae. They are microscopic in size (adults are less than 0.5 mm long) and have numerous long hairs on the legs and back. These are not insects because the adults have four pairs of legs and the body is indistinctly divided into two parts.
and involves two nymphal stages. Maximum population growth of 500 times per month can occur under optimal conditions. Risk period: Mostly hot and humid conditions. Damage: This mite causes direct damage by eating the germ of grain and spreading fungi in the commodities. During heavy infestations, the dead and live mites settle as a dust layer on stacks of commodities and floors of stores and emit a lemon-flavoured odour leading to the popular name ‘lemon-scented mite’. The markets reject heavily infested commodities. Cultural control: General storage hygiene can minimise infestations. Lowering the humidity of the storage below 50% and increasing the temperature above 35°C will drastically reduce the mite infestations.
Psocids or booklice Liposcelis spp. Psocoptera: Liposcelidae Distribution: Cosmopolitan, occurs both in temperate and tropical regions with a preference for humid tropics. Pest status: Major, widespread, regular. Identification: Tiny, pale to dark brown, softbodied, up to about 2 mm long with protruding eyes and long thread-like antennae. May be confused with: Flour mites, which have eight legs. Host range: Common in grain stores, mills, central storages, export terminals, processing plants and kitchens. They can infest a range of
May be confused with: Other grain mites. Host range: Mostly occur in damp or moist grain, residues, oilseeds and animal feeds. Can also infest beehives in apiaries, nut-in-shell peanuts, cheese, salami sausage, straw and hay stacks. Life cycle: Like all other mites, this species multiplies rapidly under high humidity conditions (over 65%) with an upper temperature limit of 35–37°C. Under the optimal conditions of 32°C and 98–100% relative humidity, a female lives up to 100 days and lays approximately 500 eggs. The life cycle from egg to adult under these conditions takes 8–12 days
Fig. 2.47. Adult psocid (2 mm long). (SARDI: G. Caon)
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stored commodities including wheat, barley, sorghum, maize, rice, beans, pulses, cassava and oilseeds. Life cycle: Members of Liposcelis spp. specifically, L. bostrychophila Badonnel, L. entomophila (Enderlein), L. decolor (Pearman) and L. paeta Pearman, are the most important pests of stored commodities and in all these species the developmental period from egg to adult roughly takes up to 3 weeks under the optimal conditions of 30°C and 70% relative humidity. Adult females may lay as many as 100 eggs. There are six nymphal stages and they all are similar in appearance to adults but smaller and paler. Liposcelis bostrychophila reproduces without mating, where all nymphs become females. Adult longevity of this species is from 72–144 days, and it is capable of surviving up to 40 days without food. Risk period: Infestations can occur throughout the year, but population outbreaks occur under high humidity conditions after rainfall. Damage: These scavengers contaminate grain but in large numbers can cause economic damage by feeding on the germ of the grain. In damp or high humidity conditions or grain with high moisture content, dense aggregations appear as a moving, slippery carpet of brown dust and endanger workers. Large numbers of dead psocids can cause rejection of contaminated grain. Monitoring: Pest populations can be monitored throughout the year by using corrugated cardboard traps baited with flour in the corrugations. Action level: As required. Chemical control: Fumigation in sealed storage. Treatments applied to the surface of grain can afford some control as adults leave the grain. Storage surface treatments can also reduce the chances of reinfestation. Cultural control: General storage hygiene and fabric treatments can minimise infestations.
Cigarette beetle or tobacco beetle Lasioderma serricorne (Fabricius) Coleoptera: Anobiidae N (Main entry in Chapter 12 Tobacco.)
Pest status in grain: Minor, widespread, regular. Identification: Adult is small, 3–4 mm long, oval, stout, brownish-red, head strongly deflexed under prothorax and with long saw-like (serrate) antennae. The larva is a small, white hairy C-shaped grub. Risk period: All year. Damage: Pupal cocoons and dead bodies of short-lived adults contaminate grain. Monitoring: Light traps and pheromone traps. Internal feeding stages can be detected through X-ray photography and acoustic detectors.
Drugstore beetle Stegobium paniceum (Linnaeus) Coleoptera: Anobiidae Distribution: Cosmopolitan, but more common in temperate regions. Pest status: Major, widespread, regular. Identification: The adults are oval, globular beetles, 3–4 mm long and possess antennae, which are waved rapidly when walking. The antennae have a loose club formed from the last three segments and the elytra have longitudinal grooves. May be confused with: The cigarette beetle, which has a curved underside to the wing covers. Host range: This beetle can attack a wide range of commodities, but prefers processed foods such as chocolate, confectionery and biscuits, and high value commodities such as dried herbs, spices and drugs. Life cycle: Adults live for 2–6 weeks during which a female lays up to 100 eggs loosely on the commodity. On hatching, the scarabaeiform or C-shaped larvae burrow into the commodity, feeding as they go. After passing through four to six instars, the larvae pupate in a cell made out of fragments of food bound up with silk. On emergence, the adult spends a couple of days inside the cocoon before biting its way out. Adults are short-lived and do not feed. They can run quickly and readily fly, and when disturbed they can ‘play dead’ by remaining motionless.
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Under optimal conditions of 30°C and 60–90% relative humidity, the life cycle takes up to 26 days.
May be confused with: The auger beetle, powder-post beetle, and larger grain borer (not in Australia).
Risk period: All year.
Host range: Both an internal and external feeder and is a serious pest of both whole kernel stored grain and cereal products. Primarily a pest of stored wheat and corn, but can infest tobacco, nuts, beans, birdseed, biscuits, cassava, cocoa beans, dried fruit, peanuts, spices, rodenticide baits, and dried meat and fish.
Damage: This pest can cause significant economic damage when it attacks high value processed commodities. Infested materials become contaminated with pupal cocoons and dead bodies of short-lived adults. Mature larvae and adults readily chew through packaging materials leaving holes. Monitoring: This pest can be monitored by using light traps as well as by synthetic pheromone traps. Internal feeding stages can be detected through X-ray photography and acoustic detectors.
Lesser grain borer Rhyzopertha dominica (Fabricius) Coleoptera: Bostrichidae Distribution: Cosmopolitan; occurs in both temperate and tropical regions. In Australia, it is found in all grain-growing states. Pest status: Major, widespread and regular. It is a primary pest of stored grain. Identification: Dark brown in colour and about 2.5–3 mm long. The body is cylindrical and the head is hidden under the round neck-shield. The neck-shield is covered with rasp-like teeth. The elytra are bare, rounded behind, and have distinct rows of pits running their length. The antennae have 10 segments with the last three enlarged, forming a loosely segmented club (Fig. 2.48).
Life cycle: Adults are strong fliers and females lay eggs singly or in groups of up to 30. The eggs are laid on the outside of the grain and a female can lay from 300–500 eggs. Young larvae initially feed on the outside of seed but later stages bore into the grain and remain there. When fully developed, the larvae pupate and adults subsequently bore their way out. A full generation takes 4 weeks at 35°C and 7 weeks at 22°C. Development ceases below 18°C. The optimum temperature is about 31°C. It develops in grain of lower moisture content than most other important pest species and can survive in grain with moisture content as low as 8 or 9 per cent. Population growth rate per month can be up to 20 times. Risk period: All year Damage: Damage caused by this pest is quite prominent and heavy. Adults and larvae bore into undamaged kernels, reducing them to hollow husks. They are also able to survive and develop in the accumulated ‘debris’ produced as the seeds are chewed up (Fig. 2.49).
Fig. 2.48. Adult lesser grain borer (length 3 mm). (DAFWA: R. Graham)
Fig. 2.49. Grain damaged by the lesser grain borer. (DAFWA)
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Monitoring: Pheromone traps placed around the grain. Action level: When detected. Chemical control: Fumigation in sealed storage. Cultural control: This insect does not occur in standing crops, therefore good hygiene with storage and handling equipment should minimise infestations.
during its lifetime. Eggs are laid singly in holes dug in grain and covered with a waxy plug by the female; larva grows inside the grain, excavating a cavity as it grows and pupates inside it. On emergence, the adult may spend a few days inside the cavity and finally chews its way out. The total development period from egg to adult takes about 25 days at optimal conditions of 30°C and 70% relative humidity but this period is greatly prolonged during cold weather. The maximum population growth rate per month is 25 times.
Rice weevil
Risk period: All year
Sitophilus oryzae (Linnaeus) Coleoptera: Curculionidae
Damage: Larval feeding leaves large cavities inside grain and emerging adults leave large emergence holes. Adults feed on the damaged grains and large numbers produce heat and moisture, encouraging mould growth and mites, both of which reduce quality (Fig. 2.51).
Distribution: Cosmopolitan, established in all regions except the coolest temperate regions. In Australia, this pest occurs in all grain-growing states. Pest status: Major, widespread, regular. A primary pest of stored grain. Identification: Adult is 2–2.5 mm in length, head with a snout, varies from reddish brown to nearly black and usually marked on the back with four light reddish or yellowish spots and has fully developed wings beneath the wing covers (Fig. 2.50). May be confused with: Granary weevil, maize weevil. Host range: Major pest of whole cereal grain including wheat, rice and sorghum, but also infests cereal products such as pasta.
Monitoring: Commercially available pitfall and probe traps can either be placed on the grain surface or inserted into the grain bulk. Acoustic detectors and X-ray photography are also available commercially to monitor the developing stages feeding inside the grain. Action level: When detected. Chemical control: Fumigation in sealed storage, grain protectants. Cultural control: Good hygiene with storage and handling equipment should minimise infestations.
Life cycle: Adult weevils can fly and live 4–5 months and each female lays 300–400 eggs
Fig. 2.50. Adult rice weevil (length 2.5 mm). Two pale areas (arrowed) on each wing cover distinguish the species from granary weevil. (DAFWA: R. Graham)
Fig. 2.51. Rice weevil damage to wheat. (DAFWA)
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Granary weevil Sitophilus granarius (Linnaeus) Coleoptera: Curculionidae Distribution: Distributed throughout the temperate regions of the world, uncommon in tropics except in cool upland areas. In Australia, it is restricted to southern temperate regions. Pest status: Major, widespread, regular. Identification: Small (adult about 2.5–3 mm long), blackish or chestnut-brown, head extends into a long slender snout with a pair of stout mandibles or jaws at the end (Fig. 2.52). The characters that distinguish this insect from the closely related rice weevil (Fig. 2.50) are absence of light spots on the wing cover, no wings under the wing covers and the thorax is well marked with longitudinal punctures. May be confused with: Rice weevil, maize weevil. a
Host range: Mostly infest wheat and barley, but can breed on other grains. Life cycle: On average, adult weevils live 7–8 months and a female lays 50–250 eggs during this period. Like the rice weevils, the female granary weevil bores a hole in the grain kernel and deposits an egg and covers it with a gelatinous fluid that seals the hole. After hatching, the larva burrows inside the kernel and grows to a pupa and then to an adult. In warm weather, this weevil takes about 4 weeks to develop from egg to adult and this developmental period prolongs during cold weather. Maximum population growth rate per month can be up to 25 times. Risk period: All year. Damage: Larvae eat large cavities inside grain and by emerging adults leave large emergence holes. Adults feed on damaged grains (Fig. 2.53), large numbers of which produce heat and moisture, encouraging mould growth and mites. Monitoring: This pest can be monitored with a range of commercially available pitfall and probe traps that can either be placed on the grain surface or inserted into grain bulk. Acoustic detectors and X-ray photography are also available commercially to monitor the developing stages feeding inside the grain. Action level: When detected.
b
Fig. 2.52. (a) Adult granary weevil (3.0 mm long) which has no yellow areas in its wings, and (b) detail of the snout of the adult weevil. (DAFWA: R. Graham)
Chemical control: Fumigation in sealed storage.
Fig. 2.53. Adult granary weevils feed on damaged grain. Larvae are primary damagers of grain. (DAFWA)
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Cultural control: Good hygiene with storage and handling equipment minimises infestations.
Maize weevil Sitophilus zeamais Motschulsky Coleoptera: Curculionidae Distribution: Cosmopolitan, more prevalent in the tropics. In Australia, it is found in northern sub-tropical grain growing areas.
probe traps, which can either be placed on the grain surface or inserted into grain bulk. Acoustic detectors and X-ray photography are also available commercially to monitor the developing stages feeding inside the grain. Action level: When detected. Chemical control: Fumigation in sealed storage. Cultural control: Good hygiene with storage and handling equipment should minimise infestations.
Pest status: Major, widespread, regular. Identification: Adult is 2–3 mm in length, head has a snout, varies from reddish-brown to nearly black and is usually marked on the back with four light reddish or yellowish spots, has fully developed wings beneath the wing covers. With these characters, the maize weevils are indistinguishable from the rice weevils by external characteristics. Only through dissection and examination of genitalia, can these two pests be distinguished from each other. May be confused with: Rice weevil, granary weevil. Host range: Predominantly found on maize, but can infest other cereals including milled and paddy rice. Life cycle: The adults are long-lived (9 months to 1 year), and a single female lays up to 150 eggs during its lifetime. Eggs are laid individually in small cavities chewed into the grains by the female. Each cavity sealed and the egg is thus protected by a sticky secretion. Eggs hatch in 6 days and, after passing through four instars, the larvae pupate inside the grain. The total life cycle from egg to adult takes up to 35 days under optimal conditions of 27ºC and 70% relative humidity.
Warehouse beetle Trogoderma variabile Ballion Coleoptera: Dermestidae Distribution: Believed to have originated in central Asia, now established throughout the northern hemisphere, including southern Europe, south-west Asia, Russia, Mongolia, China, Canada and the USA. It occurs in southern Australia. Pest status: Major, widespread, regular. Identification: Adult is 2–3 mm long, oval, brown, hairy, has wing cases with irregular pale markings; larvae are hairy (Fig. 2.54). May be confused with: Khapra beetle (not found in Australia), other native and pest Trogoderma as well as some carpet beetles. Separation of larvae must be done by trained persons. Host range: Cereal grain, rice, canola, pulses, and any other dried plant or animal material.
Risk period: All year. Damage: Extensive damage is done to the commodity by feeding larvae, which leave large cavities inside grain, and by emerging adults, which leave behind large emergence holes. Adults feed on the damaged grains and high numbers produce heat and moisture, both of which encourage mould growth and reduce quality. Monitoring: This pest can be monitored with a range of commercially available pitfall and
Fig. 2.54. Larvae of warehouse beetle on canola seed. (DAFWA)
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Life cycle: Eggs are laid randomly; larvae pass through five to six instars and pupation. Under optimal conditions of 32ºC and 70% relative humidity, the total life cycle from egg to adult takes 27 days. Maximum population growth rate per month can be up to eight times. Risk period: All year, although adults are more active during summer. Damage: Large quantities of cast larval skins can accumulate in and around infested grain and cause allergenic conditions in workers. Widespread infestations could mask an incursion of khapra beetle. Monitoring: This pest can be monitored by using pheromone traps with commercially available synthetic aggregation pheromone. Adults fly, so traps may be hung in nearby trees. Action level: When detected. Chemical control: Fumigation in sealed storage, protectants. Cultural control: This insect does not occur in standing crops therefore good hygiene with storage and handling equipment should minimise infestations.
fish, skin, woollen articles and museum specimens. Life cycle: Under favourable conditions of 35ºC and 75% relative humidity, the eggs pass through five to six larval stages to become adults. Under adverse conditions, some larvae can undergo a diapause state for several months to years. Maximum population growth rate per month can be up to 13 times. Risk period: All year. Damage: This is the world’s worst pest of stored products. Grain damaged by this pest has somewhat the same appearance as grain attacked by the lesser grain borer. Under hot and humid conditions, populations of this pest can cause total loss of infested commodities. Large quantities of cast larval skins can accumulate in and around infested materials and cause allergenic conditions in workers. Monitoring: Commercially available pheromone traps can be used to monitor this pest. Adults do not fly so traps must be placed on a surface. Action level: When detected. Chemical control: Extended fumigation in sealed storage, protectants. Cultural control: None are effective.
Khapra beetle Trogoderma granarium Everts Coleoptera: Dermestidae Distribution: Believed to have originated in India, now established in hot dry areas of the Middle East, Africa, South Asia and also in protected environments in countries with unfavourable climate. It has not been reported from Australia. Pest status: Major, widespread (in some countries), regular. Identification: Adult is an oval, reddish-brown, hairy, 2–3 mm long beetle. Larvae are very hairy and grow up to 5 mm. May be confused with: Warehouse beetle, several other native and pest Trogoderma as well as some species of carpet beetle from which it can only be distinguished by dissection. Host range: Infest a wide range of commodities, including cereal grain, oilseeds, pulses, dried
Host-plant resistance: None is known.
Furniture carpet beetle Anthrenus flavipes LeConte Coleoptera: Dermestidae Distribution: Cosmopolitan, originated in Oriental regions, but now widespread in tropical and subtropical parts of the world. Pest status: Minor, widespread, regular. Identification: Adult is a small (2–3 mm long), oval, ‘seed-like’ beetle. When alarmed, the insect tucks in its legs and antennae. The upper surface of the adult body is covered with scales, occasionally with a pattern. Larvae are about 5 mm long, hairy with long tufts extending from the last abdominal segments (Fig. 2.55). May be confused with: warehouse beetle and also Khapra beetle from which it can only be distinguished by dissection. 49
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Pest status: Major, widespread, regular. Identification: Adult is up to 2 mm long, very flattened, light brown with elongate antennae that are about two-thirds as long as the body (Fig. 2.56). May be confused with: Saw-toothed grain beetle and foreign grain beetle, which are not flattened.
Fig. 2.55. Carpet beetle larvae (each about 5 mm long). (SARDI: G.J. Baker)
Host range: This pest attacks mostly dried artefacts of animal origin, including preserved museum specimens, skins, hides and woollen goods including clothes, carpets and untreated woollen insulation. Life cycle: Adults live on average 2–4 weeks, during which a female lays 30–100 eggs. On hatching, larvae feed and burrow through the commodity and moult several times. Under adverse conditions the larvae undergo diapause and, in temperate regions, they usually overwinter in this state. Pupation takes place inside the skin of the last larval instar. On emergence, adults feed on nectar and pollen from flowers. Under optimal conditions of 35°C and 70–80% relative humidity, the life cycle is completed in 70–80 days.
Host range: Grain and grain products. They prefer broken grain and, being highly flattened, they can enter very small cracks and crevices in grains and they can enter well-packaged processed food. They can also infest oilseed cake, dates and other dried fruits, often following infestation by other insects. Life cycle: A single female lays up to 200 eggs and, after four larval instars, pupation occurs in a silk cocoon. Total life cycle takes up to 3 weeks under optimal conditions of 33°C and 70% relative humidity. Adults are fast moving and do not fly. Maximum population growth rate per month is 60 times. Damage: A secondary pest, the larvae attack the germ, thus reducing the percentage germination a
Risk period: All year. Damage: Damage is caused by larvae, which bore holes in infested materials and packing materials. Significant economic loss can be caused in museums and in households. Monitoring: This pest can be monitored by using commercially available synthetic pheromone traps.
b
Flat grain beetle Cryptolestes spp. Coleoptera: Laemophloeidae Distribution: Cosmopolitan with a range extending from the tropics to temperate regions.
Fig. 2.56. (a) Flat grain beetles in stored food, and (b) adult beetle, 2 mm long. (DAFWA: R.Graham)
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of seed grain and causing weight loss and loss of quality in food grain. Monitoring: Both pitfall and crevice traps can be used to monitor this pest. Action level: When detected. Chemical control: Fumigation in sealed storage, protectants. Cultural control: This insect does not occur in standing crops therefore good hygeine with storage and handling equipment should minimise infestations.
Minute mould beetles Corticaria spp. Coleoptera: Lathridiidae Distribution: Cosmopolitan, widespread in temperate regions. Pest status: Minor, widespread, regular. Identification: Adult is very small in size (1.3–2 mm), hairy and brown in colour. May be confused with: Hairy fungus beetle which is yellowish. Host range: Larvae and adults feed on mould and generally do not attack stored commodities directly. Life cycle: Adult is long-lived and, under optimal conditions of high humidity and 18°C, the life cycle is completed within 40 days. Risk period: All year. Damage: This insect does not cause any significant damage to the stored commodities as both adults and larvae mostly feed on mould. It does not breed or persist in dry commodities stored under good conditions. Monitoring: Both pitfall traps and crevice traps can be used for successful monitoring of this pest.
Driedfruit beetle Carpophilus hemipterus (Linnaeus) Coleoptera: Nitidulidae Distribution: Cosmopolitan, widespread in warm-temperate to tropical regions. In temperate regions, it occurs mostly in sheltered or heated environments.
Pest status: Minor, widespread, regular. Identification: Adult is a flattened and oblong beetle, 3–4 mm in length and light brown to black in colour. Elytra or the wing cases do not completely cover the abdomen and have large yellow spots. May be confused with: Many other Carpophilus species. Host range: Serious pests of dried fruits but can infest cereals, particularly maize, before harvest and a range of fruit and vegetable matter, especially if it is damp and decomposing. Life cycle: Adults live for an average of 3 months, but can live up to a year. A female lays on an average 1000 eggs, which hatch in 2–3 days and the larval stage lasts 6–8 days and pupation up to 5–7 days. Under optimum conditions of 32ºC and 70% relative humidity, the life cycle takes only 12 days. Damage: This pest mostly flourishes under damp mouldy conditions and often acts as a vector of various spoilage fungi and bacteria, both in storage and in the field. The beetle feeds on the flesh of the dried fruits and on any moulds and yeasts present in the commodity. Damage is not readily identifiable as being specifically caused by this pest. Monitoring: Flight traps with synthetic aggregation pheromone and/or food bait such as whole wheat bread dough or fermenting fruit juice can be used to monitor this pest. Action level: As required. Chemical control: Fumigation in sealed storage. Cultural control: General storage hygiene can minimise infestations.
Sawtoothed grain beetle Oryzaephilus surinamensis (Linnaeus) Coleoptera: Silvanidae Distribution: Cosmopolitan. Pest status: Major, widespread, regular. Identification: Adult is up to 3 mm long, slender, flat, dark brown to grey with six sawtooth-like projections on each side of thorax (Fig. 2.57). 51
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a
Risk period: All year. Damage: A secondary pest, the larvae feed on the damaged and broken kernels with a preference for germ. Larvae also attack germ in whole cereal grains, thereby changing the nutritional content and reducing the percentage germination. Adults and larvae are able to enter small cracks, so they can often attack packaged food or nuts in shell. Monitoring: This pest can be monitored by using corrugated cardboard traps baited with flour sprinkled inside the corrugations. Action level: When detected. Chemical control: Fumigation in sealed storage, protectants.
b
Cultural control: This insect does not occur in standing crops therefore good hygiene with storage and handling equipment should minimise infestations.
Foreign grain beetle Ahasverus advena (Waltl) Coleoptera: Silvanidae Distribution: Cosmopolitan, most common in tropical Africa and Asia, can thrive successfully in unheated locations in cool temperate regions such as northern Europe. Fig. 2.57. (a) Adult sawtoothed grain beetles (3 mm long), and (b) the sawtooth projections on thorax. White bar spans 0.2 mm. (DAFWA: R. Graham)
May be confused with: Merchant grain beetle and flat grain beetle, which do not have the sawtooth projections. Host range: Both adults and larvae attack grain, flour, meals, breakfast cereals, stock and poultry feeds, copra and dried fruits. Life cycle: Adult lives on average for 6–10 months but some may live as long as 3 years. Each female beetle lays 40–280 eggs during its lifetime, dropping them loosely among the foodstuff or inserting them into the crevice of a grain kernel. Eggs hatch in 3–5 days and the larvae feed actively. The developmental period from egg to adult takes up to 3 weeks under optimal conditions of 33ºC and 80% relative humidity. Adults are fast moving and rarely fly. Maximum population growth rate per month is 50 times.
Pest status: Minor, widespread, regular. Identification: The adult is small (2–3 mm long) but robust with stouter body and a wide prothorax, which has a single blunt tooth at each front angle. The antennae are clubbed and moderately long. May be confused with: Hairy fungus beetle and also flat grain beetle, which do not have the ‘teeth’ at the front of the wing covers. Host range: This is a minor secondary pest of a wide range of stored commodities including cereal grains, oilseeds, copra, groundnuts, dried fruits, herbs and cocoa beans. The numbers become abundant when the commodity is mouldy. Life cycle: Adults do not breed below 65% relative humidity and a female lays several hundred of eggs during its life span among the substrate or in cracks and crevices in grains. The life cycle takes about 22–30 days under optimal conditions of 27°C and 75% relative humidity.
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Risk period: All year. Damage: This pest flourishes under damp mouldy conditions, and both larvae and adults are general feeders. Damage is not readily identifiable as being specifically caused by this insect. Monitoring: This pest can be monitored by using commercially available food baits based on grain oil.
Hairy fungus beetle Typhaea stercorea (Linnaeus) Coleoptera: Mycetophagidae Distribution: Cosmopolitan, but is most common in the humid tropics. Pest status: Minor, widespread, regular Identification: The adult is a hairy, light to dark brown coloured oblong beetle, about 2.5–3 mm long. The antennae have a threesegmented club. May be confused with: Minute mould beetle which is a darker brown. Host range: This pest is mostly a minor pest of freshly harvested or slightly damp grain. Life cycle: Adults are long lived and females lay eggs randomly or loosely attach them to grains. The translucent larvae are very mobile and move through the commodity. Under optimal conditions of 25°C and 80–90% relative humidity, the life cycle is completed between 21–33 days. Risk period: All year. Damage: This pest flourishes under damp mouldy conditions and both larvae and adults are general feeders. Damage is not readily identifiable as being specifically caused by this insect. Monitoring: This pest can be monitored by using commercially available pitfall and crevice traps.
Rust-red flour beetle Tribolium castaneum (Herbst) Coleoptera: Tenebrionidae
Fig. 2.58. Damage by rust-red flour beetles (4 mm long) is not readily identifiable. (DAFWA)
Distribution: Cosmopolitan. It is thought to have originated in India, but now found throughout all tropical, subtropical and warm temperate regions of the world. It is found in all grain growing states of Australia. Pest status: Major, widespread, regular. Identification: The adult is 3–4 mm long, flattened, reddish-brown (Fig. 2.58); larvae are elongate and light brown. The last three segments of the antennae are abruptly much larger than the other segments, forming enlarged tips. The head margins are nearly continuous at the eyes and do not have a ridge over the eye. The gap between the eyes is equal to one eye diameter and the pronotum is widest at the middle (Fig. 2.59). May be confused with: Confused flour beetle and longheaded flour beetle, neither of which have the three-segmented antennal club. Host range: Mainly a pest of milled cereals, but occurs widely in stored whole grain, animal feed and processed food. Can also infest groundnuts, nuts, spices, coffee, cocoa, dried fruits and occasionally peas and beans. It is considered a secondary pest because it cannot damage sound grain. Life cycle: Adults are strong fliers and live for many months, even several years, under temperate conditions. The adult female lays 2–10 eggs each day throughout its life and larvae are quite active. Development of an egg to adult takes up to 21 days under optimal conditions of 35ºC and 75% relative humidity. Damage: See confused flour beetle. 53
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a
b
c
d
Fig. 2.59. Distinguishing the rust-red flour beetle (a, b) from the confused flour beetle (c, d). The antennae of the rust-red flour beetle end in a distinctive three-segmented club (a) and, in the ventral view (b) the eyes are about one eye diameter apart. The confused flour beetle lacks distinctively clubbed antennae (c) and the eyes in ventral view are about two and a half eye diameters apart (d) (DAFWA: R.Graham) . Scale: horizontal white bar is 0.5 mm.
Monitoring: This pest can be easily monitored by using pitfall, crevice and pheromone traps.
Pest status: Minor, widespread, regular.
Cultural control: This insect does not occur in standing crops, therefore good hygiene with storage and handling equipment should minimise infestations.
Identification: The adult is a 3.5–4 mm long, shiny, flattened, oval, reddish-brown beetle (Fig. 2.60). The head and upper parts of the thorax are densely covered with minute punctures; the head margins are expanded and notched at the eyes, with a ridge over the eye (2.59c). The gap between the eyes is equal to 2.5 eye diameters and the pronotum is widest at the anterior third (Fig. 2.59d).
Confused flour beetle
May be confused with: The rust-red flour beetle, which has the three-segmented antennal club.
Action level: When detected. Chemical control: Fumigation in sealed storage, protectants and dichlorvos.
Tribolium confusum Jaquelin du Val Coleoptera: Tenebrionidae Distribution: Believed to have originated in Ethiopia, uncommon in tropical regions, but widespread in cool temperate zones.
Host range: A secondary (i.e. only attacks damaged grain) pest and pest of milled cereals, whole grain, animal feed and processed food. Life cycle: The average life of this pest is about 1 year, but there are records of some individuals
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regions, it occurs mostly in sheltered or heated environments. Pest status: Minor, widespread, regular Identification: The adult is about 3 mm long, slender, flattened and yellowish to brown coloured beetles. The antennae are short, with a five-segmented club.
Fig. 2.60. Adult confused flour beetles (4 mm long). (DAFWA)
living as long as 3 years. Adults do not fly and the female lays an average of about 450 eggs, loosely in flour or other food materials in which adults live. The eggs are covered with sticky secretion and hatch in 5–12 days. Larvae are slender, cylindrical and wormlike, and when fully grown transform into small naked pupae. In summer (35ºC and 75% relative humidity), the developmental period from egg to adult takes about 6 weeks, whereas the life cycle is greatly prolonged by cold weather. The maximum population growth rate per month can be up to 60 times. Risk period: All year. Damage: Both larvae and adults are general feeders and the damage caused by them is not readily identifiable. A heavy infestation in commodities discolours grain and emits a foul odour due to a secretion from the abdominal glands of the adult.
May be confused with: Rust-red flour beetle and confused flour beetle, neither of which have the five-segmented antennal club. Host range: This is a secondary pest of cereals and oilseeds. Life cycle: The female lays eggs randomly in the food media, which hatch in about 4 days. The larval stage lasts about 15 days with approximately seven moults, and the pupal period lasts up to 4 days. Under optimal conditions of 35°C and 85% relative humidity, the life cycle is completed in 22 days and, under these conditions, this pest has a population growth rate of 10 times per month. Risk period: All year. Damage: Damage is not readily identifiable as being specifically caused by this pest. Infestation can lead to persistent foul odours in the commodity due to chemical secretions from abdominal glands of the pest. Monitoring: This pest can be monitored by using pitfall and crevice traps.
Monitoring: This pest can easily be monitored by using pitfall, crevice and pheromone traps.
Angoumois grain moth
Action level: When detected.
Sitotroga cerealella (Olivier) Lepidoptera: Gelechiidae
Chemical control: Fumigation in sealed storage, protectants and surface treatments. Cultural control: This insect does not occur in standing crops, therefore good hygiene with storage and handling equipment should minimise infestations.
Distribution: Cosmopolitan, widely distributed in mild and warm temperate to tropical regions. Pest status: Major, widespread, irregular.
Longheaded flour beetle
Identification: The adult is small, 5–6 mm long with a wingspan of 12 mm. The forewings are yellowish-brown with small black spot towards the tip; hind-wings have a long fringe of hairs and are sharply pointed at the tip (Fig. 2.61).
Latheticus oryzae Waterhouse Coleoptera: Tenebrionidae
May be confused with: Mediterranean flour moth and Ephestia species.
Distribution: Cosmopolitan, but most common in the tropics, particularly Asia. In temperate
Host range: An important primary pest of whole cereal grains including maize, wheat, rice, 55
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Cultural control: General storage hygiene can minimise infestations.
Indian meal moth Plodia interpunctella (Hübner) Lepidoptera: Pyralidae Distribution: Cosmopolitan in warm climates and can survive in heated buildings in cool temperate countries. Pest status: Minor, widespread, regular. Fig. 2.61. Adult angoumois grain moths (6 mm long) showing black-tipped wings. (DAFWA)
sorghum and millet. It can infest the grain both in the field and in storage, with a preference for sorghum and maize. Life cycle: Adults live 5–10 days and, during this period, a single female lays up to 200 eggs. The eggs are laid on or near grain, and after hatching the larvae bore into the grain. Each larva completes its development in about 19 days, entirely within a single grain and spins a silken cocoon for pupation. Adults do not penetrate far into bulk grain. Under optimal conditions of 30°C and 75% relative humidity, the development period from egg to adult takes 25–30 days. The maximum population growth rate per month can be up to 50 times. Risk period: Generally summer months. Damage: This moth can infest all grains both in the field and in storage. It prefers damp grain and is most serious when these grains are harvested without threshing or shelling. Larvae eat large cavities within the infested grain. It is a serious pest of bagged and traditionally stored produce or commodities stored on-farm. Heavy infestations from this pest can produce heat and moisture in the dry grain encouraging the development of mould growth and resulting in quality loss.
Identification: The adult moth is up to 10 mm in length with a wingspan of 18–20 mm (Fig. 2.62). The forewing of the adult is cream-coloured in the basal one-third and the rest of the wing is dark reddish-brown with some grey marking. The full-grown larva is 12.5 mm long and dirty white in colour, sometimes varying to greenish and pinkish hues. May be confused with: Mediterranean meal moth and warehouse moths (Cadra spp.), especially when specimens have wing scales rubbed off. Host range: Infests a range of stored grain including wheat, barley, sorghum, oats, rice, maize, buckwheat and rye. It also infests dried fruits such as raisins, apricots, almonds, peanuts, hazelnuts, walnuts and many processed products. Life cycle: The adult moth is short lived (1–2 weeks). Each female lays 100–300 eggs, singly or in groups directly on food material or on its packaging. Eggs hatch in 3–4 days and undergo four to seven instars before spinning cocoons for
Monitoring: Traps with synthetic pheromones are generally used to monitor this pest. Action level: When detected. Chemical control: Fumigation in sealed storage. Protectants applied to the surface of the grain can afford some control as adults leave the grain.
Fig. 2.62. Freshly emerged adult of the Indian meal moth. (DAFWA)
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pupation in the food material or just below the surface. Under optimal conditions of 30°C and 75% relative humidity, the development period from egg to adult takes 4 weeks. The maximum population growth rate per month can be up to 60 times. Risk period: All year. Damage: This pest causes serious economic damage to processed, packaged and manufactured food and confectionary products. Adults do not feed and larvae cause most of the damage by eating the embryo of the grain and milled cereal products, nuts, dried fruits and chocolates, etc. Larvae always leave silken threads or webbing that can produce foul flavour, block machinery and act as a harbourage for other insect pests. The ability of larvae to bore into packaged food materials has made this pest a major concern for retail industry, trade and domestic situations. Monitoring: Pheromone-baited traps can be used to monitor this pest. Action level: As required. Chemical control: Fumigation in sealed storage. Treatments applied to the surface of grain can afford some control as adults leave the grain. DDVP vapour at dusk also helps. Cultural control: General storage hygiene can minimise infestations.
Tropical warehouse moth Cadra cautella (Walker) Lepidoptera: Pyralidae Distribution: Distributed predominantly throughout the tropics and subtropics, in temperate regions, however it needs a heated environment to survive the winter. Pest status: Minor, widespread, regular. Identification: The adult is 8–10 mm long with a wingspan of 15–22 mm. The forewings are reddish-brown with fairly coloured white crosslines and the hind wings are pale grey. Fullgrown larvae are 12–15 mm long, pale grey with many setae and small dark spots and a dark head capsule (Fig. 2.63). May be confused with: Other species of Pyralidae.
Fig. 2.63. Larva of a tropical warehouse moth (15 mm long) showing dark spots along the body. (DAFWA)
Host range: It is a major pest of flourmills and food-processing plants. It can infest a range of commodities including flours, grains, dates, dried fruits, cocoa beans, nuts, spices and seeds. Life cycle: Adults are short lived (maximum up to 2 weeks), and during this period a single female lays up to 250 eggs loosely on the surface of the commodity. Eggs hatch in about 3 days and the larval development is completed in about 3 weeks, which involves five instars. The mature larva wanders and seeks a suitable place to pupate, such as the walls of the store or the spaces between bags. Pupation takes place inside a silken cocoon prepared by the mature larva. Under optimal conditions of 30ºC and 75% relative humidity, the development period from egg to adult takes 25–30 days. The maximum population growth rate per month can be up to 50 times. Risk period: Generally summer months. Damage: Consumption by larvae is the main type of damage, but frass and silk-filled galleries cause significant contamination of the commodity. Monitoring: Adults can be monitored by using sticky traps baited with synthetic sex pheromones. Refuse traps can be used to monitor larvae. Action level: As required. 57
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Occasional contaminants of grains include various snail species, the bronze field beetle, vegetable beetle, spring beetles Liparetrus spp., carab beetles, ladybirds of various species, the desiantha weevil, European earwig, brown
shield bug, coon bug, and Rutherglen bug. These insects are accidental contaminants collected from the field by the harvester or by insects sheltering in windrowed crops. They cause no damage to stored grain although some of the
Chemical control: Fumigation in sealed storage. Treatments applied to the surface of the grain can afford some control as adults leave the grain. Cultural control: General storage hygiene can minimise infestations.
Mediterranean flour moth Ephestia kuehniella Zeller Lepidoptera: Pyralidae Distribution: Cosmopolitan, but most prevalent in temperate regions. Pest status: Minor, widespread, regular. Identification: Adults 5–7 mm long with a wingspan of 15–16 mm. The forewings are brownish-grey, crossed with two light coloured bands, and the hind wings are uniformly grey. Mature larvae are 10–15 mm long, creamy white shaded with yellow brown or pink. Pupae turn from light brown to black immediately before adult emergence. May be confused with: other Ephestia species and the Indian meal moth.
(true) bugs may taint grain by their odour. Contaminants may cause grain to be rejected on delivery at storage depots. Any insects in export shipments may be rejected by importing countries.
Host range: Infests a range of stored commodities including cereals, processed flours, tobacco, chocolate, cocoa beans, coffee, cottonseed, dried fruits, nuts, seeds and shelled maize. Life cycle: Adults live up to 3 weeks and during this period a single female lays 150–200 eggs, mostly during night time. Under optimal conditions of 25°C and 70% relative humidity, the development period from egg to adult takes 6–7 weeks. Risk period: Generally summer months. Damage: Severe infestations from this moth pest in flourmills can cause webbed lumps in the flour drastically reducing its quality, and these may clog the mill machinery. Action level: As required. Chemical control: Fumigation in sealed storage. Treatments applied to surface of grain can afford some control as adults leave the grain. Cultural control: General storage hygiene can minimise infestations.
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Beneficial organisms commonly found in Australian grain stores
Pseudoscorpions various spp. Arachnida: Scorpionida Pseudoscorpions are similar in appearance to scorpions, however they lack the tail and sting. They are flattened, oval shaped, and only grow to about 1–5 mm. They have large scorpion-like claws and four pairs of legs, and possess poison glands at the base of their pincers. They are usually brown with some species lacking eyes. Juvenile stages may be confused with mites. Pseudoscorpions are predators often found in moist grain. They feed on insect eggs, spring-tails, booklice, mites, small larvae and silverfish. They ingest their meal by anaesthetising and liquefying it so that they can suck it up through special grooves in their mouthparts. Prior to mating, the adults often undergo a courtship dance involving posturing, various dance steps, and grasping with the pedipalps. Following courtship, the male places a spermatophore onto the substrate for the female to retrieve. Females place the fertilised eggs into a brood sac which is attached to their genitalia. The female provides nourishment to the embryos in the form of a nutritive fluid and the young remain in the brood sac throughout the protonymphal
stage. Two additional nymphal stages occur via moulting before maturity is reached.
Mites Acarina: Acaridae One species of mite, Cheyletus malaccensis Oudemans is sometimes found in tropical storages and is both beneficial and a nuisance to grain workers. Adults of the mite are relatively large (about 0.5 mm), whitish in colour, usually with a typical white stripe along the middle of the body, have long legs and prominent mouthparts with large mandibles. These mites prey on mould mites and eggs, larvae and nymphs of other stored product pests including psocids. The eggs of this predatory mite hatch and develop through a larval and two nymphal stages before emerging as adults. Under optimal conditions of 30°C and 75% relative humidity, the life cycle is completed in 19–20 days. These mites can also reproduce parthenogenetically (in absence of male forms). Other predator mites sometimes found in grains include: Pyemotes ventricosus (Newport), Acarophenax triboli Newstead and Duval, Pyemotes tritici (Lagreze-Fossat and Montagne), Acaropsis docta Berlese, Chortoglyphus gracilipes Banks and Blattisocius tarsalis (Berlese).
Parasitic wasps Choetospila elegans Westwood, Anisopteromalus calandrae (Howard), Lariophagus distinguendus (Foster), Pteromalus semotus (Walker), Bracon hebetor Say, Trichogramma evanescens Westwood, Holepyris sylvanids (Brethes), Cephalonomia meridionalis Brethes, Cephalonomia waterstoni Gahan, Dinarmus basalis (Rondani), Israelius carthami Richards and Cephalonomia gallicola (Ashmead) and Zeteticontus utilis Noyes.
Predatory bugs Xylocoris flavipes (Reuter) and Amphibolus venator (Klug).
Protozoans Nosema oryzaephili Burges, Canning and Hurst; Mattesia oryzaephili Ormiers, Louis and Kuhl; Mattesia dispora (Weiser), Adelina triboli Bhatia, Nosema heterosporum Kellen and Lindegren, N. plodiae Kellen and Lindegren.
Bacteria Bacillus thuringiensis Berliner.
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Sources of information Allen, P.G. (1973). Biology of Desiantha caudata Pascoe (Coleoptera: Curculionidae) in South Australia. Journal of the Australian Entomological Society 12: 201–206. Allen, P. (1986a). Blackheaded pasture cockchafer. Department of Agriculture South Australia and Technical and Further Education, Fact Sheet FS 3/86. Allen, P. (1986b). Pasture webworm. Department of Agriculture South Australia and Technical and Further Education, Fact Sheet FS 25/85. Allen, P. (1986c). Yellowheaded cockchafer. Department of Agriculture South Australia and Technical and Further Education, Fact Sheet FS 23/85. Baker, G.H. (1986). The biology and control of white snails (Mollusca: Helicidae), introduced pests in Australia. Division of Entomology Technical Paper No. 25. Broadley, R.H. (1979). Armyworms in south Queensland field crops. Queensland Agricultural Journal 105: 433–443. Broadley, R.H. (1986). Parasitism of Mythimna convecta (Walker) (Lepidoptera: Noctuidae) larvae in South East Queensland. Journal of the Australian Entomological Society 25: 61–62. Burgess, E.P.J. (1987). Population dynamics of Mythimna separata and its parasitoid, Cotesia ruficrus, on maize in New Zealand. New Zealand Journal of Agricultural Research 30: 203–208. Calder, A.A. (1996). Click Beetles: Genera of Australian Elateridae (Coleoptera). Monographs on Invertebrate Taxonomy, Volume 2. CSIRO Publishing, Collingwood. Carver, M. (1984). Metopolophium dirhodum (Walker) (Homoptera: Aphididae) newly recorded from Australia. Journal of the Australian Entomological Society 23: 192. Cereal aphid forecasts: and search for ‘cereal aphid forecast’. Chapman, R.B. (1984). Pasture pests. In: New Zealand Pest and Beneficial Species (Scott, R., ed.). Lincoln University. pp. 121–122. Common, I.F.B. (1954). A study of the ecology of the adult bogong moth, Agrotis infusa (Boisd.) (Lepidoptera: Noctuidae), with special reference to its behaviour during migration and aestivation. Australian Journal of Zoology 2: 86–89. Common, I.F.B. (1993). Moths of Australia. Melbourne University Press, Melbourne. Edwards, E.D. (1992). A second sugarcane armyworm (Leucania loreyi) (Dupobchel) from Australia and the identity of L. loreyimima Rungs (Lepidoptera: Noctuidae). Journal of the Australian Entomological Society 31: 105–108. Elder, R.J., Brough, E.J. and Beavis, C.H.S. (1992). Managing Insects and Mites in Field Crops, Forage Crops and Pastures. QDPI Brisbane, Brisbane. 143pp. Emery, R.N. and Szito, A. (1999). Western Australian native cockchafers can become serious pests of agricultural crops. 7th Australasian Conference on Grassland Invertebrate Ecology, Perth, Western Australia, 4–6 October 1999. Emery, R., Mangano, P. and Michael, P. (2005). Crop Insects: The Ute Guide, Western Grain Belt Edition. Western Australia Department of Agriculture and Grains Research and Development Corporation. Goodyer, G. (1985). Cutworm caterpillars. Department of Agriculture New South Wales Agfact AE.40. Goodyer, G. (1993). Pests of winter cereals. Department of Agriculture New South Wales Agfact P3. AE.3.
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Goodyer, G. (Undated). Identifying major noctuid caterpillars. New South Wales Agriculture and Rhone-Poulenc. Booklet. 16pp. Hadlington, P. and Gerozisis, J. (1995). Urban Pest Control in Australia. UNSW Press, Sydney. 294pp. 3rd rev. edition. Hill, M.G. (1988). Analysis of the biological control of Mythimna separata Lepidoptera: Noctuidae) by Apanteles ruficrus (Braconidae: Hymenoptera) in New Zealand. Journal of Applied Ecology 25: 197–208. Hopkins, D. (1982). Cutworms. Department of Agriculture South Australia, Fact Sheet FS 44/81. Hopkins, D. (1987). Cereal aphids. Department of Agriculture South Australia and Technical and Further Education, Fact Sheet FS 42/85. Hopkins, D. (1996). Cereal curculio and maculate curculio. Primary Industries SA, Fact Sheet. FS 22/85/96. Hopkins, D. (1999). Armyworm control in cereals. Primary Industries and Resources SA Fact Sheet. Agdex 110/622. Hopkins, D. and Miles, M. (1998). Insects: the GRDC Ute Guide for the Southern Region. Primary Industries and Resources, South Australia. Hopkins, D., Leyson, M., Baker, G., Charwat, S., Cavagnaro, V., Keller, M., Rohitha, H., Dillon, T., Long, B., Mayfield, A., Desboilles, J., Heidenreich, C., Sharma, S., and Richards, M. (2003). Bash ’Em Burn ’Em Bait ’Em: Integrated snail management in crops and pastures. South Australian Research and Development Institute. 40pp. Horne, P.A. and Edward, C.L. (1995). Phenology and food preferences of Labidura truncata Kirby (Dermaptera: Labiduridae) in western Victoria. Journal of the Australian Entomological Society 34:101–104. Leyson, M., Hopkins, D.C., Charwat, S., Baker, G. (2003). Release and establishment in South Australia of Sarcophaga penicillata (Diptera: Sarcophagidae), a biological control agent for Cochlicella acuta (Mollusca: Hygromiidae). 2003 BCPC Symposium Proceedings No. 80: Slugs & Snails: Agricultural, Veterinary & Environmental Perspectives, pp. 295–300. May, B.M. (1977). Immature stages of Curculionidae: Larvae of the soil-dwelling weevils of New Zealand. Journal of the Royal Society of New Zealand 7: 189–228. McDonald, G. (1991a). Oviposition and larval dispersal of the common armyworm, Mythimna convecta (Walker) (Lepidoptera: Noctuidae). Australian Journal of Ecology 16: 385–393. McDonald, G. (1991b). Development and survival of Mythimna convecta (Walker) and Persectania ewingii (Westwood) (Lepidoptera: Noctuidae) on cereal and grass hosts. Journal of the Australian Entomological Society 30: 295–302. McDonald, G. (1995). Insect migration in an arid continent. 1. The common armyworm Mythimna convecta in eastern Australia. In: Insect Migration: Tracking Resources Through Space and Time (Drake, V.A. and Gatehouse, A.G., eds) Cambridge University Press, New York. pp. 131–150. McDonald, G. and Smith, A.M. (1986). The incidence and distribution of the armyworms Mythimna convecta (Walker) and Persectania spp. (Lepidoptera: Noctuidae) and their parasitoids in major agricultural districts of Victoria, south-eastern Australia. Bulletin of Entomological Research 76: 199–210. McDonald, G, Moritz, K, Merton, E. and Hoffmann, A.A. (1995). The biology and behaviour of redlegged earth mite and blue oat mite on crop plants. Plant Protection Quarterly 10: 52–55.
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Narayan, D.S. (1962). Morphological, biological and ecological studies of the winter grain mite, Penthaleus major (Dugès), Penthaleidae: Acarina. Part 1. Journal of the Zoological Society of India 14 (1): 45–63. Robinson, M.T. and Hoffmann, A.A. (2001). The pest status and distribution of three cryptic blue oat mite species (Penthaleus spp.) and redlegged earth mite (Halotydeus destructor) in southeastern Australia. Experimental and Applied Acarology 9: 699–716. Smith, A.M. (1984). Larval instar determination and temperature-development studies of immature stages of the common armyworm, Mythimna convecta (Walker) (Lepidoptera: Noctuidae). Journal of the Australian Entomological Society 23: 91–97. TIMERITE® model for timing redlegged earthmite control: . Umina, P.A. and Hoffmann, A.A. (2004). Plant host associations of Penthaleus species and Halotydeus destructor (Acari: Penthaleidae) and implications for integrated pest management. Experimental and Applied Acarology 33: 1–20. Umina, P.A., Hoffmann, A.A. and Weeks, A.R. (2004). The biology, ecology and control of the Penthaleus species complex (Acari: Penthaleidae). Experimental and Applied Acarology 34: 211–237. Williamson, R.C. and Potter, D.A. (1997). Oviposition of black cutworm (Lepidoptera: Noctuidae) on creeping bentgrass putting greens and removal of eggs by mowing. Journal of Economic Entomology 9: 590–594 Woods, W., Lawrence, P.J. and Booth, P. (1980). Armyworm – a damaging pest of coarse grain on the south coast. Journal of Agriculture, Western Australia 21: 22–25. Woods, W. and Michael, P. (1987). Insect pests of broadacre farming. Western Australian Department of Agriculture, Bulletin 4113 2nd edition.
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3 COTTON PESTS L.J. Wilson, G.P. Fitt, S. Deutscher, M. Khan and B.A. Pyke
(a) Cotton seedlings (CSIRO)
(b) Cotton at squaring (production of flower buds) (CSIRO)
(c) Cotton plant (CSIRO)
(d) Cotton crop at boll burst (CSIRO)
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seedling squaring (flower buds) and boll setting boll maturation harvest
Jul Aug Sep Oct Nov Dec Jan Feb Mar Apr May
Phenology of a NSW–Qld summer cotton crop. The crop is chemically defoliated prior to harvest.
PEST (major pests in bold) Seedling stage field crickets silverleaf whitefly cotton aphid green peach aphid leafhoppers green mirid Rutherglen bug thrips wireworm false wireworms flea beetles cotton tipworm lightbrown apple moth cutworms lesser armyworm cluster caterpillar Squaring and boll-setting stages two-spotted mite silverleaf whitefly cotton aphid mealybugs leafhoppers apple dimpling bug green mirid cotton seed bug flower beetles redshouldered leaf beetle cotton leaf perforator cotton looper native budworm and cotton bollworm Boll maturation stage two-spotted mite locusts silverleaf whitefly
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PEST (major pests in bold) cotton aphid green mirid pale cotton stainer cotton harlequin bug green vegetable bug green stink bug redbanded shield bug pink bollworm and pinkspotted bollworm yellow peach moth rough bollworm cotton bollworm and native budworm
Cotton production in Australia Cotton is grown as a summer crop in NSW and Qld, which are the main producing states and where an average of 296 000 ha is under cotton, reaching 500 000 ha in good years, depending on cotton prices, rainfall and availability of irrigation water (Fig. 3.1). Australia produces high yields (most growers exceed 1800 kg lint per ha) and high quality fibre that is sought after in international markets. About 95% is exported. There is potential for cotton production in the Ord and Broome regions of WA, the Katherine region of the NT and the Burdekin region of Qld. In experimental production systems in the NT and WA, cotton is grown during the winter or ‘dry season’ to avoid the prolonged cloudy periods, heavy rainfall and higher insect abundance, especially of Helicoverpa, that occurs during summer or the ‘wet season’. Most (80–90%) of cotton in eastern Australia is irrigated, and the remainder is dryland cotton. The main rotation crops for irrigated cotton are winter cereals (wheat, barley) and vetch. Other common summer crops include sorghum and sunflowers. Sowing usually occurs in late spring (early October) with the onset of warmer weather. Squaring (flower bud production) begins after about 8 weeks and the first flowers appear in about 10 weeks. Most fruit (boll) is set and begins maturation about 3–4 months after planting, with boll burst (opening of cotton fruits to reveal the cotton lint with seed) usually beginning after about 5 months. Crops are chemically defoliated about 5 and a half months after planting and picking begins shortly after.
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Australian cotton is typically grown under a high input system: it is fertilised, irrigated and, until recently, arthropod pests have been controlled with pesticides, many of which are broad spectrum controlling not only the target pests but also reducing the abundance of natural enemies. Presently, about 85% of cotton is transgenic Bt (Bollgard II®), which produces the Cry1Ac and Cry 2Ab proteins that control Helicoverpa species and most other lepidopteran pests. Insecticide applications against noctuid caterpillars in these crops are reduced by 90% with higher survival of beneficials. However, insecticides formerly applied against Helicoverpa species masked a number of other pests, such as green mirids, cotton aphids and green vegetable bugs, which have now become more significant. A number of arthropods have developed resistance to pesticides: Helicoverpa armigera is widely resistant to endosulfan, pyrethroids, carbamates, and there is low level resistance to organophosphates, spinosad and indoxacarb; spider mites are resistant to organophosphates, bifenthrin and chlorfenapyr; aphids are resistant to the carbamate pirimicarb and to organophosphates, and silverleaf whitefly is resistant to some extent to almost all available insecticides. Resistance to insecticides is managed through the Insecticide Resistance Management Strategy, which is reviewed and updated annually. A comprehensive resistance management plan has also been developed for Bollgard II® cotton, and compliance with the plan is a condition for using transgenic cotton. Pest management, and increasingly agronomic management (irrigation, nutrition, growth), in eastern Australian cotton is largely done by 65
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Darwin Wyndam Derby
Northern Territory
Townsville
Queensland Emerald
Western Australia
Rockhampton
Theodore/Biloela South Australia
Perth
Darling Downs St George Brisbane Border Rivers New Bourke Gwydir South Namoi Wales Macquarie Tandou Newcastle
Lachlan Sydney
Adelaide
Canberra
Victoria
Melbourne
Existing cotton regions Tas. Hobart
Fig. 3.1. Cotton-producing areas of Australia.
cotton consultants, most of whom have tertiary training. The industry has a well-trained and coordinated extension network and strong links between industry, extension, research and funding agencies (e.g. the Cotton Research and Development Corporation) ensure regular publication of extension materials that are widely disseminated. The are also sophisticated decision support programs available for desktop and handheld computers which help with sampling, decision-making and data storage (e.g. CottonLOGIC). The Cotton Catchment Communities Co-operative Research Centre has also developed a Cotton Production Course, delivered primarily through the University of New England, which has been completed by many consultants and agronomists, and an integrated pest management (IPM) short course for cotton growers. A detailed IPM strategy has been developed for Australian cotton (Deutscher et al. 2005). Information on the effect of insecticides and miticides on predators and parasites of pests can also be found in these guidelines.
S E E D L I N G S TAG E
Field crickets N (Main entry in Chapter 15 Pastures—winter rainfall)
Pest status on cotton: Minor, widespread, irregular. Field crickets, Teleogryllus spp. may damage cotton occasionally when present in plague numbers. The adults and late-stage nymphs feed on the leaves and stems of seedlings and may reduce a stand to the extent that replanting is necessary.
Silverleaf whitefly Bemisia tabacii (Gennadius) Hemiptera: Aleyrodidae Distribution: The silverleaf whitefly (SLW) is a pest of crops in the USA, Europe, Asia, Africa and Australia. It has been recorded in cotton crops from all growing areas in eastern Australia and from Katherine in the NT, but not yet from
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the Ord River Irrigation Area. However, it has only reached levels regularly requiring the use of insecticides in Central Qld though it has occasionally required control in the Darling Downs, Byee and St. George areas (Kelly et al. 2002a). Pest status: Minor south of 25°S latitude, major north of about 25°S latitude, widespread in eastern Australia, irregular. Identification: SLW adults are small white insects about 2 mm long (Fig. 3.2a). SLW eggs are tiny and are attached to the undersides of leaves by a short stalk (pedicel). The nymphs are small (< 1 mm) discs attached to the lower leaf surface. May be confused with: Three main types of whitefly are found on cotton in Australia: the greenhouse whitefly Trialeurodes vaporariorum (Westwood) (Fig. 3.2b); and two biotypes of Bemisia tabaci, the B. tabaci B-Biotype, known as the silverleaf whitefly (SLW, Fig. 3.2a), and the eastern Australian native (EAN) biotype. Greenhouse whitefly and EAN are the most commonly found, particularly in cotton fields adjacent to sunflowers but are not significant pests of cotton. Greenhouse whitefly are about twice the size of B. tabaci. The main distinguishing feature is the way they hold their wings. B. tabaci has a split between the wings, whereas the Greenhouse whitefly has a
b
overlapping wings that form a heart shape. It is not possible to separate the two B. tabaci biotypes in the field. A biochemical test such as an esterase or the polymerase chain reaction test is needed to identify the biotype. SLW poses a greater pest threat than other whiteflies because of its greater host range, quicker reproductive rate, and its ability to rapidly develop resistance to insecticides. Under suitable conditions, SLW can outbreak on an area-wide scale to a point where management with insecticides is difficult. Host range: SLW is found on a wide range of hosts, including ornamental plants in nurseries, herbs such as basil, and on weeds and crops (sunflower, soybean, melons) in the field. Sow thistle is a preferred host. Life cycle on cotton: SLW eggs are generally laid haphazardly on the underside of leaves. The newly laid eggs are yellow–green, changing colour to dark tan as they are about to hatch. They are very small (difficult to see with the naked eye), oval-shaped, and sit on top of a pedicel (stalk) that fits into a small slit in the leaf made by the female. The pedicels enable moisture to be drawn from the leaf, preventing dehydration of the eggs. Leaves with high densities of whitefly eggs often have a dark sheen on the underside. The first instar (crawler stage) is the only nymph stage that is mobile. They travel a few centimetres to find a suitable part of the leaf to start feeding by inserting their mouthparts into the leaf, extracting plant sap. Instars 2 through to early 4 continue feeding without moving from the position selected by the crawler. During the late fourth instar, the insect stops feeding and becomes a pupa (or ‘red eye’), out of which emerges an adult. Female whitefly are produced from fertilised eggs and males from unfertilised eggs. Most of the adults emerge as females and, as females tend to live longer, they make up a significantly higher portion of the population. Each female can lay up to 300 eggs. The life cycle from egg to adult can be as short as 18 days in the summer; this is slowed in cooler weather. Risk conditions: Outbreaks are favoured by warm weather and host availability (De Barro 1995).
Fig. 3.2. (a) Four adult silverleaf whitefly (2 mm long) showing the characteristic narrow body shape and split between the wings. Nearby are translucent pupal cases from which they have emerged (CSIRO: C. Mares) . (b) Adult greenhouse whitefly (3.5 mm long) (SARDI: G.J. Baker) .
Climate: SLW does not have an overwintering diapause stage so its distribution is limited to those areas where it can survive winter conditions. In areas with cold winters, populations can persist in glasshouses. 67
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The growth rate of SLW decreases and generation time increases in cooler weather. It is a potential pest in areas where the winter generation time is 80 days or less (as occurs in areas of Qld from Biloela north). Continuity of suitable hosts: A long period of continuous availability of suitable hosts is essential for whitefly outbreaks. The discontinuity of host availability in some Australian cottonproducing areas such as northern NSW reduces the likelihood of outbreaks. In contrast, the almost continuous availability of suitable hosts in the Emerald Irrigation Area (Qld) increases the probability of further outbreaks. Here, whitefly may reach damaging levels late in the cotton season (Kelly et al. 2002a) Damage: Large numbers of SLW can retard growth by removing assimilates, creating damage similar to aphid feeding. SLW also secretes large quantities of sticky honeydew that reduces photosynthesis and tarnishes cotton fibre. Whitefly honeydew is more difficult to remove in processing than aphid honeydew because it has a higher boiling point. SLW whitefly is also a highly efficient carrier of viruses. SLW is a vector of gemini-viruses such as cotton leaf curl in Pakistan and the USA. None of these viruses are known in Australia, but should they enter the country the SLW is now present to spread them (Kelly et al. 2002b). Monitoring: Proper identification and regular estimations of the numbers of adults and nymphs are important for monitoring. Standard sampling techniques have been adapted to Australia (Sequeira et al. 2006) from Arizona (Ellsworth et al. 1995).
Species identification Samples of whitefly need to be tested biochemically to separate the EAN and SLW biotypes. Collect 200 leaves from the field, selected at random from the fifth to eighth mainstem leaf below the terminal. Pack the leaves into a paper bag, seal, then place this inside a plastic bag. Put the sample into a small insulated box with a frozen icebrick and send by express courtier to: NSW DPI, Tamworth Agricultural Institute, Calala Lane, Tamworth NSW 2340, marked ‘Urgent Whitefly Samples’. Include a note with your name and contact information, as well
as information on the farm and field from which the sample was taken. Population monitoring: Once the whiteflies have been identified: 1
Designate a management unit of 20–35 ha. Each management unit should have a minimum of two sampling sites. Sample 15 leaves per site (30 leaves per management unit). Sample at least weekly when SLW is confirmed.
2
Choose a plant to sample. Move at least 10 m into the field before sampling. Avoid sampling plants disturbed by other pestsampling. Choose only healthy plants at random. Sample along a diagonal or zigzag line moving over several rows. Take five to 10 steps before selecting a new plant.
3
Choose a leaf. Sample the fifth mainstem node leaf from the top of the plant (the first node is that with an unfolded leaf)
To estimate numbers of adult SLW, a two level or binomial scoring system is used. Keep shadow off the plant and carefully turn the leaf over by the tip of the leaf blade or the petiole. Tally the leaf as ‘infested’ if the whole leaf has three or more whitefly adults. Tally the leaf as ‘uninfested’ if it contains less than three whitefly adults. Calculate the % of infested leaves: % infested = (infested leaves/total leaves sampled) × 100 Then calculate the number of SLW adults per leaf using the following table: Average number adults/leaves* 1 2 3 4 5 6 7 8 9 10
14% 28% 39% 49% 57% 64% 70% 75% 79% 82%
*An infested leaf has three or more adults.
Sampling for SLW nymphs can either be done with % presence or absence of leaf discs with SLW, or
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by using actual numbers (nymphs per leaf disc). Binomial sampling often gives more consistency between observers. Sample nymphs within a 3.88 cm2 (10 cent piece-sized) disc. Place the disc between the main veins on the left side. Count large (third and fourth instars) nymphs in area. Large nymphs are those that are visible to the naked eye. These third- and fourth-instar nymphs appear as flattened, oval discs or ‘scales’. Use the disc to record presence or absence of SLW on each of 30 leaves. Discs with one or more large nymphs are scored as positive.
leaves, or 1.6–2.3 adults/leaf. Average nymphs/ disc of greater than 0.25 at this time would support the decision to use the IGR. The decision to use the IGR should not be delayed beyond either 1600 DD or greater than 50% infested leaves or 2.3 adults/leaf as yield can be lost, the efficacy of the IGR may be reduced and substantial lint contamination can occur before the product has time to control the population.
Calculate the % of positive leaves:
Application of the IGR earlier than 1500 DD increases the chance that the population will recover towards the end of the season and expose the crop to lint contamination from honey dew. Between 1300 and 1500 DD, if the population has infested no more than 10–20% of leaves (or is >1 adult/leaf), the opportunity for whitefly suppression should be considered when making control decisions for other pests. Endosulfan and diafenthiuron are registered for a range of key pests, and when applied at this time will also suppress SLW populations. This suppression may be sufficient to prevent the population reaching the IGR threshold, but is not a substitute if high populations are present during this period. Similarly, if SLW populations are below the IGR threshold at 1600 DD, but 10–20% of leaves are infested before 1800 DD is reached, choosing to use diafenthiuron for aphid control at this stage would also reduce the whitefly population to below damaging levels. While late populations are less likely to reduce yield, the moderate–low density of 10–20% infested leaves at this time is sufficient to contaminate lint with honey dew.
% infested leaves = (infested discs/total discs) × 100 OR Calculate the average number of large nymphs per disc: average number of large nymphs per disc = total nymphs/total discs sampled Management decisions should be based mainly on adult numbers (% infested leaves or mean adults/leaf) (Sequeira et al. 2006). The disc counts of large nymphs provide supporting information. The management options available in-crop and the thresholds that trigger their use are dependent on the crop’s stage of development. The Decision Support Matrix should be used between peak flower and first open boll to identify the threshold that applies to the crop.
Using the IGR Insect growth regulators (IGRs) are the cornerstone of effective SLW management in cotton. They provide excellent control of SLW across a broad range of population densities. They are very selective, allowing survival of whitefly predators and parasitoids. However there is a very high risk that resistance will develop and lead to control failures in the field. ENSURE ONLY A SINGLE APPLICATION OF AN IGR OCCURS WITHIN A SEASON. Twice weekly monitoring from peak flower will ensure that if thresholds are reached, the IGR can be applied at the time when it will be most effective. The optimum time to use the IGR is when the crop is between 1500 and 1600 DD and population monitoring shows 37–50% infested
Early season suppression and late season knockdown
Chemical control: IGRs selectively kill SLW in relation to beneficial populations. Determine if the whitefly is SLW, EAN or greenhouse whitefly. Use IGRs first when numbers exceed threshold. Effects of these products will not be seen over a week after application. Delay use of other control options until after IGR application. Later in the season influxes of whitefly may require control with other compounds such as pyrethroids or organophosphates. Adhere to Insecticide Resistance Management Strategy (IRMS). Action thresholds and control recommendations are updated periodically; check the Cotton pest management guide (NSW DPI) for details.
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SLW control decision support matrix Crop growth stage Cumulative day degrees Suppression Mean adults/leaf1 % infested leaves Control with IGR Mean adults/leaf1 % infested leaves Nymphs/disc Knockdown Mean adults/leaf1 % infested leaves 1 Expected
Peak flower 1300 1350 1400 > 0.5 > 10
SLW thresholds relative to crop development First open boll Open cotton 1500 1550 1600 1650 1700 1750 1450
1800
< 1.0 < 25
1.0 25
1.2
1.3 30
1.
1.6 37
1.8
2.3 50 >0.25
2.4 55
2.8 60
3.3 70
4.2 80
0.2
0.2
0.2
0.3
0.3 5
0.4
0.5 10
0.6
0.7 15
0.8
0.9 20
population density
Notes Sampling protocol
Day degrees
Suppression early season
Control with IGR
Knockdown late season
Sample 30 leaves at 5th node below the terminal/25 ha weekly from first flower (777 DD) and twice weekly from peak flowering (1300 DD). Use % infested leaves (recommended) or mean adults/leaf. Infested leaves are those leaves with 2 or more adults. Disc counts of large nymphs provide supporting information. Day degrees are calculated using the formula: Daily day degrees (DD) = [(Max °C – 12) + (Min °C – 12)] ÷ 2 Access the cumulative day degree information for your nearest SILO weather station from the Cotton CRC website – www.cotton.crc.org.au Long-term averages suggest the periods from 1300–1500 DD and 1500–1600 DD are: • 12 days and 9 days at Emerald, • 15 days and 9 days at Dalby, and • 13 days and 7 days at St George. Twice weekly monitoring is critical to identify trends in population development during these periods when suppression and control strategies can be most effective. Endosulfan and diafenthiuron are products that may be used to control aphids and some other pests. When used for these purposes, these products can slow or suppress the development of SLW populations if they are present at 10–20% infested leaves (< 1 adult/leaf). Population suppression will not occur if there is > 25% infested leaves or crop development is past first open boll. The window for endosulfan application by ground rig closes on 15 January 2007. See label directions and IRMS. ENSURE ONLY A SINGLE APPLICATION OF AN IGR OCCURS WITHIN A SEASON. Check with your local Department of Primary Industries officer or the APVMA for information on registration/permit status for IGRs to control SLW in cotton. Optimum efficacy will be achieved with IGRs if the application is made when the SLW population is consistently equal to or above the corresponding threshold values, and crop development is between 1500 and 1600 DD. Early application (before 1500 DD) may see populations rebuild before the end of the season. Delaying IGR use beyond 50% infested leaves or 1600 DD can result in significant lint contamination and potential yield loss. Diafenthiuron may be used to prevent lint contamination by late season aphids. When used for this purpose, diafenthiuron can slow or suppress the development of SLW populations if they are present at 10–20% infested leaves (< 1 adult/leaf) after 1600 DD. Population suppression to below threshold levels may not occur if there is > 25% infested leaves.
Cultural control: Crop residues from all susceptible crops should be rapidly destroyed after harvest. The management of farm weeds that can act as a host is particularly critical over the winter period. Host species include: bladder ketmia, native rosella, Rhynchosia spp., morning glory vines (cow, bell, potato), sow thistle, rattlepods, native jute, burr gherkin and other cucurbit weeds, Josephine burr, sunflowers
when young, euphorbia weeds, poinsettia, and volunteer cotton. Where possible, allow a hostfree period for the pest over winter. Plant cotton early in the season to avoid latematuring crops, which act as a bridge to the next host. Moisture stress, and excessive amounts of nitrogen are associated with increased SLW numbers and late-maturation of the crop.
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Table 3.1. Deciding whether to spray insect growth regulator (IGR) against SLW on cotton (modified from Management in Arizona Cotton 1996), using two alternative sampling methods: presence/absence OR actual number of SLW per leaf disc Whitefly adult levels
Silverleaf whitefly large nymphs
Presence or absence of large nymphs
Actual numbers of nymphs per leaf disc
Less than 27% infested
Less than 0.5
At least 27–40% infested
At least 0.5–1.0
Host-plant resistance: Plant smooth-leafed varieties as hairy leaf cotton varieties are more susceptible to SLW than smooth-leaf varieties. Most Siokra varieties (okra leaf shape) are less susceptible than normal leaf. Conservation of natural enemies: A complex of predators and at least 14 species of parasitoids (sometimes giving high parasitism rates) have been found in Australia. Parasitoids of whitefly include several wasps in the genera Encarsia (Fig. 5.28) and Eretmocerus (Aphelinidae). Predators of whitefly nymphs include big-eyed bugs (Fig. 3.35), lacewing larvae (Fig 3.41) and ladybirds. A management strategy needs to preserve and promote the activity of predators and parasites, which includes the avoidance of early-season use of broad-spectrum insecticides, particularly pyrethroids and organophosphates.
Fewer than 3/leaf Wait and resample in 3–7 days Resample in 3 days OR apply an IGR
At least 3–5/leaf Resample in 3 days OR apply an IGR Apply an IGR
feeding on a healthy cotton crop, they will be larger and darker in colour, although this will range from dull pale green, dark green, brown and black (Fig. 3.3). Aphids that are parasitised by small wasps become brown and distended (Fig. 3.3). May be confused with: Two other aphid species may feed on cotton but rarely develop to damaging levels: green peach aphid (Myzus persicae (Sulzer)) and cowpea aphid (Aphis craccivora Koch). The cowpea aphid is similar to the cotton aphid in appearance but the wingless adults of this species are a shiny black, in contrast to the cotton aphid, which is a dull colour. The cowpea aphid colonises a range of hosts but prefers legumes and is often found on medics. This species is often found on cotton early in the season but seldom establishes.
Cotton aphid Aphis gossypii Glover Hemiptera: Aphididae Distribution: Cosmopolitan, widely distributed throughout the world on agricultural, horticultural and glasshouse crops. Widely distributed in Australia, found in all states. Pest status: Major, widespread, irregular. Identification: Cotton aphids are small insects about 1–2 mm in length. They have a rounded body shape with long antenna and at the tail end two distinctive siphunculi resembling dual exhaust pipes. Siphunculi produce defence chemicals and alarm pheromones. At the very tip of the tail end is the cauda, used to flick away honeydew, a sticky fluid that the aphids excrete. Cotton aphids vary widely in colour. If raised over several generations on poor food they will be small and yellow, known as yellow dwarfs. If
Fig. 3.3. Colony of cotton aphid showing wide variation in size and colour. Two large pale brown parasitised aphids (‘mummies’) are at the top. (CSIRO: P. Reid)
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The green peach aphid is a pale yellow–green and is more teardrop-shaped than the cotton aphid. Colonies tend to be uniform in colour compared with cotton aphid. Seen with a hand lens or microscope, green peach aphid has a small tubercle at the junction of the antenna and head, which is absent in cotton aphid and cowpea aphid. The area between these tubercles is Ushaped in green peach aphid, whereas it is flat in cotton aphid. Also, the green peach aphid has a pair of long, pale, tube-like siphunculi at the tip of its abdomen, whereas those of cotton aphid are quite short and usually dark. The green peach aphid prefers cooler conditions. It is sometimes found on cotton early in the season but populations do not usually persist in hot weather. Other aphid species occasionally found on young cotton are mostly the winged forms of species that have migrated from other hosts, especially leguminous weeds. They test feed but do not normally establish. These include the pea aphid (Acyrthosiphon pisum (Harris)), the blue green aphid (Acyrthosiphon kondoi Shinji), the spotted alfalfa aphid Therioaphis trifolii f. maculata (Monell), the corn aphid Rhopalosiphum maidis (Fitch) and the oat or wheat aphid Rhopalosiphum padi (Linnaeus). Host range: The cotton aphid has a broad host range and has been recorded on members of the following families: Fabaceae (legumes, lucerne), Solanaceae including thornapples, ground cherries, nightshades, Cucurbitaceae including paddymelons, Malvaceae including bladder ketmia, marshmallow, paddy’s lucerne, and Asteracae including sunflower, capeweed, daisies, thistles, and Bathurst burr. Life cycle on cotton: Cotton aphids have winged (alate) and non-winged forms (aptera). The winged forms are adult females and migratory. They are produced when aphid colonies are stressed due to overcrowding or poor food quality or can be induced by reduced day length. Alate aphids settle on plants and test feed. If the plant is unsuitable, the winged adult will resume the flight, settle, and test feed sequence until it finds suitable food or dies. If the plant is suitable, production of live young commences quickly. These mature through four nymphal stages into wingless adults. Female aphids give birth to live female young which are clones of
themselves, inheriting all of their characteristics including insecticide resistance. Within a cotton region there may be different aphid population clones. A female aphid can produce live young at the rate of four to six per day, which in summer can mature into adults in 4–7 days and immediately begin producing live young, leading to many generations per year. Aphids are all female and have within them several live young at various stages of development. These cloned offspring already have clones developing within them before they are born allowing production of many generations in a short period. Populations can explode if conditions such as food quality and climate are suitable. The wingless cycle continues until some aspect of their environment triggers the switch to the production of dispersing (winged) forms. When this occurs, adult female aphids then produce live young, which develop wing-buds. These nymphs mature into winged adults, which then fly off to find a new host plant. In winter, cooler temperatures slow the growth rate of aphids dramatically. In Australian cotton regions, cotton aphids persist through winter at low levels of abundance on available host plants. In spring, as temperatures increase, aphid populations begin to build rapidly again. Risk period: Throughout crop life. The cotton aphid is favoured by warm temperature and does well on cotton through the peak growing period. Damage: Damage symptoms from cotton aphids initially appear as crinkled and curled leaves, with the margins of the leaves curling downwards. Aphids cause damage to cotton in four ways: 1
Aphids compete with young growth and developing fruit (squares and bolls) for assimilate. If this is beyond the capacity of the plant to compensate for, then some reduction in growth is likely.
2
The presence of aphids on leaves reduces photosynthesis. The cause of this effect is not well understood but could be due to a number of factors including the damage caused by insertion of stylets, especially when there are many aphids, the effects on assimilation depletion, or the effects of saliva secreted into the plants by the aphids.
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3
4
Secretion of honeydew. As aphids feed on the phloem vessels they imbibe fluid rich in sugars which is excreted as honeydew. Honeydew on leaves can reduce photosynthesis (Fig. 3.4) and contaminates lint, making it sticky and discoloured. Severe downgrading of sticky lint may result because of the difficulties of processing it in high speed spinning machinery. Spread of the disease cotton bunchy top (CBT), which reduces yield (Fig. 3.4). This disease results in shortening of internodes and petioles, reduced leaf size, boll size and yield and is characterised by the presence of angular mottling on leaves.
Pre-squaring cotton appears to be able to fully compensate for aphid damage if aphid feeding ceases. However, prolonged high densities up to cut-out (when fruit production slows or stops) can a
b
Fig. 3.4. Honey dew contamination of leaves: (a) uncontaminated and contaminated (CSIRO: L. Wilson) , and (b) leaf showing angular mottling symptoms characteristic of cotton bunchy top (CSIRO: A. Reddall) .
cause reduction of yield. Populations in excess of 90% of plants infested with aphids for 3 or more weeks are likely to result in economic loss. Monitoring: Aphid sampling should begin from seedling emergence and continue at least weekly. Aphids generally prefer younger growth, so this is a good place for visual monitoring. Score a plant as infested only if non-winged forms are present (winged forms could be another similarlooking species that is just test feeding before moving on to another host). If a high proportion of plants do have winged forms then resample within a few days to check if they have settled and produced young. If reproducing aphids are found it is important to confirm the species, as green peach aphid causes more severe damage than cotton aphid and has a different insecticide resistance spectrum. Action level: Cotton may recover from aphid damage even if high numbers (> 90%) of plants are infested, provided the infestation doesn’t persist for too long (< 10 days). Control should be applied if the standard threshold of 90% of plants infested occurs during the growing season, or slightly earlier if it is clear that aphid populations will soon exceed threshold. Once cotton has open bolls, the threshold should be reduced to 10% of plants infested if honeydew is present on leaves or open bolls. The recent advent of the CBT disease, for which cotton aphid is a vector, means that application of a standard threshold is more difficult. Not all aphid populations carry the disease. Early infestations of cotton aphid should be monitored for the presence of CBT symptoms on plants. It may be possible to spot spray these areas if CBT symptoms are found. It is important not to exacerbate the spread of aphids by reducing their predators, which may occur if broadspectrum insecticides are used to which the aphids are resistant. Chemical control: A limited range of insecticides may be applied as foliar sprays. Insecticides may also be applied as seed treatments, or can be applied as granules with the seed at planting, and can provide control of aphids on seedling cotton for 3–6 weeks after emergence and may help reduce the risk of infection of the crop with CBT. Care must be taken in selection of insecticides as many populations of cotton aphid
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are resistant to particular insecticides, including some organophosphates and carbamates. There is also cross-resistance in aphids between some organophosphates and some carbamate insecticides. Care must also be taken in the choice of seed treatment or at planting insecticides. These also select for resistance in aphid populations (Wilson and Spora 2001; Wilson et al. 2001). Cultural control: Where resistance aphids are detected in previous seasons, a non-host rotation crop, such as a winter cereal, together with the on-farm control of weeds, cotton stubble and cotton volunteers, may reduce reservoirs of aphid resistance and disease transmission. Host-plant resistance: None of the current commercial cotton varieties show strong resistance to aphids. There are several CBTresistant varieties available. These varieties are still susceptible to aphids and aphid damage but do not develop CBT symptoms. Conservation of natural enemies: A range of parasitoids and predators will help to reduce aphid survival. Predators of aphids include: ladybird larvae, damsel bugs, big-eyed bugs, and the larvae of green lacewings and hoverflies. Wasp parasitoids (Aphidius colemani Viereck, Braconidae and Lysiphlebus testaceipes Cresson) mummify and kill aphids (mummified aphids appear as bloated pale brown aphids that do not move (Fig 3.3)). A range of coccinelids, the red and blue beetle, predatory Hemiptera and Diptera also attack aphids (Pyke and Brown 1996). When conditions are only moderately favourable to aphids, natural enemies may successfully limit populations. However, under conditions favourable for mass dispersal and rapid aphid increase, natural enemies may not effectively prevent economic damage. Some insecticides (directly, or by drift) increase aphid numbers because of their differential mortality against beneficials (see www.cotton.crc.org.au and Wilson et al. 1999).
A strategy for managing resistance in cotton aphids (Johnson and Farrell 2004): 1 A maximum of two sprays of any registered aphicide against aphids, unless the product is otherwise restricted. 2 Rotation of chemistry; that is, do not use the same chemical group consecutively. 3 The first aphicide spray should not be from the same chemical group as any seed treatment or at-planting insecticide used that also controls aphids. 4 There is cross-resistance between some carbamates and some organophosphates and these specific insecticides are considered as the same group for aphid control.
The green peach aphid (Fig. 5.17) is often confused with the cotton aphid. The life cycle, economic damage and thresholds for green peach aphid in cotton are similar to that of cotton aphid. It has a wide host range and is often found on the plant families Fabaceae (legumes, lucerne and lupins), Asteraceae, Brassica sp., and peach and plum trees, and also a number of weed hosts common in cotton or nearby fields. These include cow vine, turnip weed, marshmallow and thornapples. Green peach aphid is sometimes found on young cotton, but it does not tolerate hot conditions so rarely becomes a problem. Regular monitoring will help determine the need for control. The green peach aphid causes more severe stunting of cotton plants than does cotton aphid. The threshold for control is correspondingly lower, at 25% of plants infested.
Leafhoppers Hemiptera: Cicadellidae Vegetable leafhopper, Austroasca viridigrisea (Paoli)
Green peach aphid
Cotton leafhopper, Amrasca terraereginae (Paoli)
Myzus persicae (Sulzer) Hemiptera: Aphididae
Pest status on cotton: Minor, widespread, irregular.
Pest status on cotton: Minor, widespread, irregular.
Identification: Leafhoppers, or jassids as they are commonly known in the Australian cotton
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industry, are small, leaf-feeding insects ranging in colour from green, through yellow–green to brown. The vegetable leafhopper is the most common leafhopper on cotton in NSW (Fig. 3.5a), while the cotton leafhopper also occurs in NSW but is more abundant in Qld (Fig. 3.5b). Other species such as the common brown leafhopper, Orosius argentus Evans and the spotted leafhopper, Austroagilla torrida Evans are also occasionally found. Adults are small, elongate, wedge-shaped insects about 3 mm long, quick to hop and fly off when disturbed.
Action threshold: The current threshold for control through the early season is 50 leafhoppers per metre of row.
Life cycle on cotton: Eggs are laid in slits made in soft plant tissue. Nymphs resemble adults but are smaller, paler and wingless and generally slower moving.
Distribution: The green mirid is a native species widely distributed across Australia. It is a pest in both eastern and northern Australian cotton regions.
Risk period: Leafhoppers occasionally damage seedlings and new growth, but they are more likely to attack mature leaves from the flowering stage onwards.
Pest status: Major, widespread, irregular.
Damage: Leafhoppers feed on the upper surface of cotton leaves, their stylets penetrating to about half of the thickness of the leaf, and their damage appears as a pale stippling effect. In unsprayed cotton or cotton sprayed infrequently and with selective insecticides, leafhoppers can build to large numbers in the mid to late season, and the cumulative feeding damage can make leaves appear almost white. Photosynthesis in damaged leaves can be reduced. The photosynthetic rate of young leaves (node 3–4 below the terminal) with 80% of the upper surface damaged is reduced by 20% compared with undamaged leaves, but the impact on yield has yet to be demonstrated.
Control: Broad-spectrum insecticides can be used, but at the risk of reducing beneficial insect populations and creating outbreaks of other pests.
Green mirid Creontiades dilutus (Stål) Hemiptera: Miridae
Identification: Green mirid adults are about 7 mm long, pale green with long antennae that have a red tip (Fig. 3.6b). The nymphs are similarly pale green with long antennae, also usually with a red tip. First and second stage nymphs are about 1.5–3.0 mm long and wingless (Fig. 3.6a). Wing pads develop at third instar. Adults and nymphs move quickly away from a
a b
b
Fig. 3.5. (a) Vegetable leafhopper adult (4 mm) (CSIRO: , and (b) cotton leafhopper adult (3 mm) (CSIRO: C. Mares) .
P. Room)
Fig. 3.6. (a) Green mirid nymph (3 mm) (CSIRO: C. Mares) , and (b) adult green mirid (7 mm) (K. Power) .
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disturbance, which makes them difficult to locate in the field. May be confused with: A related species, the brown mirid (Creontiades pacificus), is also sometimes found in cotton, but is generally less abundant than the green mirid. Brown mirids are quite common in pulse crops (Fig. 7.17). Unlike green mirid, the triangular pronotum region in the brown mirid is darker and the antennae are striped with red and white. The brokenbacked bug (Taylorilygus pallidulus (Blanchard)) (Fig. 7.15) is similar in appearance to the green mirid but is smaller and is not often found in large numbers in cotton. Host range: Mirids feed and develop on a wide range of host plants, including sunflower, safflower, lucerne and many legume, and weed species including wild turnips, verbena, common joyweed and many species of thistles (Asteraceae). Life cycle on cotton: The green mirid passes through an egg and five nymphal stages to become an adult. Within a crop, mirids lay eggs singly, preferring the leaf petiole. The egg is inserted into the plant tissue with an oval egg cap projecting above the leaf or petiole surface. Eggs hatch after 5–7 days depending on temperature; at 30–32°C eggs hatch within 4–5 days and each of the five nymphal stages takes about 2–3 days to develop. The pale green nymphs range in size from 1.5 mm to 6.7 mm and all instars have long antennae (2–9 mm) and are highly mobile. In summer, a generation (egg to adult) can be completed in about 3 weeks. Adults can live for 3–4 weeks. Mirid populations may vary significantly with climatic conditions. In sustained hot weather (three consecutive days > 35°C), numbers may decrease. Numbers also tend to be lower immediately after heavy rains or storms, though early in the season it is thought that storm fronts may also bring influxes of adults. These factors need to be considered when sampling (Khan et al. 2004). During the winter months, mirids are often difficult to locate, overwintering in low numbers as adults or eggs on wild plants. However, as temperatures rise in August, their populations increase. Mirids move into cotton crops during November as alternative host plants near cotton crops dry off and they seek a fresh food source. There is also evidence of long-distance
migration, possibly from inland areas, associated with weather fronts, and may be the cause of influxes of mirids sometimes observed in cotton-growing regions early in the season. Risk period: Likely to be a pest all seasons. Damage: During feeding, green mirids pierce the plant tissues with their sharp mouthparts or stylet and release a chemical (pectinase) that destroys cells in the feeding zone. The affected tissue rapidly dulls in colour, then blackens, desiccates and dies. Favoured feeding sites include plant terminals, young leaves (particularly on seedlings), small squares and young bolls. It is common for these structures, especially terminals and squares, to be destroyed by mirid feeding. Green mirid feeding on small squares (5–7 cm long) and small bolls (< 10 days old) causes shedding. If the bolls are larger, individual locks may be damaged. During the boll-filling stage, green mirid damage is similar to that of green vegetable bug and is characterised by the development of shiny black spots on the boll, the presence of warty growths inside the boll wall and brown staining of the developing lint. After approximately 20 days bolls are protected by a thick boll coat and do not suffer significant damage. If seedling tip damage and early square loss caused by green mirids is excessive, maturity can be delayed and yield reduced. Mirid damage is cumulative and maximum damage occurs when the insect reaches the fourth and fifth nymphal stages. Adult green mirids cause more damage than the nymphal instars 1–3 and similar levels of damage to the final instars 4 and 5. Monitoring: Mirids are very mobile pests and populations can fluctuate rapidly, so sampling needs to be done two to three times per week. Both adult and immature mirids should be counted using beat sheets, which are quick to use and give consistent results. When plants have less than nine nodes, beat sheets and visual checks give approximately the same estimates of mirid numbers. However, as plants grow beyond nine nodes, the beat sheets become more effective for sampling mirids and the ratio of mirids in visual counts to beat sheet counts is about 1:3. If using beat sheets, the numbers should be divided by 3 as the thresholds are based on visual sampling. An
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alternative method used to estimate mirid densities is sweep-netting. This technique has proven to be the best method for finding mirid adults, while the beat sheet is the best for finding nymphs. A standard sweep net sample is 20 vigorous sweeps through the crop canopy. Counts from a sample of 20 sweeps can be converted to visual count equivalents by dividing the adult count by 3 and the nymphal count by 1.6. Sampling should occur two to three times per week, becoming more frequent when approaching a control decision. It is also important to monitor plant damage, as mirids can be present yet there may be little damage. Monitoring for fruit retention should be done weekly or before an insect control decision. If retention falls below 50–60%, calculation of the fruiting factor should indicate whether yield loss is likely. A description of fruit retention monitoring and the use of the fruiting factor can be found in the IPM Guidelines and in the Cotton Pest Management Guide, both available from the Cotton Technology Resource Centre at Narrabri and from the website www.cotton.crc. org.au. Action level: Thresholds for mirids vary between warm and cool season areas due to the cotton plant’s ability to compensate for damage. Thresholds early in the season are shown in Table 3.2. Mid-season thresholds are still being developed, but early-season thresholds can indicate the need for more intense searching for mirid damage: (i) evidence of mirid numbers using Table 3.2, and (ii) evidence of damage exceeding guideline levels (below). Control decisions for mirids should use a combination of plant damage and mirid numbers. For example, if early-season fruit retention is very low (< 40%) then mirid thresholds may need to be lowered to prevent further loss. Alternatively, in crops with very high retention (> 80%), thresholds may be lifted. Thresholds for northern Australia, where cotton is grown through the winter, are similar to the cool regions in Table 3.2. Table 3.2. Early season mirid thresholds (numbers of adults and nymphs/m in visual checks)
Visual Beat sheet Sweep net: adults Sweep net: nymphs
Warm area
Cool area
1.0 3.0 3.0 1.6
0.5 1.5 1.5 0.8
Tip damage: Cotton plants have a very good ability to recover from early season terminal damage. Terminal damage may be less prevalent in Bollgard II® due to control of Helicoverpa spp. and tipworm. For light terminal damage, where only the tip is damaged, a threshold of 100% of plants each damaged once could be used with little risk of yield loss or maturity delay. For heavier terminal damage, where is the tip and one or two true leaves are destroyed, the threshold should be reduced to 50% of plants damaged once. Fruit retention: Crop yield and maturity are not affected if first position fruit (flower buds) retention is maintained around 60% at the time of flowering. This is particularly relevant in Bollgard II® cotton as very high retention may reduce growth and limit yield. Procedures for checking retention are provided in Deutscher et al. (2004). Boll damage (flowering–defoliation): Feeding damage from mirids results in shiny black spots on the outside of the bolls at the feeding sites, which are easily mistaken for normal blemishes. Boll damage is monitored by cutting open 100 bolls in the 12–15-day-old range and inspecting for internal warty growth or stained lint. External spots are not always indicative of feeding. A provisional threshold of 20% of damaged bolls is suggested. Procedures for checking boll damage are provided in Deutscher et al. (2005). Chemical control: Some insecticides applied at planting can control mirids during early crop growth but none provide control into the fruitsetting period. Those used at planting may also suppress and select for resistance in other pests, especially aphids. Care is needed in selecting the first insecticides to be used to control aphids, mites or mirids in the crop. These insecticides should be from a different mode of action to that of any insecticides used at planting (e.g. seed treatments, granular insecticides, soil sprays) so that there is not repeated use of the same chemical group. This will help reduce the selection for insecticide resistance in these pests. Most insecticides registered for control of mirids and stink bugs have moderate or high impact on beneficial insect populations and their use may contribute to flaring some pests late in the season, especially when multiple applications are made. 77
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High mirid numbers may be controlled by two sprays about 7 days apart, using a low insecticide rate with an additive (see below), which will have short residual activity but also less impact on beneficials than full label rate. The second application to control young nymphs emerging from eggs is only necessary if subsequent sampling detects enough mirids to justify a spray. Alternatively, when adults start to be found in the crop it is prudent to assume they will lay eggs. If detected early it may be possible to control the adults with a reduced-rate insecticide before there is significant egg-lay. Cultural control: Cotton is not the most preferred host for mirids. Planting crops that are more attractive to mirids close to cotton crops may reduce populations in cotton. When maintained well, lucerne strips within a field or on borders or as larger blocks on the farm can reduce mirid populations within cotton crops. See Deutscher et al. (2005) for more details. Host-plant resistance: None of the commercial cotton varieties available have useful resistance to mirids.
Pest status on cotton: Minor, widespread, irregular. Adult bugs (Fig. 5.30) are often found on cotton but they do not tend to feed and are unable to reproduce on the crop. Risk period: Seedling stage. Damage: Starving nymphs occasionally damage border cotton adjacent to drying sunflower crops. If cotton crops follow sunflowers from the previous summer, Rutherglen bugs may survive on fallen sunflower seeds or weeds through winter and migrate to cotton as the seedlings emerge. They suck cotton seedlings dry causing gaps in the crop. There have been no reports of damage to squaring or flowering cotton and they are unlikely to damage mature cotton. Monitoring: If high bug numbers are found in seedling cotton, monitor plant stands, and if in more mature crops, monitor for damage by cutting fruit open to check for blackened stamens in squares and staining of the seeds in young bolls.
Conservation of natural enemies: A number of generalist predators are known to feed on mirids. These include damsel bugs, big-eyed bugs, assassin bugs, predatory shield bugs, spiders and ants. Specific parasitoids that attack mirids have yet to be identified in Australia. Mirids are themselves predators and are known to eat mites and Helicoverpa spp. eggs.
Thrips
Beneficials may be conserved by using a selective insecticide early in the season, moving to harder insecticides later in the season if required. This helps to reduce the risk of secondary pest outbreaks that will also prove difficult and expensive to manage, as well as selecting for pesticide resistance in the secondary pests. Use of reduced rates of insecticides combined with salt or petroleum spray oils may also reduce impacts on beneficials. Information on the effect of insecticides on beneficial insects can be found on the Cotton CRC website.
Distribution: All three species are spread widely across the world and are exotic species to Australia. T. tabaci is found in all states, F. schultzei in all states except Tas. and F. occidentalis in all states except the NT. F. occidentalis was found in cotton regions during 2002.
Rutherglen bug Nysius vinitor Bergroth Hemiptera: Lygaeidae N (Main entry in Chapter 5 Oilseeds.)
Thysanoptera: Thripidae Tobacco thrips, Thrips tabaci Lindeman Tomato thrips, Frankliniella schultzei (Trybom) Western flower thrips, F. occidentalis Pergande
Pest status: Major, widespread, regular. Identification: All three species of thrips are small (about 2 mm in length) and cigar shaped (Fig. 3.7). Colouration is not a good indicator of species, but T. tabaci (Fig. 3.7a) and F. occidentalis adults tend to be straw-coloured, while those of F. schultzei are dark, almost black. However, F. schultzei also has a pale form, mostly in the warmer tropical areas. T. tabaci is more abundant on seedling cotton (up to about 50 days after emergence) and F. schultzei and F. occidentalis are
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more abundant on flowering cotton. F. schultzei in particular can often be found in large numbers in cotton flowers. First-instar larvae of all species are small (about 0.7 mm long) and pale white, while second-instar larvae are bright yellow and larger (1.5 mm). May be confused with: A range of other thrips species occur on cotton but generally at low numbers. The main source of confusion is between the three species listed. However, a fourth species, Scolothrips sexmaculatis (Pergande) (six-spotted thrips, Fig. 3.7b) is often found on unsprayed or low spray-input cotton late in the cotton season if spider mites are present. This species is an obligate mite predator and is effective in controlling mite populations. Adults of S. maculatus can be distinguished from other species by the presence of six spots, in three pairs, along the wings (a hand lens is adequate to distinguish these) (Fig. 3.7). Larvae can also be distinguished by being almost clear in colouration, and having the last two instars (3 and 4), which have obvious wing buds, complete development on the plant. Thrips may also be confused with springtails (Collembola). These insects are similar in size to thrips but are grey and soft-bodied. During cool, wet weather they may attack cotton, causing damage similar to thrips. Host range: All three species have wide host ranges, including many crops, broadleafed weeds, grasses and native species. Life cycle on cotton: Reproduction in thrips is unusual in that although they can reproduce sexually, they more often reproduce asexually. a
b
Fig. 3.7. (a) Adult tobacco thrips (1.5 mm long), a pest of cotton (CSIRO: C. Mares) , and (b) a beneficial thrips, sixspotted thrips (1.5 mm long), on the left of which is an egg of the two-spotted mite (CSIRO: L. Wilson) .
Unfertilised adult females of T. tabaci produce female young and, as a consequence, males are generally much less abundant than females. In contrast, unfertilised adult females of F. schultzei and F. occidentalis produce only male young. Female thrips have a saw-like ovipositor used to cut a slit, usually in the terminal of young leaves and in parts of flowers, into which they lay their eggs. The eggs hatch and thrips pass through two ‘larval’ stages and two ‘pupal’ stages. The larvae range from 0.5 to 1 mm in length and resemble miniature adults without wings. They are usually translucent white to yellow in colour and are frequently seen in the terminal. Before moulting, the second-stage larvae drop from the plant onto the soil or litter. Here they pass through the pre-pupa and pupa stages, which are presumably non-feeding. These two ‘pupal’ stages also resemble miniature adults but they have wing-buds. Thrips do not appear to overwinter in the soil, preferring sheltered positions on suitable host plants. The duration from egg through to egg producing adult ranges from 15 days at 30°C to 22.4 days at 20°C for F. schultzei, 18 days at 26°C and 35–45 days at 15°C for F. occidentalis, and from 16.1 days at 30°C to 28.6 days at 20°C for T. tabaci (Waterhouse and Norris 1989; Bournier 1994). Risk period: Thrips are a pest of seedling cotton mainly in eastern Australia. Typically, thrips infest cotton crops from emergence, and their feeding can cause growth distortion (Wilson and Bauer 1993). Thrips populations generally decline as plants approach squaring (production of flower buds). Thrips are found in cotton flowers and on the leaves in the mid to late season but they do not cause economic damage. Damage: Thrips feeding causes leaf distortion and can cause tipping out (death of the apical meristem) with a subsequent delay in plant development while a new terminal is produced. However, plants may compensate for this damage without loss of yield or delay in maturity (Sadras and Wilson 1998). Heavy damage can result in excessive tipping out throughout a cotton field and consequent delays in crop growth and maturity, which, in association with other damage, may cause yield loss (Forrester and Wilson 1988). In rare cases, seedling death necessitates replanting. Thrips may also feed on young cotton buds (squares) 79
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causing them to shed, although this is not common. Monitoring: Sample seedlings from emergence until plants have six to eight true leaves. Pull plants from the soil and quickly count the number of adult thrips (with wings), then count the number of larvae. Check within the plant terminal by teasing it apart with the tip of a pencil. Count thrips larvae and adults that walk out of the terminal. Also, look at the growing point and carefully check if the tip has been destroyed. Score if the true leaves are less than 1 cm in length. Sample at least 20–30 plants at random across the field. Cotton crops adjacent to cereal crops are especially at risk from thrips and should be monitored closely for signs of thrips damage. Action level: Plants will often compensate for thrips damage, so a threshold that combines both pest abundance and plant damage is used. Control would be warranted if (i) there are more than 10 thrips per plant, and (ii) most leaves are less than 1 cm long, which represents a loss of leaf area of greater than 80%. Tip damage is unlikely to occur at these threshold levels, but if thrips are causing tipping out then control should be considered. Take care in assessing tip damage as it may be due to other pests. If there are signs of tissue removal then damage is probably due to Helicoverpa spp. or other Lepidopteran pests. Chemical control: Control options fall into two groups: (i) seed- and soil-applied preventive controls (i.e. seed treatments or atplanting insecticides applied with the seed), and (ii) foliar applied insecticides. Preventive application should be based on expected yield loss. In northern Australia, thrips are not pests. In the hotter cotton regions of eastern Australia (Central Qld, St George, Bourke) thrips rarely cause economic damage. In the warm regions (McIntyre, Gwydir, Lower Namoi Valleys) thrips cause economic damage about 1 year in 10, mainly when abundant winter growth of weeds and cereal crops senesces in early spring, forcing thrips to seek alternative hosts. In these seasons, the use of a preventive treatment is justified. In the cooler regions (Darling Downs, Upper Namoi, Macquarie Valley and Lachlan) thrips will cause damage 1 year in 2 so a preventative
insecticide should be used. Choice of preventative insecticide should take into account effects on other pests, such as spider mites and selection for resistance, especially in aphids. The preventive insecticides are systemic in action and have relatively little impact on beneficial species. Foliar sprays for thrips are effective and inexpensive, especially if applied as a banded application by a ground sprayer. However, they are generally broadspectrum and disruptive to natural enemy populations, which can lead to increased survival of Helicoverpa spp. and increased risk of outbreaks of mites or aphids. F. occidentalis has yet to establish as a seedling pest of cotton, but in the event that it does, it may prove harder to control due to resistance to a wide range of insecticides. Cultural control: Avoid planting cotton crops next to cereal crops. Host-plant resistance: None of the commercial cotton varieties available show useful resistance to thrips. Conservation of natural enemies: A range of predators will attack thrips, most notably minute pirate bugs (Orius spp.) In addition to being pests, thrips are also predators of mite eggs and can play a significant role in preventing mite outbreaks especially through the early season (Wilson et al. 1996). In the mid to late cotton season, F. occidentalis is often abundant and feeding damage can be seen along the veins on the undersides of leaves in the upper canopy, but such damage is unlikely to affect yield. Instead, these thrips will often control mite outbreaks and control is rarely, if ever, warranted.
True wireworm Agrypnus variabilis (Candeze) Coleoptera: Elateridae N (Main entry in Chapter 2 Cereals.)
False wireworms Coleoptera: Tenebrionidae N (Main entry in Chapter 5 Oilseeds.)
Eastern false wireworm, Pterohelaeus darlingensis Carter
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Large false wireworm, Pterohelaeus alternatus Pascoe
a
Southern false wireworm, Gonocephalum macleayi (Blackburn) Saragus spp. Pest status on cotton: Minor, widespread, regular. Life cycle in cotton: Eggs are laid in summer and autumn, and larvae feed on organic matter in the soil until they reach full size in spring. True wireworms prefer wet soil for egg-laying (e.g. irrigated summer crops), whereas false wireworms prefer drier conditions protected by stubble or weeds. Spring infestations are generally worse in newly developed fields, in fallow fields (particularly with heavy trash cover) and in fields following summer crops (particularly soybeans). Damage: Both damage cotton by either boring into germinating seeds or chewing through young seedlings just below ground level. Damage is unlikely in fields following conventional cotton, as insecticide use usually prevents beetle survival, however Bollgard II® fields require fewer pesticide applications allowing more soil insects to survive. Risk period: Damage can occur for up to 4 weeks after sowing, resulting in patchy plant stands which may require replanting. Monitoring: Wireworms are difficult to sample. They descend in the soil as it dries out and venture near the surface only after rain or an irrigation event and are often not noticed until damage appears, by which time it is not possible to control them. Chemical control: Insecticides must be applied before, or at sowing. The most effective control is an in-furrow band application of either a granular or liquid insecticide.
Flea beetles Coleoptera: Chrysomelidae Redheaded flea beetle, Nisotra sp. Brown flea beetle, Chaetocnema sp. Pest status on cotton: Minor, restricted, irregular. Seedling damage caused by flea beetles is
b
Fig. 3.8. (a) Redheaded flea beetle (4 mm), which eats small shot holes in cotton leaves, and (b) characteristic linear damage from the brown flea beetle. (CSIRO: C. Mares)
reported occasionally. They are small, shiny, metallic beetles of the sub-family Halticinae, noted for their jumping ability. At least two species have been observed in seedling cotton: the redheaded flea beetle (Fig. 3.8a) and the brown flea beetle. The redheaded flea beetle causes characteristic shot hole perforations to leaves but rarely causes sufficient damage for control to be warranted. The brown flea beetle causes linear surface feeding patterns particularly on the cotyledons (Fig. 3.8b). Occasionally, infestations of this flea beetle are so large that control measures are needed to prevent desiccation of seedlings.
Cotton tipworm or cotton tip borer Crocidosema plebejana Zeller Lepidoptera: Tortricidae Pest status on cotton: Minor, widespread, irregular. Identification: The cotton tipworm is the larva (Fig. 3.9a) of a moth that infests a range of malvaceous plants including cotton. The small moth (Fig. 3.9c) lays oval-shaped, flattened, translucent eggs which are placed singly, often next to a leaf vein on the underside of a leaf or among the small terminal leaves. As the egg develops its colour becomes whiter and a ‘red ring’ develops just before hatching. Host range: Marshmallow weed (Malva parviflora) is a preferred host and the abundance of tipworm in cotton is closely related to the 81
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a
b
Tipworm damage is generally less important after commencement of vegetative growth and square production. On the rare occasions that conditions allow cotton tipworm to reach high numbers in fruiting cotton, squares and bolls may also be damaged. Action level: The current thresholds for control are one or two larvae per metre, and 100–200% of plants damaged lightly (e.g. just the terminal damaged, larvae not entrenched) or 50–100% of plants tip damaged heavily once (larvae entrenched). Chemical control: Control is sometimes possible when eggs have just hatched, before the larvae bore into the plant terminal. Use of broadspectrum insecticides for control often provides poor efficacy and destroys natural enemies, leading to a resurgence of tipworm and outbreaks of secondary pests such as spider mites.
c
Lightbrown apple moth or tortrix Epiphyas postvittana (Walker) Lepidoptera: Tortricidae Pest status on cotton: Minor, restricted, irregular.
Fig. 3.9. (a) Cotton tip worm neonate (CSIRO: C. Mares) , (b) tipworm in stem (5 mm) (CSIRO: C. Mares) , and (c) tipworm adults (CSIRO: C. Mares) .
abundance of marshmallow in the winter and spring in cotton regions. Damage: Tipworm larvae tunnel into the terminal (Fig. 3.9b) destroying the single stem habit of cotton seedlings and causing multiple branching. If this damage occurs in the seedling or early squaring stages of crop growth, it may result in increased vegetative growth (branching) at the expense of early reproductive growth, and crop development may be delayed. Crops may compensate for this damage often without loss of yield and only a small delay in maturity (< 7 days as damaged plants have more branches each producing fruit so they catch up). However, multiple tip damage to each plant can cause longer delays in maturity, which if compounded by other adverse factors (e.g. cool weather) can reduce fibre quality and yield.
Identification: The pale, greenish-yellow eggs are laid in flat masses of 20 to 30 eggs overlapping each other like fish scales. These egg masses are not covered in moth scales like those of the lesser armyworm, Spodoptera exigua. Young larvae feed by tying the terminal leaves together with a fine web, which becomes dotted with pellets of frass. Older larvae vary from light to dark green with a light brown head and can grow up to 20 mm long. They pupate in silk-lined burrows in the soil after feeding for about 3 weeks. Adults are brown, males are smaller than females and are two-toned brown (Fig. 3.10). May be confused with: Young larvae superficially resemble small tipworm larvae, having a black head and pale body. However, they can easily be distinguished from tipworm larvae by the presence of webbing and the fast, twisting movement of the larvae when prodded. Host range: Includes many broadleaved weeds and crops such as lucerne. Damage: Lightbrown apple moth can be found infesting cotton seedlings in low numbers, particularly in cool seasons but under hot, dry
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Fig. 3.10. Female (left, 12 mm) and male (right, 8 mm) moths of the light brown apple moth. (SARDI: G.J. Baker)
conditions it is seldom present. Damage to cotton is very rarely important.
Cutworms
Lesser armyworm is often present in low numbers on young cotton, but prefers weed hosts. Larvae may occasionally infest seedlings heavily enough to cause defoliation, and resowing may be necessary. The moth is about 10 mm in length with grey, mottled forewings. The hindwing is of a pearlywhite colour. Eggs are laid in ‘rafts’ of 10 to 30 and are covered by webbing-like scales by the female moth, similar to the cluster caterpillar. The young larvae remain near the egg raft and skeletonise the leaf. Mature larvae are often mistaken for Helicoverpa spp. but are green to brown, about half the length of a mature Helicoverpa larva, with a white stripe along each side of the back. The threshold for control of lesser armyworm is two small larvae or one large larva per metre from emergence to flowering.
Cluster caterpillar
Agrotis spp. Lepidoptera: Noctuidae
Spodoptera litura (Fabricius) Lepidoptera: Noctuidae
N (Main entry in Chapter 2 Cereals.)
N (Main entry in Chapter 7 Pulses—summer.)
Pest status on cotton: Minor, widespread, irregular.
Pest status on cotton: Minor, widespread, regular.
Cutworms are the larvae of the Bogong moth, Agrotis infusa (Boisduval) and related species. The moths are brown to black and slightly larger than a Helicoverpa spp. moth. They lay white eggs in the soil or on seedlings. Younger larvae feed on the leaves of seedlings and may be mistaken for Helicoverpa spp., tortrix or tipworm larvae. Older larvae are dark grey to brown, smooth-bodied, and 25 to 50 mm in length when fully grown. They feed in late afternoon and at night, spending the days hidden in the soil, except under very overcast conditions.
The cluster caterpillar (Fig. 7.129) was a serious cotton pest in the Ord River (WA) but not in eastern Australia. Its habits resemble those of cutworms except that eggs and small larvae are always found in groups. The white eggs are laid under leaves and are covered with fine, brown hair-like scales from the female moth. Young larvae are grey–brown and they skeletonise leaves at night. Larger larvae consume whole leaves and heavy infestations defoliate large areas, destroying squares and flowers as well. Pupation occurs in the soil and the adults have dark brown forewings patterned with grey, white hindwings bordered with grey, and a wingspan of about 35 mm (Fig. 7.125b).
Cutworms chew cotton seedlings at or above soil level and may destroy seedlings by either chopping the stem or eating entire leaves. Insecticide treatment for cutworms should be made in late afternoon or evening to minimise insecticide degradation prior to larvae emerging to feed.
Lesser armyworm Spodoptera exigua (Hübner) Lepidoptera: Noctuidae Pest status on cotton: Minor, widespread, regular.
S Q UA R I N G A N D B O L L- S E T T I N G S TAG E S
Two-spotted mite Tetranychus urticae (Koch) Acarina: Tetranychidae Distribution: Cosmopolitan, widely distributed throughout the world on agricultural, 83
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Life cycle on cotton: The TSM develops from egg to adult in about 7 days at 30ºC resulting in multiple overlapping generations on cotton through the summer. Eggs typically hatch after 1–2 days. The first stage is the larva, which is small, translucent white with six legs. The larva moults into a protonymph, a deutonymph then to an adult. Between each moulting stage are resting stages, known respectively as protochrysalis, deutochrysalis and teliochrysalis.
Fig. 3.11. Three adult two-spotted mites (0.5 mm long) with a nymph and an egg. (Graphic Science: Dennis Crawford)
horticultural and glasshouse crops. Widely distributed in Australia, found in all states. Pest status: Major, widespread , irregular. Identification: The two-spotted mite (TSM) is a small arthropod belonging to the group including ticks and mites. It is typically a pale green colour with darker green spots on either side of the body (Fig. 3.11). Adult female mites are about 0.5 mm long, eggs are small (0.1 mm diameter), round and almost clear (Fig. 3.11). The TSM typically forms colonies on the undersides of leaves especially near the junction of the petiole and leaf blade, and in leaf folds. Damage symptoms are bronzed areas on the upper leaf surface. May be confused with: The TSM may be confused with other spider mite species on cotton. These include Tetranychus lambi Pritchard and Baker (strawberry spider mite), which is smaller (adult females are about 0.3 mm in length), and has six smaller dark green spots rather than two large spots. This species rarely damages cotton and does not cause the bronzing of leaves. The other species is Tetranychus ludeni Zacher (bean spider mite), which is similar in size and damage to TSM, but the adult females are a dark red or carmine colour. Although potentially a damaging pest, this species is generally scarce. Host range: The TSM has a wide host range including many crop, broadleaf weed and grass species (Wilson 1995). Some key non-cultivated hosts include turnip weed, wireweed, burr medic, sowthistle and marshmallow weed.
The female teliochrysalis produces a pheromone attractive to male mites. Mated females produce male and female eggs in the ratio of 1:3, whereas unmated females produce only male eggs. In winter, mites mainly survive on available hosts, but in cooler regions some may change into the bright orange diapause form. This form is nonfeeding and non-reproductive and seeks sheltered locations in cracks in the soil and litter to survive through winter. Mortality of the diapause form in cotton regions is typically very high. Risk period: The TSM can damage cotton at any time of the growing season, but most often during mid to late season (January–March). Cotton crops are colonised as seedlings by mites migrating from nearby weeds or crop hosts on which they have overwintered. These overwintering populations consist of actively feeding and reproducing mites, although mite development is slow due to lower temperatures. The degree of infestation of seedling cotton determines the potential for mite outbreaks to develop, with higher infestation levels (> 10% of plants infested) having a greater risk of populations developing that could cause yield loss. Management also strongly influences whether early populations develop to damaging levels. Disruption of natural enemies by insecticides can allow mite populations to increase quickly (Wilson et al. 1998). Damage: Damage by the TSM generally first appears on the younger leaves (mainstem nodes leaves 3–5 below the plant terminal). Feeding damage is visible on the upper surface of leaves as deep maroon or carmine areas (Fig. 3.12). The effect of mite damage in plant development and ultimately yields is cumulative; that is, yield losses are more severe the longer mites are on the crop and the more quickly their populations increase. Populations
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a
Sampling protocol for mites on cotton
b
Fig. 3.12. (a) Okra leaf cotton varieties are more resistant to spider mites than (b) normal leaf varieties. (CSIRO: C. Mares)
that increase early in the season and quickly can dramatically reduce yield if left uncontrolled (losses of 90% of yield have been recorded). Populations that increase slowly late in the season may not affect yield at all. Beneficials can strongly influence the development of mite populations, and thereby their potential effect on yield. Monitoring: A presence/absence sampling technique allows rapid sampling (Wilson and Morton 1993), increasing the chances of finding scattered mite populations. As the effect of mites on yield depends on the timing and rate of mite increase, the most useful measure of damage potential is the rate of population change over time, which can be estimated by taking sequential samples or sampling at least weekly (Wilson 1993). Action level: Thresholds for mites have been developed (see p. 86) which take into account both population density and rate of increase (Wilson 1993). The decision support system, CottonLOGIC, includes mite sampling information and calculation of potential yield losses. CottonLOGIC can be run on handheld computers in the field. A detailed description of the thresholds can be found in the Cotton pest
1 Walk into the field about 40 m. (Early in the season it is also advisable to sample near the field edges to see if significant influxes of mites have occurred.) 2 Take a leaf from the first plant on the right or left. The leaf should be from the third, fourth or fifth mainstem node below the terminal. If the plant has less than three leaves sample the oldest. Note that early in the season, up to the point that the plant has about five true leaves, it is simplest to pull out whole plants. 3 Walk five steps and take a leaf from the next plant, on the opposite side to the previous one, and so on until you have 50 leaves. (Wait until you have collected all the leaves before scoring them.) 4 Once all the leaves have been collected, score each leaf by turning it over, looking at the underside, firstly near the stalk, then scanning the rest of the leaf with a hand lens. If mites of any stage (eggs or motiles) are present, score the leaf as infested. 5 Repeat this simple procedure at several widely separated places in the field to allow for differences in mite abundance within the field. Depending on the size of the field, between four to six sites are needed to obtain a good estimate of mite abundance. 6 When finished sampling, calculate the percentage of plants infested in the field.
management guide, published annually by NSW DPI, or on the Cotton CRC website. Chemical control: A number of acaricides are available and effective for control of TSM. Some TSM populations are resistant to particular acaricides (Herron et al. 1998, 2001 and 2004); information on resistance levels of TSM can be obtained from NSW DPI, which is the centre for TSM-resistance testing in Australia. Cultural control: Good farm hygiene is important, especially through autumn, winter and spring to reduce the availability of hosts on which TSM can overwinter. Reducing the size of the overwintering population reduces the level 85
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Thresholds for mites on cotton 1 Seedling emergence to squaring: Mites are normally suppressed by predators, especially thrips during this period. Mite populations only need to be controlled if they begin to increase, which indicates that natural controls are not keeping them in check. 2 Squaring to first open bolls: Control if mite populations increase at greater than 1% of plants infested per day in two consecutive checks or if more than 30% of plants are infested. 3 First open bolls to 20% open bolls: Control is only warranted if mites are well established (greater than 60% plants infested) and are increasing rapidly (faster than 3% of plants infested per day). 4. Crop exceeds 20% open bolls: Control is no longer warranted.
of colonisation of seedling cotton. Selection of rotation crops is also important as some, such as safflower, are good hosts for mites which can migrate to nearby cotton crops when the safflower senesces late in spring. Host-plant resistance: Mite populations develop more slowly, with less potential for yield loss, on okra leaf cotton varieties than normal leaf varieties (Fig. 3.12), typically saving about one acaricide application (Wilson 1994, 1995). Okra leaf shape provides less protection of mite eggs from desiccation. Bollgard II® cotton reduces the need to spray the crop for lepidopteran pests, thereby helping to conserve beneficials that will contribute to control of TSM. Conservation of natural enemies: Beneficials play a major role in controlling or delaying the development of TSM populations in cotton and every effort should be made to conserve them. These include pest thrips that are also facultative predators of mite eggs (Thrips tabaci, Frankliniella schultzei, F. occidentalis) and the predatory sixspotted thrips (Scolothrips sexmaculatus) (Fig. 3.7), predatory bugs (Geocoris lubra Kirkaldy Lygaeidae (Fig. 3.35), Nabis kinbergii Reuter Nabidae (Fig. 3.37), Orius spp. Anthocoridae, Campylomma liebknechti (Girault) Miridae), ladybeetles (especially Stethorus spp. (Fig. 15.54)
and Diomus notescens (Fig. 3.49). Chemicals used to control other pests often kill these natural enemies, allowing TSM to increase unchecked, hence their reputation as ‘secondary’ pests. Information is available on the Australian Cotton CRC website and in the Cotton Pest Management Guide on the effects of all registered insecticides and acaricides on beneficial insect groups.
Mealybugs Hemiptera: Pseudococcidae Pest status on cotton: Minor, restricted, irregular. Mealybugs are small, sucking insects related to aphids. They are occasionally seen on cotton in most areas and have, on rare occasions, reached minor outbreak levels or ‘hotspots’ in commercial crops in central Qld. Species include hibiscus mealybug, Maconellicoccus hirsutus (Green), striped mealybug, Ferrisia virgata (Cockerell) and Phenacoccus solani (Ferris). They form colonies on stems and leaves developing into dense, waxy, white masses. Infestations are usually close to field borders. Control is seldom warranted.
Apple dimpling bug Campylomma liebknechti (Girault) Hemiptera: Miridae Pest status on cotton: Minor, widespread, regular. The apple dimpling bug (yellow mirid) is about one-third the size of the green mirid. Adults are generally pale green with spines on the legs. The nymphs are small and yellow in colour. The apple dimpling bug damages small squares (flower buds) early in the fruiting cycle. It is generally regarded more as a predator of Helicoverpa spp. eggs and spider mites than a pest. Apple dimpling bugs are best sampled by visual inspection of the crop. Beat sheet sampling is also effective at sampling adults but the yellow nymphs are very hard to see against a yellow beat sheet. The threshold for the apple dimpling bug is generally five times the threshold for the green mirid. If the apple dimpling bug is in high numbers it is worthwhile to monitor the plant’s fruit set by either checking fruit retention or by
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using the ‘fruiting factor’ technique (Deutscher et al. 2004). This will indicate if damage is occurring which may require control. A range of insecticides is available for control, including some that have relatively low negative effects on beneficial populations.
Cotton seed bug
a
b
Oxycarenus luctuosus (Montrouzier) Hemiptera: Lygaeidae Pest Status on cotton: Minor, widespread, regular. Adult cotton seed bugs (Fig. 3.13) can be found sheltering on cotton plants from the late seedling stage onwards but do not feed or reproduce until cotton bolls open and ripe seeds are available. The eggs are laid in open bolls, and the bright red nymphs can be found in clusters among the lint (Fig. 3.13a). Cotton seed bug feeding may reduce seed weight in late bolls by as much as 15% and also reduce viability. Because this damage tends to be only to seeds in late bolls, the cotton seed bug does not normally require control. a
Fig. 3.14. (a) Flower beetle larva (2.5 mm) (CSIRO: P. Room) , and (b) adults (3.5 mm) (CSIRO: P. Room) .
Flower beetles Carpophilus spp. (Jacoby) Coleoptera: Nitidulidae Pest status on cotton: Minor, widespread, regular. Small groups of black beetles, flower beetles, Carpophilus spp. are a common sight in cotton flowers. The larvae are white with a pale brown head. Adults (Fig. 3.14b) and larvae (Fig. 3.14a) appear to feed on pollen but do not cause any damage.
Redshouldered leaf beetle Monolepta australis (Jacoby) Coleoptera: Chrysomelidae
b
Fig. 3.13. (a) cottonseed bug adults and nymphs (CSIRO: , and (b) adult cotton seed bug (3.5 mm) (CSIRO: C. Mares) .
C. Mares)
Pest status on cotton: Minor, restricted, irregular. The redshouldered leaf beetle is 7 mm long, golden yellow in colour, with a red spot on each wing and a red band on the wings directly behind the head (Fig. 7.79). The redshouldered leaf beetle rarely attacks cotton, and when it does, damage is usually confined to the edges of fields. It chews terminal buds, leaves (Fig. 3.15), squares and the surfaces of bolls. Breeding apparently takes place in bushland where the eggs are laid in the soil. Larvae feed on plant roots and pupation is in the soil. Adults also feed on maize, citrus, avocado, macadamia and stone fruits. The redshouldered leaf beetle is 87
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a
Fig. 3.15. Redshouldered leaf beetle damage. (NSWDPI:
b
A. Bishop)
usually active from October to April and has three to four generations per year.
Cotton leaf perforator Bucculatrix gossypii Turner Lepidoptera: Buccalatricidae
c
Pest status on cotton: Minor, restricted, irregular. The cotton leaf perforator occurs mainly in Qld but has been recorded on occasions in NSW. The young larvae, instars 1–3, mine between the upper and lower surfaces of leaves creating characteristic trails (Fig. 3.16d). The fourth instars emerge and feed directly on the leaf surface. The fourth-instar larva forms a thin, silk shelter in which it moults into the fifth instar. The fifth instar causes the most obvious damage, skeletonising the leaf by feeding on the lower leaf surface leaving numerous windows and small holes (Fig. 3.16c). Older larvae (less than 10 mm long) are pale grey with four black spots just behind the head. Pupae are formed in white, ribbed cocoons attached to the leaf (Fig. 3.16a). Adults are cream–grey moths with wings that are densely fringed with hairs, particularly the hind wings (Fig. 3.16b). Damage rarely warrants control. Larvae are occasionally noticed surviving early season endosulfan sprays for Helicoverpa spp. and are often mistaken for endosulfan resistant H. armigera. They are, however, not resistant to endosulfan, merely protected from the spray within their mines from which they emerge as late-instar larvae.
Cotton looper Anomis flava (Fabricius) Lepidoptera: Noctuidae
d
Fig. 3.16. (a) Cotton leaf perforator in the early leafmining stage, (b) older larva of the cotton leaf perforator and damage (5 mm), (c) pupa of cotton leaf perforator (8 mm), and (d) adult cotton leaf perforator (4 mm). (CSIRO: C. Mares)
Pest status on cotton: Minor, widespread, regular. The cotton looper is named after the looping movement of its greenish larva (Fig. 3.17b). The small, bluish-green eggs are laid on leaves (Fig. 3.17a). The larvae prefer to eat older leaves and defoliation (up to 80%) progresses upwards on the plant (Fig. 3.17). The pupae are found in rolled up leaves and emerge as brown moths with patterned wings. Sprays applied against Helicoverpa spp. coincidentally control looper infestations and the species is well-controlled by insect-resistant transgenic cotton varieties
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a
b
Fig. 3.17. (a) Eggs of cotton looper (CSIRO: C. Mares) , and (b) larva of cotton looper (40 mm) (CSIRO: C. Mares) .
containing the Cry1Ac and Cry2Ab proteins (Bollgard II ® ).
Native budworm Helicoverpa punctigera Wallengren Lepidoptera: Noctuidae
Cotton bollworm or corn earworm Helicoverpa armigera Hübner Lepidoptera: Noctuidae For crop specific information about Risk period, Damage, Monitoring, Action level and Control on crops other than cotton, see the relevant crop chapter. Distribution: H. punctigera is endemic to Australia and is widespread throughout all states in both summer and winter cropping regions and in natural rangelands. It has been recorded as an occasional migrant to New Zealand, Norfolk Island and the Cocos Islands, although the existence of breeding populations has not been established in these locations. H. armigera is distributed in southern Europe, Africa, east and south-east Asia to Australia, Papua, and many Pacific islands. In Australia, it
is more common in eastern Australia, particularly in crop-growing regions, and in northern Australia, and declines in abundance towards the south but is recorded in SA and Vic. as a pest of summer sweet corn. H. armigera extends into the arid inland and can be found in low numbers on several native plants (Zalucki et al. 1994). Pest status: Both species: major, widespread, regular. H. punctigera and H. armigera combined represent the most significant insect pests of broadacre agriculture in Australia. Estimated total costs of control and damage are up to $328 million. The proportion of this total cost due to H. punctigera is difficult to estimate, since both species feed on many crops. Although H. armigera is regarded as the major pest because of its capacity to develop pesticide resistance and its difficulty to control, H. punctigera may often be the most numerically abundant of the two species. H. punctigera occurs on crops most commonly during spring and early summer, where it feeds on maturing winter crops then summer crops such as sunflower and cotton. Much of the cost of control and damage on grain legumes in SA, Vic. and WA is due to H. punctigera. Identification: Heliothinae is a sub-family of noctuid moths containing the Genera Adisura, Australothis, Helicoverpa, Heliocheilus and Heliothis, which occur in Australia. The species are distinguished by their genitalia, details of which may be found in Matthews (1999). The most abundant and frequently occurring heliothine moths found in Australian field crops are Helicoverpa armigera and H. punctigera. Both of these species were previously assigned to the genus Heliothis. Heliothis rubrescens (Walker) has been recorded on tobacco in Qld (Titmarsh et al. 1990), and huge flights of Heliothis punctifera (previously Neocleptria punctifera) Walker are occasionally recorded across eastern and southern Australia (Hopkins, unpublished). Eggs: Eggs are spheroidal, being slightly wider than high. When newly laid, eggs are pale yellow–white, turning to pale brown within 36 hours (at 25°C) (Fig. 3.18) and then briefly to black just prior to hatching (Fig. 3.18). Larvae: The larvae exhibit many colour forms, but are predominantly green, brown, black or pink (Fig. 3.19). They have three pairs of thoracic 89
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Fig. 3.18. Developmental stages of Helicoverpa eggs. Newly laid, white eggs develop through the brown ring stage (middle) to the pre-emergent black head stage. The distinct black head of the larva distinguish healthy eggs from parasitised eggs. (DPI&F Qld: B. Scholz)
(‘true’) legs, four pairs of abdominal pro-legs and a pair of anal pro-legs (Fig. 3.20). Early instar larvae are morphologically indistinguishable until the third instars (Fig. 3.20), when a characteristic saddle usually, but not invariably, develops on larvae of H. armigera (Fig. 7.127) (Stanley 1978). H. punctigera larvae do not have saddle markings on the first and second abdominal segments. Final-instar larvae may be distinguished by the colour of the setae on the dorsal part of the prothoracic shield, just behind the head. These are black in H. punctigera and light in H. armigera (Fig. 3.21). All immature stages of H. punctigera can be distinguished from H. armigera using antibody-based test kits not presently available in commercial form (Trowell et al. 2000). Pupae: The pupae are brown to dark brown. Pupae of the two species may the distinguished
Fig. 3.20. Developmental stages of Helicoverpa punctigera eggs and larvae. Newly laid egg (white) and mature egg (black) with six larval instars. (SARDI: G. Caon)
by two small tail spines (cremastal spines) present on the final (10th) abdominal segment (Fig. 3.22), which are separated by a gap approximately equal to their diameter in H. punctigera, whereas the gap is approximately twice their diameter in H. armigera. Adults: H. punctigera is similar in appearance to H. armigera and many errors in identification have undoubtedly occurred prior to Common (1953) and Hardwick (1965) providing distinguishing features of adult moths. Both are medium-sized to large, straw coloured to brown, heliothine moths. The forewing is variably marked. The hindwing has a well-defined dark marginal band with a pale basal patch when viewed from above (Fig. 3.23). Males may generally be paler in coloration than females in both species. Colour may be influenced by temperature; characteristically dark adults follow exposure of H. punctigera pupae to low temperatures (Hill 1993). Matthews (1999) provides taxonomic descriptions of both species. a
Fig. 3.19. Variations in colour and markings of Helicoverpa armigera larvae. (DPI&F Qld)
b
Fig. 3.21. (a) Fully grown larva of H. punctigera, showing dark hairs on the head and anterior thorax, and (b) larva of H armigera showing pale hairs on the anterior thorax. (DPI&F Qld: J. Wessels)
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a
b
Fig. 3.22. Posterior end of a pupa of (a) Helicoverpa punctigera, and (b) H. armigera showing the cremastal spines at the end of the pupae. The two species may be distinguished by the distance between the cremastal spines. (DPI&F Qld: J. Wessels)
Adults of the two species can generally be distinguished in the following ways (Fig. 3.23): • H. punctigera is usually more greyish brown or straw coloured than H. armigera, which is a more orange (‘warmer’) brown or greenish. • In the hindwing, the contrast between the dark outer band and the paler inner part is more pronounced in H. punctigera than in H. armigera. • H. armigera generally has a pale area (window) in the darkened hindwing margin. The two species may also be distinguished by both male and female genitalia, details of which are given by Matthews (1999). May be confused with: Helicoverpa assulta Guenée adult, like H. armigera, has a pale patch in the marginal dark patch of its hind wing
Fig. 3.23. Adult Helicoverpa armigera moths (right) have a pale area on the back edge of the hind wing (arrowed) not present in H. punctigera (left). (H. Brier)
when viewed from above. However, it is a smaller moth with the base of the hind wing a dull orange (Common 1990). Its larval food does not include field crops. Host range: Both Helicoverpa species are highly polyphagous. In Australia alone, H. armigera has been recorded on over 100 plant species from 36 families, while H. punctigera has been recorded on over 200 plant species from 47 families (Matthews 1999). Despite their wide host range, there are clear differences between the species. H. armigera feeds on both dicotyledonous and monocotyledonous plants, whereas H. punctigera occurs mostly on dicotyledonous plants although it occasionally attacks cereals (Chapter 2). Hosts of Helicoverpa armigera include nearly all the major field crops in Australia: cotton, sorghum, maize, sunflower, chickpea, lupins and most other legumes, lucerne, tobacco and wheat. In addition, horticultural crops such as tomatoes, lettuce, capsicum and sweet corn, and flowers such as chrysanthemums and roses suffer high rates of damage (Matthews 1999). H. armigera is more closely associated with cultivated crop hosts than with native host plants and its population dynamics is largely determined by cropping patterns in a region. By contrast, H. punctigera feeds on a wide range of native, ephemeral hosts across large parts of inland Australia as well as many crops. Most H. punctigera hosts are dicots, predominantly of the families Asteraceae, Fabaceae, Malvaceae, Solanaceae, Amaranthaceae and Brassicaceae. In cropping areas of eastern and western Australia H. punctigera feeds on cotton, cereals (Chapter 2), maize (Chapter 4), oilseeds (Chapter 5), poppies (Chapter 6), summer pulses (Chapter 7), winter pulses (Chapter 8), and tobacco (Chapter 12). There is some evidence for geographic variation in host-plant preference of H. punctigera (Firempong and Zalucki 1990); population genetics of host selection are yet to be examined. The species is well-adapted to exploit numerous plants in many families spread across inland and coastal parts of Australia. Many of these incidence records may not represent significant host plants (Walter and Benfield 1994). The highly generalised host selection pattern of H. punctigera and its extreme abundance in some seasons probably result in many instances of inadvertent oviposition on unusual hosts (e.g. 91
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records on citrus and pine trees). Similarly, the later-instar larvae may move freely among, and feed on, many plants in diverse communities. Life cycle on field crops: There are considerable differences in the seasonal dynamics of the two species in the crop growing areas of eastern Australia. Egg production in both species is influenced in part by adult size and diet. The mean number of eggs laid is 1400 to 2000, with a maximum number of around 3000 depending on female longevity. In southern Qld and much of NSW, H. armigera passes through four to five generations per season, and the winter in pupal diapause (Fitt 1989; Fitt and Daly 1990). Development time at 21.3°C constant temperature is 24 days for larval development and 22.6 days for pupal development, at 24.6°C the larval stages total 18.5 days duration and 15.7 days for pupal development, at 27.1°C larval development takes about 14.5 days and pupal duration is 11.5 days, and at 33.9°C larval stages are completed in 11.7 days and the pupal stage in a further 10 days (Twine 1978). Males take a little longer to develop than females. During the summer, subpopulations develop through first generation on spring weeds, winter legumes (chickpea), and on early sown sorghum or sunflower. During this period, populations are relatively synchronised, but later in the summer considerable overlap of generations occurs. The next two to three generations may occur on cotton, although some populations also breed on sorghum, sunflower, maize and legumes during the summer. Hence, at any one time, the H. armigera populations occur as a complex mosaic of sub-populations developing on many host plants, at slightly different rates, with differing levels of mortality and fitness effects on future fecundity, and there is considerable adult movement linking all sub-populations (Dillon et al. 1996). In southern cropping areas, H. punctigera has four to five overlapping generations between egg-laying of immigrants in spring and pupation of the last summer generation. During spring and summer, eggs hatch in about 7 days at 19°C and about 4 days at 25°C constant temperature. Most larvae in a population pass through six instars (Fig. 3.20), a few have an extra (seventh) instar. At 19°C constant
temperature, larvae complete development in about 36 days and pupal development takes a further 34 days. At 25°C all larval stages are completed in 17 days and pupal development in a further 16 days. Pupae may enter diapause starting from the first summer generation. Towards the end of summer an increasing proportion of pupae enter a winter diapause of up to 318 days duration (Cullen 1969). Non-migrating adults mate 1–2 days after emergence and may continue to mate throughout their lifetime of about 11 days in summer temperatures. Female moths need to feed on nectar for mating and to mature eggs. Plants producing nectar when moths are active are attractive to moths, especially plants with extra-floral nectaries. Some flight activity may be observed in crops during the day, but most flying, feeding and mating starts at dusk and continues into the night (Cullen 1969). Females of both species produce a pheromone to attract males. Females are capable of laying about 1400 eggs during their lifetime. Females appear to select rough surfaces on which to lay eggs in preference to smooth surfaces. The intensity of egg laying by H. punctigera on spring-flowering crops in southern cropping areas appears to depend on the time of arrival of migrating moths and the time of flowering of the crop in relation to flowering of weeds and other plants in the area surrounding the crop. If these weeds have been flowering for several weeks prior to crop flowering, moths lay most eggs on these weeds (Cullen 1969). Further, H. punctigera eggs that are laid on crop plants may suffer significant mortality. Cullen (1969) found between about 50 and 60% egg mortality mostly caused by rainfall washing eggs onto the ground. It has been difficult to relate spring moth catches in light or pheromone traps to subsequent crop damage. Low trap catches do not necessarily mean a low likelihood of crop damage, while high trap catches do not correlate well with crop damage. Nonetheless the timing of spring peaks of adults resulting from migration events can be used to predict the timing of subsequent population peaks. Larvae move on their host plant to eat leaves, buds, flowers and fruit (seeds) during their development. The importance of each of these appears to depend on their availability in
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relation to the stages of development of the larval population, which may vary between years. Migration: The seasonal pattern for H. punctigera in cropping areas is tied to rainfall patterns over wide areas of Australia (Maelzer and Zalucki 1999; Oertel et al. 1999). In some areas, phenology of H. punctigera is understandable only by invoking substantial migratory movement. In SA and WA, H. punctigera is rare during autumn, when there are few larval hosts, but reappears each spring (Cullen 1969; Walden 1995). Overwintering populations of H. punctigera are extremely difficult to locate in the irrigated areas in northern NSW and south-east Qld, where the overwintering populations are almost exclusively H. armigera (Fitt and Daly 1990). Despite this, large numbers of H. punctigera reappear in these regions each spring, as massive migrations from inland Australia recolonise the crop-growing regions (Gregg et al. 1995). Linking the spring peaks of moths caught in light traps in cropping areas to inland source areas involves a combination of evidence (Gregg et al. 1995; Rochester et al. 1996): • ground surveys of winter breeding habitats to detect the presence and age structure of larval populations • use of development models to predict emergence times of these generations • light trap catches and analysis of wind patterns associated with spring weather patterns, backtracking from trap locations to possible source areas. Catches of moths flying higher than 20 m indicate a migrating population. • presence of pollen of inland plants from source areas on moths trapped in cropping areas • the generally unmated status of trapped moths which suggests recent immigration Some of the possible source areas of moths that invade cropping areas during early spring are shown in Fig. 3.24, together with possible migration paths of the moths. In both WA and eastern Australia, moths appear in cropping areas during spring following warm north to
north-westerly winds which precede cold fronts traversing Australia at that time. Flights from inland breeding areas to cropping areas may take as little as one or two nights. The spring migrations generate breeding populations of H. punctigera on spring weeds and winter pulse and oilseed crops before moving to cotton and other crops in late spring and early summer. After the spring generation, H.punctigera generations progressively decline in abundance and become the minor species during the second half of summer (Cullen 1969; Wardhaugh et al. 1980; Wilson 1983; Fitt 1989; Maelzer and Zalucki 1999). This decline in population density, the scarcity or absence of overwintering populations of H. punctigera in many crop growing areas, and the near simultaneous appearance of large numbers in several areas each spring (September–October) indicate migration to be an important component of the life-history of this species. The extent of winter breeding on ephemeral native plants (Zalucki et al. 1994), and hence the magnitude of spring migratory events, is determined by the timing and extent of autumn rainfall through particular regions of arid inland Australia (Oertel et al. 1999) and to broad-scale climatic patterns such as the Southern Oscillation Index (Maelzer and Zalucki 1999, 2000). As a result it is possible, although presently difficult, to forecast long-distance movements (Rochester et al. 1996) or population dynamics of this species (Maelzer and Zalucki 2000). Risk period in cotton: All growth stages from seedling to maturity. Damage to cotton: The main damage caused by larval feeding is to squares, flowers and bolls, which are the preferred feeding sites. Feeding on stems and leaves is rarely extensive and has little effect on the numbers of fruiting points and mature bolls. Larvae damage seedlings by eating terminal buds. In maturing cotton, larval injury to squares, flowers and young bolls causes them to be shed, followed by compensatory production of new buds. Older bolls, when attacked, remain on the plant. Surface feeding on large bolls has little effect on lint and seed weights, but larval feeding inside bolls reduces seed weight, lint quality and often completely destroys lint (Fig. 3.25). 93
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Darwin
NT QLD
WA
Brisbane
Perth
NSW Adelaide
VIC Possible winter breeding areas of native budworm Possible spring migration routes of native budworm; curvature of the arrows indicates movement of moths across frontal weather patterns
ACT
Sydney
Canberra Melbourne
TAS Hobart
Fig. 3.24. Possible source areas and migration routes of spring flights of native budworm moths. From data in Gregg et al. 1995 and Walden 1995.
Monitoring on cotton: A number of sampling strategies have been utilised for Helicoverpa spp. in cotton, most seeking to optimise the accuracy of density estimates with the time required for sampling. Visual sampling for eggs and larvae is the main technique used for routine sampling, although sweep-netting or beat sheets may also be used. Helicoverpa eggs are usually concentrated in the upper parts of cotton canopies, where they are found on the upper and lower surfaces of both immature and mature leaves. Smaller numbers may be found on squares, flowers, bracts, petioles and stems. Current sampling techniques for Helicoverpa are based largely upon those originally developed for the SIRATAC computer-based pest management system now replaced by the CottonLOGIC suite of decision support systems (http://cotton.crc.org.au/ CottonLogic/HTML/About.shtml). The standard
procedure recommended by CottonLOGIC is to scout a field at least every 3 days, recording two age categories of Helicoverpa eggs and four size categories of larvae. Plants for sampling are selected using a stratified random method by which six or more groups of five adjacent plants are chosen and carefully searched. A minimum sample size of 60 plants per 100 ha field is recommended. CottonLOGIC supported three sampling procedures for Helicoverpa: numbers per metre, numbers per plant, or a binomial (presence/ absence) system rather than recording absolute numbers. The latter system saves time in sampling and recording. Up until the plant starts to produce fruit (squaring), the whole plant is sampled, but thereafter sampling is limited to the upper terminal portion of the
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Fig. 3.25. Fully grown larva of H. armigera feeding on a cotton boll. (CSIRO: C. Mares)
plant, where the majority of Helicoverpa eggs and larvae are to be found. Both these components (binomial sampling and sampling terminals only) save considerable time. Binomial sampling assumes a consistent relationship between the proportion of infested samples (pi) and the mean density of insects (m). CottonLOGIC uses a series of equations to convert the proportion of plants (or terminals) infested with Helicoverpa to actual densities. Action level (cotton): Action thresholds have been well-defined for Helicoverpa eggs and larvae in cotton. Pest management in cotton is comprehensively described in the IPM Guidelines for Australian cotton (Deutscher et al. 2005), and in the Australian Pest Management Guide (Johnson and Farrell 2004) is based on regular structured sampling at approximately 3 day intervals. Thresholds are available for each life stage and for differing stages of crop development. In seedling cotton up to flowering the threshold is defined as two larvae per metre row. After flowering, an egg threshold of five brown eggs per metre is also utilised. Thresholds are higher for Bt cotton varieties, where no egg threshold is defined and larvae must be present above threshold for two consecutive samples. Chemical control (cotton): A number of chemical pesticides are registered for H. punctigera control in Australia. Although the larvae feed in concealed locations in many crops, well-timed sprays applied at the time of egg hatch or when larvae are relatively small (first to third instar) are most effective in limiting damage. Although both H. armigera and H. punctigera have been subject to intense selection pressure by
pesticides in cotton, H. punctigera has not developed sustained resistance to any chemical (Gunning and Easton 1994). This is in contrast to H. armigera and is largely due to ecological factors. H. punctigera is highly mobile, has an extensive geographic distribution and occurs on many native and introduced uncultivated hosts, in addition to its many crop hosts. Thus, this species maintains adequate populations in unsprayed refugia to dilute selection acting in the cropping areas. The susceptibility of H. punctigera to pesticides means that single chemicals can often be effective providing control of even large larvae, in contrast with H. armigera which often requires mixtures of two or three chemicals to provide control of early instars. Cultural control: Many agronomic practices have been suggested for Helicoverpa management in various cropping systems including: manipulation of crop planting dates, stubble cultivation and destruction of crop residues, the use of closed seasons, destruction or manipulation of alternate hosts, diversionary hosts or trap crops. However, none of these are widely used for management of H. punctigera. The widespread recommendation to use cultivation for control of Helicoverpa pupae in cotton in Australia (Fitt and Daly 1990) has most impact on H. armigera, since it is the predominant species in overwintering populations. H. punctigera is not present in most crops in late autumn and does not overwinter in significant numbers in cropping regions. During winter, most of the H. punctigera populations may well be distributed throughout inland Australia where patches of ephemeral host plants are located. Host-plant resistance (cotton): Many crop plants display characters which can be exploited by plant breeders to reduce their attractiveness to ovipositing adults or suitability for Helicoverpa larvae. These include morphological characters such as glabrousness, nectariless and okra leaf in cotton, and biochemical factors such as increased levels of terpenoids and tannins, again in cotton (Fitt 1987). Although some progress has been made with natural host-plant resistance characters, there are no crops in which these provide adequate protection. The introduction of transgenic crops provides examples of engineered host-plant resistance. Transgenic cotton varieties which express the insecticidal protein genes 95
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(Cry IAc, Cry 2Ab) from Bacillus thuringiensis are commercially available in Australia (Fitt 2004). These provide excellent protection against both Helicoverpa species and have resulted in significant reductions of up to 90% in the use of pesticide during the growing seasons. Conservation of natural enemies: Cotton fields typically harbour a rich diversity of arthropods. In Australia up to 450 different species have been recorded in unsprayed fields and many of these are beneficials. It is striking that the key beneficial groups in cotton are similar in many parts of the world (Hearn and Fitt 1992), but their impacts and value have often proven difficult to demonstrate. While predators and parasites are important components of IPM systems there are often severe limitations in their capacity to control some pests, particularly the heliothine moths. An important area of research, beyond simply minimising the use of disruptive chemicals, has been to identify means to conserve, augment or manipulate beneficial populations. Conservation of natural enemies requires considerable ecological understanding of their seasonal phenology, habitat and prey requirements. The majority of predators are generalists, able to sustain populations on a diversity of prey types. Predator abundance can be readily monitored and estimates of abundance utilised in decision making through a predator:prey ratio which indicates when predators are sufficiently abundant to have impact (Mensah 2002a,b). Mechanisms to encourage beneficial insects include the use of nursery crops such as perennial lucerne crops that are attractive to many insects and offer a permanent habitat onfarm, so providing some buffer against the unpredictability of natural populations. Integrated pest management: Over the last 5 years adoption of integrated pest management (IPM) approaches has expanded dramatically in Australian cotton production in some other crops where ‘soft option’ pesticides are now widely available. A set of IPM guidelines is now freely available on the Australian Cotton CRC website at http://cotton.pi.csiro.au/Publicat/ Pest/, together with a set of 12 supporting documents, on-line insect identification aids and the computer-based decision support aid CottonLOGIC (Hearn and Bange 2002). The IPM
guidelines and CottonLOGIC provide practical approaches for producers and consultants and reflect the current state and Australian IPM research (Wilson et al. 2005), which emphasises the integration of plant compensation, host-plant resistance, beneficial insects, habitat complexity, and use of selective pesticides.
B O L L M AT U R AT I O N S TAG E
Locusts and grasshoppers N (Main entry in Chapter 18 Locusts and grasshoppers of
pastures and rangelands.)
Pest status on cotton: Minor, widespread, irregular. The most common species of locust throughout northern NSW and south-west Qld are the Australian plague locust, Chortoicetes terminifera. This insect will not generally damage cotton, although some light damage has been observed on field margins as swarms move through and ‘test feed’. The spur-throated locust, Austracris guttulosa, if present in large swarms, can cause damage to cotton. Migratory locust, Locusta migratoria, is mainly a pest in the Central Highlands of Qld where outbreaks are frequent and often severe, however, they will not damage cotton, as they are primarily grass feeders, attacking pastures and crops such as sugarcane, sorghum, wheat, barley, oats and maize.
Pale cotton stainer Dysdercus sidae Montrouzier Hemiptera: Pyrrhocoridae Pest status on cotton: Minor, restricted, irregular. The pale cotton stainer is suppressed in cotton crops which rely on insecticides for pest control, but reduced insecticide use in Bollgard II® transgenic crops and uptake of integrated pest management (IPM) may result in the cotton stainer becoming a more frequent pest. Identification: The adult bugs are about 12 mm long and light reddish brown with the ends of the forewings dark brown (Fig. 3.26b). Near the
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centre of each forewing there is a small black spot. There is a black triangular marking ahead of the centre of the back, and black markings also occur across the front of the thorax and on the head. The underside of the body is yellow– green with red and black markings. Adult females lay eggs in the soil. Host range: The pale cotton stainer is found mainly on malvaceous plants, such as hibiscus, and related plant families. Life cycle on cotton: Small nymphs (Fig. 3.26a) are gregarious and feed on seeds that can be reached by their short stylets, either in very small bolls or in open bolls. Feeding introduces fungi into the bolls that renders the seeds unviable and stains the lint a yellowish colour. The adults remain quiescent through the winter until suitable wild hosts become available in spring and summer.
Insects and Ord cotton Summer cotton was grown commercially in the Ord Irrigation Area WA, for 10 years from 1964, in spite of evidence from experimental crops that pink bollworm, rough bollworm, cluster caterpillar and native budworm were likely to cause high levels of damage. By 1974, 40 insecticide sprays applied between February and July were necessary to protect diminished yields of cotton. During this period, the hitherto uncommon cotton bollworm developed resistance to most insecticides and become the pest that finally made commercial growing uneconomic. ‘All who have viewed the Ord history will be careful to avoid the costly mistake of depending too heavily on chemicals to control pests’ (Michael and Woods 1980).
Risk period: Autumn. Monitoring: The best method to sample for the pale cotton stainer is by using a beat sheet. Damage should also be assessed regularly from boll set to boll maturity, by randomly selecting 14-day-old bolls, and squashing them into the a
palm of the hand to look for the presence of warts or stained lint. Action level: The current proposed threshold for the pale cotton stainer is three bugs per metre in beat sheet samples (Khan and Bauer 2002) or 20% of young bolls damaged (those < 14 days old).
Cotton harlequin bug Tectocoris diophthalmus (Thunberg) Hemiptera: Scutelleridae Pest status on cotton: Minor, widespread, irregular. b
Fig. 3.26. (a) cotton stainer nymph, and (b) adult of the cotton stainer (10 mm). (CSIRO: C. Mares)
Identification: The cotton harlequin bug (Fig. 3.27) is a member of the jewel bug family (Scutelleridae). The adults are 15–20 mm long and brightly coloured, females are yellow– orange with six to eight small patches of metallic green or blue scattered over the body while males are smaller and metallic green and blue with red patches or extensive areas of red. The females lay eggs in a whorl around a branch, and the female tends these until they hatch (Fig. 3.27). The nymphs are brightly coloured, often with metallic patches, and are gregarious, feeding on open bolls or very small bolls where they can reach the seeds with their short stylets. 97
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Historically, the green vegetable bug (GVB) has been suppressed by insecticides applied against Helicoverpa spp. Insect-resistant transgenic cotton (Bollgard II®) has reduced the frequency of broad-spectrum insecticides and resulted in GVB becoming a more frequent pest in cotton. Identification: See Chapter 7.
Fig. 3.27. Cotton harlequin bug nymph (17 mm). (CSIRO: P. Room)
Host range: Mostly plants of the Family Malvaceae, such as hibiscus and cotton. Risk period: In cotton this pest usually occurs in the late season. Damage: Bug feeding on the seeds in mature bolls stains the lint. It is rarely a pest, as insecticides applied against other pests control it. However, in transgenic cotton insecticide application is reduced and the pest has become more frequent. Monitoring: The threshold for cotton harlequin bug is four adult bugs per metre using a beat sheet (Khan and Bauer 2002). Damage to bolls should also be monitored regularly from boll set to boll maturity, by randomly selecting 14day-old bolls, and squashing them into the palm of the hand to look for the presence of warts or stained lint. A threshold of 20% of young bolls damaged indicates the need for control.
Green vegetable bug Nezara viridula (Linnaeus) Hemiptera: Pentatomidae N (Main entry in Chapter 7 Pulses—summer.)
Distribution: Cosmopolitan, found in Asia, Europe, Africa and South and North America where it is called the southern green stink bug. An introduced pest widely distributed in Australia. Pest status on cotton: Minor, widespread, irregular.
May be confused with: Other shield shaped bugs may resemble GVB in shape but are easily distinguished by key differences. The green stink bug, Plautia affinis (Dallas) is much smaller (about 8 mm long) and has brown forewings. The redbanded shield bug, Piezodorus hybneri (Gmelin) has a pink or yellow band across its shoulder and is smaller (about 9 mm long). The brown stink bug, Dictyotus caenosus (Westwood) is about 8 mm long and evenly dull brown. Life cycle on cotton: The female lays between 20–150 eggs in a large raft (Fig. 3.41), usually on the underside of the leaf. The cylindrical eggs are laid upright and stuck together on to the surface where they are laid. Newly laid eggs are creamy white and are about 1 mm long. They change to pinkish then orange, and hatch in 5–7 days. There are five nymphal instars, each taking about 5 days to develop to the next stage, and the life cycle takes between 5 and 6 weeks in summer. The adults live for 10–15 weeks. The GVB overwinters as adults, hibernating (in regions with mild winters) or diapausing (in regions with severe winters) under the bark of trees, in crevices, in stubble corn stalks or similar refuges. During August to September, with the rising temperature, numbers increase with large recruitment during October to midNovember on wild hosts such as wild turnip, variegated thistle Silybum marianum, marshmallow, wild castor oil amaranths of Amaranthus spp., and some field crops. As dry weather begins during November to December, these wild hosts dry and small numbers of GVB move onto cotton if no other attractive hosts such as soybean or mung bean are available. Risk period: Young boll stage. Bolls older than 20 days usually do not suffer significant damage. Damage: GVB adults and nymphs cause similar damage symptoms but later stages are more damaging. First stages usually do not feed and cluster around the egg remnants. From the second stage they start to feed and from the
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third stage they start to disperse. Adults and fourth- and fifth-stage nymphs cause almost similar amounts of damage. Third-stage nymphs cause half of the damage caused by the adults. If GVB populations are left unchecked, crop losses could exceed 50%. GVB feeding on developing fruit (bolls) causes black external feeding spots on the boll wall and internal wart-like growths or lint staining. Internal inspections can be a better guide to the extent of the damage according to the age of a boll. GVB prefer young bolls (< 12 days old) and cause shedding of bolls less than 10 days old. Damage to older bolls reduces the growth of the seeds resulting in segments of the bolls (locks), which are small and have stained lint. Plants can compensate for some loss because they produce more bolls than they can normally retain. Monitoring: Crops should be checked for GVB from boll set to boll maturity using a beat sheet. Beat sheets should be long enough (at least 1.5 m) so that the distant end of the sheet can be draped over plants in the adjoining rows to catch adults flying through these plants. GVB can be monitored with visual checking, but the efficiency of this method is almost half that of the beat sheet count. The distribution of GVB is very patchy; therefore, thorough inspections throughout a crop are necessary. Early to mid morning is the best time to inspect for GVB in the field. During this time they move to the top of the crop to bask in the sun, making location easier. Damage should also be assessed regularly, by randomly selecting 14-day-old bolls and squashing them into the palm of the hand to look for the presence of warts or stained lint. Action level: The threshold for control is 0.5 adults per metre in a visual check or one per metre using a beat sheet or 20% boll damage. Instars 4 and 5 and adults are regarded as equivalent, and instar 3 is 0.5 of instars 4 and 5 and adults. A cluster of first or second instars, clumped around the egg remnants, is equivalent to one later instar or adult (Johnson and Farrell 2004). Boll damage can be determined by sampling 12–15-day-old bolls and inspecting for internal warty growths or stained lint. Control of the GVB needs to be based on both numbers present and bolls damaged. Control: Chemical control should be applied if insect numbers and damage reaches the
threshold level. Selection of a pesticide should aim to have minimum impact on beneficial insects. Reduced rates (one-third to half of the recommended rate) of certain chemicals plus salt (NaCl; 5–10 g per litre of water) is as effective as the full rate of insecticide, while being less disruptive to beneficial insects. At the late stage of the crop when maintaining an IPM strategy is of less concern, broad-spectrum chemicals such as synthetic pyrethroids or organophosphates may be appropriate. Chemical control may be cost-effective but beneficial arthropods should be conserved by using the most selective insecticide. Natural enemies: An important natural enemy of the GVB is the egg parasitoid Trissolcus basalis (Wollaston) Hymenoptera: Scelionidae (Fig. 7.53). The parasitised eggs appear black and not the orange colour of healthy eggs. Another important natural enemy is the tachinid fly Trichopoda giacomellii (Blanchard) (Fig. 7.49), which attacks fourth- and fifth-instar nymphs and adults of GVB, attaching small white eggs predominantly on the thorax and pronotum (just behind the head). The parasitoid may cause death of some nymphs, while egg production of females that survive the parasitoid is suppressed.
Green stink bug Plautia affinis (Dallas) Hemiptera: Pentatomidae Pest status on cotton: Minor, restricted, irregular. The green stink bug looks similar but smaller than the green vegetable bug (Fig. 7.50). It has rarely been a pest of cotton until recently, when it has been recorded causing damage in crops that receive little insecticide spray and are close to areas of weeds or native vegetation, such as near riverbanks or billabongs. Adults of the green stink bug migrate from the weeds into nearby cotton crops. A number of parasitoids control this pest in weeds but not in cotton crops, possibly because the parasitoids have not moved into cotton or because insecticides applied against other pests have killed them. Green stink bug can be sampled using beat sheets. Damage should also be monitored regularly from boll set to 99
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boll maturity, by randomly selecting 14-dayold bolls and squashing them into the palm of the hand to look for the presence of warts or stained lint. Currently, the threshold for the green stink bug is three per metre in a beat sheet sample with 20% damage to small bolls (14 days old).
Redbanded shield bug Piezodorus hybneri (Gmelin) Hemiptera: Pentatomidae N (Main entry in Chapter 7 Pulses—summer.)
Pest status on cotton: Minor, widespread, irregular. Identification and life history: See Chapter 7. The redbanded shield bug (Fig. 7.60) is found in most cotton regions, although it is generally more abundant in the northern areas such as central Qld, the Ord river irrigation area (WA) and Katherine (NT). The bug breeds in cotton where nymphs are often found. Damage to young cotton bolls (fruit) caused by this pest is visible as external feeding marks, which appear as small dark puncture marks, internal wart like growths on the inside of the boll coat and/or lint staining. Thorough inspections are important as redbanded shield bug infestations tend to be patchy. Bugs are most visible in the mornings. Sample using a beat sheet. Damage should also be assessed regularly from boll set to boll maturity, by randomly selecting 14-day-old bolls and squashing them into the palm of the hand to look for the presence of warts or stained lint. Currently, the threshold for the redbanded shield bug is three per metre with 20% damage to small bolls (14 days old).
The pink bollworm is a major cotton pest in other cotton-producing countries, but in Australia it occurs only in the NT and north of WA. The pinkspotted bollworm (Fig. 3.28) occurs in coastal and central Qld, and will attack cotton as well as its primary hosts. Host range: Hosts of pinkspotted bollworm include cotton trees and broadleaf bottle tree, Brachychiton australis. Life cycle on cotton: Small, inconspicuous eggs of both species hatch into larvae that tunnel into large squares, flowers or bolls, usually completing development in the one structure. Mature larvae pupate in bolls, stems or surface trash. The whole life cycle takes about 6 weeks, but usually only one generation of pinkspotted bollworm of economic importance occurs at Biloela/Theodore (Qld) during late January–March. As far as is known, pinkspotted bollworm does not have an overwintering diapause. However, larvae can remain alive for long periods and survive by feeding on dry cotton seed in trash. Live larvae have even been found in seed cotton in modules waiting ginning. Action level: The threshold for control for pinkspotted bollworm is 5% of bolls infested in the period from flowering to harvest. Cultural control: Pink spotted bollworm: Effective burial of cotton crop residues by late August has been found to minimise spring emergence and reduce subsequent infestations in cotton. Pink bollworm: In NT and WA cotton areas, growing cotton in the winter months (April– October) avoids major damage from pink
Pink bollworm and pinkspotted bollworm Lepidoptera: Gelechiidae Pectinophora gossypiella (Saunders) and P. scutigera (Holdaway) Pest status on cotton: Minor to major, restricted, irregular. Neither of the two species of bollworm are pests of cotton in NSW or southern Qld.
Fig. 3.28. Pinkspotted bollworm larva (16 mm). (CSIRO: C. Mares)
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Fig. 3.30. A boll damaged by tunnelling of rough bollworm larva. (CSIRO: C. Mares) Fig. 3.29 Rough bollworm larva (10 mm). (CSIRO: C. Mares)
bollworm. Current research in northern Australia is focussing on winter-production systems. Quarantine: Pink bollworm could cause serious damage if it spread to, and established in, the eastern cotton-producing regions. Pink bollworm has a diapause stage and can be spread to new areas within cotton seed. Within Australia, fumigation of seed cotton or cotton seed being transported from known pink bollworm-infested areas to non-infested areas is mandatory.
Yellow peach moth Conogethes punctiferalis (Guenée) Lepidoptera: Pyralidae Pest status on cotton: Minor, restricted, irregular. Yellow peach moth is a rare late-season pest of cotton in coastal Qld. It has not been recorded from cotton in NSW. Eggs are laid on bolls and the grey–pink larvae bore into bolls and sometimes stems. Small larvae resemble those of the pinkspotted bollworm but produce masses of webbing and excreta at the entrances to their tunnels. The moth is a bright yellow–orange dotted with black spots and has a wingspan of about 25 mm (Fig. 4.9). The wide range of host plants includes sorghum and maize.
Rough bollworm Earias huegeliana Gaede Lepidoptera: Noctuidae Pest Status on cotton: Minor, widespread, irregular. Rough bollworm is the larva of a moth (Fig. 3.29) which occurs widely on malvaceous plants including cotton in Australia and the Pacific Islands. The weed, bladder ketmia is the major host of rough bollworm. Upsurges of rough bollworm in cotton are frequently associated with the maturation of wild hosts. Damage is caused directly to bolls by larval tunnelling (Fig. 3.30), although larvae can also be found in squares. Larvae may also tunnel down the mainstem causing destruction of the primary growing point at any growth stage including seedlings. Damage may retard growth and reduce yields, although this pest rarely exceeds threshold. The recommended threshold in cotton is two larvae per metre from planting to flowering, then three larvae per metre or 3% of bolls damaged for the rest of the season. A range of products is registered for control including a selective insect growth regulator. This pest is generally well-controlled by the transgenic Bollgard II® cotton. One other species is of minor importance on cotton in northern Australia: Earias vittella (Fabricius), the northern rough bollworm. It is widely distributed throughout Asia and is an important pest of cotton in India.
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BENEFICIALS R.K. Mensah and B.A. Pyke
Adult green lacewing (K. Power)
BENEFICIAL ORGANISMS spiders
predatory bugs (true bugs)
lacewings beetles
common parasitoids of cotton pests
PAGE jumping spiders lynx spiders orbweavers wolf spiders assassin bugs bigeyed bug brown smudge bug damsel bug glossy shield bug pirate bug predatory shield bug green lacewing brown lacewing green carab beetle green soldier beetle ladybirds red and blue beetle egg parasitoids larval parasitoid
Generalist predators are particularly important in cotton production because they can survive on alternative food if pest species are in short supply. Predators consume their prey in a short time compared with parasitoids, which kill their prey over a longer period. Sucking predators typically have hollow mouthparts which enable them to inject enzymes into their prey and then suck out their fluids. Chewing predators use their mandibles to chew and consume their prey.
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In contrast to predators, adult parasitoids do not feed on their host but deposit their eggs so that their offspring can use the host as food during development. Parasitoids complete their development on a single host which is usually killed either when the parasitoid larva approaches completion of its development or the adult parasitoid emerges from the host. Since parasitoids develop within and at the expense of their host, they cannot be larger than their host.
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Adult parasitoids may lay one or more eggs either externally or internally, in the egg of the host or in or on the larva, pupa or adult of the host. Depending on the stage of the host that is attacked by the parasitoid, development must be completed before the end of the host development stage. An exception can be the larval stage where parasitoid development is sometimes not completed until the host’s pupal stage.
Major beneficial insects in cotton The abundance of beneficial insects in cottonfarming systems is affected by food availability, mating partners, overwintering sites, shelter, climatic conditions and pesticide sprays (Mensah 2002a). In Australia, some 123 species of predators have been recorded in cotton-farming systems. Almost two-thirds of these are insects. There are 80 predatory insect species in 10 orders, followed by spiders (41 species in 11 orders), one mite and one species of collembola (springtail). Most major predators are listed in Table 3.3. Most of the parasitoids recorded in cotton crops are parasitoids of Helicoverpa spp. The major parasitoids are Trichogramma spp., Telenomus spp. and Microplitis demolitor. Trichogramma and Telenomus spp. attack eggs but M. demolitor prefer to lay their eggs in small Helicoverpa larvae.
Fig. 3.31. Jumping spider, 8 mm long, on cotton leaf. (W. Sterling)
Lynx spiders Oxyopes spp. Araneida: Oxyopidae Lynx spiders (Fig. 3.32) are active hunters and are usually seen in the foliage of the cotton plants. These spiders do not make a web but rely on their good eyesight to locate and capture prey. They have numerous spines on their legs and their abdomen is narrow and pointed. The young spiders or their offspring disperse by ‘ballooning’ from tall plants where the silk is played out into the breeze and is carried away. Lynx spiders feed on small Helicoverpa larvae and other insects.
Orbweavers Common key predators on cotton S PI D E R S Numerous species of spiders occur in cotton fields throughout the season, but only four of the most commonly found genera are important. Spiders are generalist predators and do not discriminate between pests and other beneficial insects as food source.
Araneus spp. Araneida: Araneidae Orbweavers (Fig. 3.33) build circular orb webs in the foliage of cotton plants or between cotton plants to trap their prey. The webs are easily seen when the cotton plants are large and before the canopy closes over the furrows. The behaviour of these spiders in capturing prey is to wait under nearby leaves for flying insects to become trapped in their webs before attacking and capturing them. They capture and eat Helicoverpa moths, and often remains of moths
Jumping spiders Araneida: Salticidae Jumping spiders (Fig. 3.31) have two large, forward-facing eyes which gives them good binocular vision to detect and locate prey. The spiders usually rest during the night but actively forage and search plants during the day for prey. They capture their prey by jumping on them. The legs are relatively short except the first pair. Most species found in cotton are dark or grey in colour.
Fig. 3.32. Lynx spider, 12 mm long, with egg sac on cotton leaf. (CSIRO: C. Mares) 103
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Fig. 3.33. Araneus sp. (15 mm long) in cotton. (CSIRO: C. Mares)
can be found hanging in the webs. The webs are usually rebuilt in every evening after capturing trapped prey.
Wolf spiders Lycosa spp. Araneida: Lycosidae Several species of wolf spiders occur in cotton fields. Most make a shallow burrow in the soil as a retreat and range over the ground searching for prey. The colour of the wolf spiders merges well with the soil background and this allows them to hide and capture prey and other predators. They prey on large Helicoverpa larvae. Females of these spiders carry their eggs in a grey ball attached to their abdomen. Wolf spiders occur throughout the cotton season. However, they are easily seen early in the cotton season when cotton plants are small, but when the cotton closes its canopy they are difficult to see.
PR E DATO R Y B U G S ( T R U E B U G S )
Fig. 3.34. Assassin bug, Pristhesancus sp. attacking Helicoverpa sp. larvae on cotton (size: 20 mm). (M. Shepard)
up to 300 eggs in a lifetime. The eggs are barrelshaped and laid upright in clusters or rows on leaves or stems of plants. Eggs hatch within 2 weeks and the developmental period is about 25 to 35 days. The nymphs resemble adults but do not have wings. Assassin bug nymphs and adults feed on a range of pests including lepidopteran larvae, aphids and many other insects such as ladybirds and spiders.
Bigeyed bug Geocoris lubra Kirkaldy Hemiptera: Lygaeidae The bigeyed bug (Fig. 3.35) is one of the more common key predators on Australian cotton crops (Pyke and Brown 1996; Room 1979). The insect is easily identifiable by its large eyes. They have piercing and sucking mouth parts which they use to kill prey by piercing and sucking their body fluids. They occur in cotton crops in early to mid season and numbers are highest in cotton during flowering. The numbers of bigeyed bug in cotton are reduced
Assassin bugs Pristhesancus spp. Coranus triabeatus Hemiptera: Reduvidae Assassin bugs (Fig. 3.34) range from 10–30 mm in length, and have distinct heads with prominent eyes. They have narrow heads and powerful piercing and sucking mouth parts. The front legs are enlarged to grasp prey and the hind legs are long and slender. Assassin bugs ambush prey that passes by. The adults can lay
Fig. 3.35. Bigeyed bug (3 mm long). (K. Power)
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Fig. 3.37. Adult Damsel bug (8 mm long) on cotton leaf. Fig. 3.36. Adult brown smudge bug (4 mm long).
(CSIRO: C. Mares)
(CSIRO: C. Mares)
by insecticide sprays, rainfall, temperature and high humidity. Adult females lay their eggs singly on leaves and stems of cotton plants. The eggs are grayish-white to pink and sausage-shaped. Eggs hatch within 5 to 8 days and a bright red spot develops in the eggs a few days after the eggs are laid. Both adults and nymphs are important predators of many cotton insect pests including eggs and larvae of Helicoverpa spp., aphids, mites, whiteflies and plant bugs. Bigeyed bugs feed on cotton nectar and sometimes on plant sap.
Brown smudge bug Deraeocoris signatus (Distant) Hemiptera: Miridae The brown smudge bug (Fig. 3.36) feeds on Helicoverpa spp. eggs, aphids, mites, apple dimpling bugs and small nymphs of green mirids. Both adults and nymphs are very effective predators of mites. Brown smudge bug numbers are usually high in unsprayed cotton during the mid-cotton season, which is December to early February (Pyke and Brown 1996; Room 1979).
Damsel bug Nabis kinbergii Reuter Hemiptera: Nabidae Damsel bugs, Nabis kinbergii Reuter (Fig. 3.37) are very common in cotton during the early and middle cotton season, from November to
January and sometimes February. Females lay white and cylindrical eggs that are inserted in plant tissues. The eggs hatch in 8–12 days and nymphs develop within 3–4 weeks. Damsel bugs occur in cotton crops in all cottongrowing areas in Australia (Pyke and Brown 1996; Room 1979). Damsel bugs are slender, dull tan to grey with long antennae and legs, and prominent eyes. Nymphs resemble the adults but have no wings. Damsel bug adults insert their eggs in the cotton plant tissues. Both nymph and adult are predators of mites, aphids, lepidopteran larvae, mirids, white flies and occasionally on other predators such as lacewing larvae, pirate bugs and bigeyed bugs. In cotton fields, damsel bugs feed on Helicoverpa spp. eggs, neonate (first to third instar) and medium and large (fourth to sixth instar) larvae. Both nymphs and adults use their piercing and sucking mouthparts to suck fluid from eggs and larvae or nymphs of insects. Damsel bugs are usually found in cotton crops infested with caterpillar eggs and larvae as well as mites. Adult damsel bugs can also be found in winter on weeds and crops during winter and also in perennial crops such as lucerne.
Glossy shield bug Cermatulus nasalis (Westwood) Hemiptera: Pentatomidae The glossy shield bug (Fig. 3.38) is found in cotton mostly in the mid to late cotton season, from mid January to March–April. The glossy shield bug lays black eggs in a raft of 50 or more on a plant (Pyke and Brown 1996; Room 1979). 10 5
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a
b Fig. 3.38. Glossy shield bug nymph (12 mm long) sucking out a cluster caterpillar. (DPI&F Qld: J. Wessels)
The eggs have short, white spines around the rim. The nymphs that hatch from the eggs are dark red and brown. The early instar nymphs are bright red. The development of glossy shield bug from egg to adult takes about 3 weeks. Adults and older nymphs feed on larvae of caterpillars, particularly Helicoverpa spp. High numbers of glossy shield bug adults and nymphs are found during midsummer or late in the cotton season in unsprayed cotton fields or fields that have received few insecticide sprays against Helicoverpa spp. In early summer, glossy shield bugs can be found in lucerne, linseed and any summer crops infested with caterpillar larvae.
Pirate bug Orius spp. Hemiptera: Anthocoridae The pirate bug moves into cotton early in the cotton season to feed on thrips and mites. They insert their eggs in leaves and other soft plant tissues. The eggs hatch after about 3 days depending on temperature. The nymphs become adults within 20 days and the adults may live for about 3 weeks. The adult pirate bugs are very small, long, flat and oval-shaped (Pyke and Brown 1996; Room 1979). They are black with a white pattern at the back and have a prominent, forward-projecting beak. Adult and nymphs of pirate bugs have piercing and sucking mouth parts. They feed on thrips, mites, aphids, plant bugs, whiteflies, and eggs and larvae of Helicoverpa spp., loopers and armyworms. In cotton, pirate bugs are very effective egg predators and also predators of thrips, and may also feed on pollen.
Fig. 3.39. (a) Predatory shield bug nymph (CSIRO: C. Mares) , and (b) adult predatory shield bug (12 mm long) on cotton leaf (CSIRO: C. Mares) .
Predatory shield bug Ochelia schellenbergii (Guérin-Méneville) Hemiptera: Pentatomidae Predatory shield bugs (Fig. 3.39) are found all year round on weeds and other crops including cotton infested with caterpillars. Adults and nymphs are usually most abundant in cotton from December to February. The predatory shield bug adult can lay about 200 eggs in its lifetime. Eggs of the predatory shield bug have long white spines around the rim and are laid in rafts usually in multiples of 14 on cotton plants. The developmental period from eggs to adult takes about 3 weeks and the adult can live for 4–8 weeks. Newly hatched nymphs remain clustered around the egg mass before dispersing. The nymphs are dark red and brown, and young nymphs feed on plant sap. Adults feed on all stages of Helicoverpa spp. larvae. Both adults and nymphs are killed by insecticide sprays.
L AC E W I N G S
Green lacewing Chrysopa spp. Neuroptera: Chrysopidae Female green lacewings lay stalked eggs which are white and oval (Fig. 3.40) and hatch within 6
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a
b
Fig. 3.41. Larva of a brown lacewing (10 mm long) preying on aphids. (Graphic Science: © Denis Crawford)
Brown lacewing Fig. 3.40. (a) Green lacewing eggs are laid on stalks, and (b) a fully grown green lacewing larva sucks bodily fluids from its prey and carries residual corpses on its back. The larva is a voracious predator, but the adult is not predatory. (Graphic Science: © Denis Crawford)
days. The larvae are pinkish-brown with welldeveloped legs and jaws. The jaws are used to suck fluid from prey. Larvae are very mobile and cannibalistic, and can grow to 6–8 mm (Fig. 3.40). They are active predators of Helicoverpa eggs, aphids and other soft-bodied insects The larvae have three instars and the duration of the instar stage is about 2–3 weeks. The pupae are round, silky cocoons and are usually attached to the underside of the leaves. The pupation period takes about 2 weeks. The adult lacewing feeds only on nectar and aphid honeydew. Adult green lacewings are pale to bright green, about 12–20 mm long with long antennae. The wings are large and transparent and are usually held upright over the abdomen. The body is very fragile and the species in cotton has prominent golden eyes. The adults are active fliers during the morning, evening and night. They occur in cotton during spring and summer and the female lays eggs on a long flexible stalk which she attaches to the plant. Adults are not predaceous. The pre-oviposition period is about 6 days and the female lays over 200 eggs (Fig. 3.40). The green lacewings have two to three generations in a year.
Micromus tasmaniae (Walker) Neuroptera: Hemerobiidae The brown lacewing (Fig. 3.41) has a life cycle similar to the green lacewing but both adult and larvae of brown lacewings are active general predators of aphids, eggs and other soft-bodied insects. Adults are brown, with wings and body covered in fine hairs. They are about half the size of green lacewings and are more secretive in their habits. The adults fly during the evenings or night. Female brown lacewings deposit hundreds of small, cream, oval eggs on the underside of cotton leaves in the field. Unlike green lacewings, the brown lacewing eggs have no stalks. The eggs become pink or purple within a few days. The larvae are reddish-brown and cream, and similar in appearance and habits to the green lacewing larvae. The larval stage lasts for about 1–3 weeks depending on temperature. Brown lacewing larvae have dark legs, antennae and mouthparts. The pupa is cream and is found in protected places in the cotton foliage.
BEETLES
Green carab beetle Calosoma schayeri Erichson Coleoptera: Carabidae 107
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Fig. 3.43. Green soldier beetle adult (10 mm long). (K. Power)
Fig. 3.42. Green carab beetle adult (24 mm long). (SARDI: G.J. Baker)
Carab beetles (Fig. 3.42) are predators, dark in colour and shiny with long slender legs and antennae. Adult green carab beetles occur in cotton fields during the flowering, boll setting and maturation stages. The adults hunt caterpillars and other slow-moving insects. They fly into cotton fields in the night and respond positively to light. The carab beetle runs very quickly and is very active at night. The larvae are worm-like but have a well-developed legs and jaws. Both adult and larvae are very active on the soil surface and the adults can climb into the cotton canopy in search of prey. Carab beetle adults hide in soil cracks and under loose soil and leaf debris during the day and hunt for prey at night. They lay their eggs in or on the soil. The larvae of the green carab beetle are also predaceous but have not been found in cotton (see Pyke and Brown 1996; Room 1979). They are also common on inland rangelands following good seasons where they prey on noctuid larvae, including Helicoverpa punctigera.
Green soldier beetle Chauliognathus pulchellus (W. S. Macleay) Coleoptera: Cantharidae Green soldier beetles (Fig. 3.43) occur in large swarms in eucalypt plantations where they are the main predators of eucalyptus leaf beetles, Chrysophtharta bimaculata (Olivier). Adults of green soldier beetle are usually found in swarms
during the late cotton season. Some swarms seem to be mating aggregations, while others seem to be mass dispersal flights. The numbers of these insects are usually very low in cotton but they prey on insect eggs, lepidopteran larvae and also slow-moving insects such as aphids. Adults respond positively to sugar solutions and honeydew of aphids. The larvae of green soldier beetles are predaceous, although they have not been found in cotton fields.
Ladybirds (Coccinellidae) Many species of ladybirds occur on cotton crops across the cotton-growing regions in Australia. The most common are the common spotted ladybird Harmonia conformis (Boisduval) (Fig. 3.45), mite-eating ladybirds Strethorus spp. (Fig. 15.54), striped ladybird Micraspsis frenata (Erichson) (Fig. 10.1), threebanded ladybird Harmonia octomaculata (Fabricius) (Fig. 3.47), transverse ladybird Coccinella transversalis Fabricius (Fig. 3.47), white collared ladybird, Hippodamia variegata Goeze (Fig. 10.1), variable ladybird Coelophora inaequalis (Fabricius), mealybug ladybirds, Cryptolaemus montrouzieri Mulsant (Fig. 3.48), and two-spotted ladybird Diomus notescens (Blackburn) (Fig. 3.49). Adult ladybirds are small, round to oval and dome-shaped. Adult females lay their eggs in groups of five to 30 on the plant, mostly near prey such as aphids (Fig. 3.44). A female ladybird may lay from 200 to more than 1000 eggs over a 1–2 month period, commencing from spring to summer. Eggs hatch within 4 days and larvae (Fig. 3.44b) feed for 2–3 weeks before turning into pupae. The larvae forage on
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a
b
Fig. 3.46. Adult common spotted ladybird shelter in protected refuges. (SARDI: P.T. Bailey)
plants to feed on the eggs, larvae/nymphs and adults of cotton pests. When the larvae are fully grown, they pupate on the plant where they emerge as adults after about a week, and have pre-oviposition period of 10 days. Ladybirds overwinter as adults often as aggregations in holes and bark of trees or under leaf litter, under rocks or protected places (Fig. 3.46). Some ladybirds change from bright yellow to a dark colour during winter periods. Ladybirds are killed by chemical sprays, rainfall, high temperatures over 35oC and high relative a
Fig. 3.44. (a) Ladybird eggs (1.5 mm long) are often laid among prey, and (b) the larva of the common spotted ladybird (12 mm long). (SARDI: G.J. Baker)
b
Fig. 3.45. Adult striped cocinella. (CSIRO: C. Mares)
Fig. 3.47. (a) Threebanded ladybird, and (b) transverse ladybird (4 mm long). (K. Power)
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Fig. 3.48. Mealybug ladybirds, Cryptolaemus montrouzieri Mulsant (adult 5 mm long) are effective predators of mealybugs. The larva (foreground) has a similar appearance to its prey. (SARDI: P.T. Bailey)
humidity over 80%. They also have their own natural enemies. Ladybirds are important predators in cotton and, depending on species, eat aphids, mites, Helicoverpa spp. eggs and larvae, whitefly and jassids. Two-spotted ladybirds (Fig. 3.49) and the mite-eating ladybirds are common predators of two-spotted mites and their eggs. Ladybirds are commonly found in cotton farms during the early cotton season and when chemicals have not been used. Furrow irrigation creates artificial humidity, which reduces ladybird abundance. Cultivation of weeds when the soil is dry produces dusts and disturbs cotton plants, forcing adults to fly away from the plant. Most common ladybirds are found in cotton growing areas in NSW and Qld (Pyke and Brown 1996; Room 1979).
Red and blue beetle
The red and blue beetle is an important predator on cotton crops and occurs throughout the cotton season. The red and blue beetle eggs, larvae and pupal stages occur in the soil. The red and blue beetles lay their eggs in clusters on soil debris. The larvae feed on small worms and other soil organisms. The adults feed on Helicoverpa eggs and sometimes very small and small (first to third instar) larvae and other insects. Most adults can also be found aggregating on pollen and flowers. Adults are found in the cotton canopy and stem in the mornings and after sunset. In hot afternoons, they may shelter in soil cracks under the cotton plants and may thus sometimes escape insecticide spray. The species overwinter as adults. Rainfall, very high humidity, furrow irrigation and soil cultivation reduce numbers (Stanley 1997).
C O M M O N PA R A S I TO I D S O F C OT TO N PE S T S
Egg parasitoids Trichogramma spp. (Hymenoptera: Trichogrammatidae) and Telenomus sp. (Scelionidae) Trichogramma spp. are egg parasitoids of Helicoverpa spp. and other Lepidoptera pests (Figs 3.50. 3.51). A number of species emerge from eggs of moths, the most common species in cotton is Trichogramma pretiosum (Fig. 3.50) but T. carverae is also common (Fig. 3.51). Trichogramma spp. adults are minute, mostly < 0.5 mm long (Fig. 3.51). The female wasp lays
Dicranolauis bellulus (Guerin-Meneville) Coleoptera: Melyridae
Fig. 3.49. The minute two-spotted ladybird Diomus notescens (3 mm long). (CSIRO: C. Mares)
Fig. 3.50. Female Trichogramma pretiosum. (DPI&F Qld: B. Scholz)
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Percentage parasitism is calculated as: total number of unhatched eggs in the egg tray/ total eggs in the egg tray × 100
Inoculative releases of Trichogramma spp.
Fig. 3.51. Trichogramma carverae Oatman & Pinto, adult (size: 0.4 mm) seeking to lay eggs on an egg mass of light brown apple moth. (SARDI: G. Caon)
one or more eggs in an egg of the host insect, and one or several parasitoids may develop. The eggs turn black after 3 days and one or more parasitic wasps may emerge from parasitised eggs 8–10 days after parasitisation. The adult female T. pretiosum can parasitise about 58 Helicoverpa eggs in her lifetime. Egg parasitoids, including Telenomus sp., are more abundant during middle to late cotton season (i.e. January to March). In Australia, population levels are much higher in the northern cotton areas compared with the southern ones (Scholz and Parker 2004).
Sampling parasitoids and estimation of Trichogramma parasitism Helicoverpa egg parasitism is measured by collecting Helicoverpa spp. eggs that are brown (> 2 days old) rather than younger white eggs (< 2 days old). Twenty eggs should be collected from leaves or squares of different cotton plants in the field (Scholz and Parker 2004) and placed individually in a multi-celled egg tray to avoid cannibalism. The egg trays should be securely sealed with sticky tape or food wrapping plastic film to restrict larval movements between trays and stored at about 24oC. Numbers of black parasitised eggs are counted within 2–3 days after egg collection when unparasitised Helicoverpa brown eggs have hatched.
In Australia, Trichogramma pretiosum are used extensively as a biocide for control of Helicoverpa spp. in cotton crops. The wasps are purchased commercially by growers and pest managers and released into cotton crops to kill Helicoverpa eggs. T. pretiosum are released from small cardboard capsules which can hold 1000 wasps and are stapled onto cotton leaves prior to release of the wasps (Scholtz and Parker 2004). Inoculative releases are used to kick-start Trichogramma field populations, by two releases of about 30 000 wasps per hectare 1 week apart in cotton or in adjacent crops, e.g. sorghum or maize crops near commercial cotton fields. The impact of inoculative releases may not be noticed until the wasps have completed one generation in the field, about 10 days after release (Scholz and Parker 2004).
Larval parasitoid Microplitis demolitor (Wilkinson) Hymenoptera: Braconidae M. demolitor is a common larval parasitoid of Helicoverpa in cotton and grain crops. It is common during mid to late cotton season. The adult lays its egg in a small Helicoverpa larva. M. demolitor larvae feed and develop inside its host after hatching. The host stops feeding after a few days and is eventually killed by the parasitoid larvae. Host larvae are usually killed before inflicting significant damage. After the death or during the moribund stage of the host, the fully developed M. demolitor larva eats its way out of the dead Helicoverpa larvae and forms a small cocoon beside it. Prior to this stage, the only way to determine if a Helicoverpa is parasitised is to open up the third- or fourth-stage larvae and examine the contents for a maggot-like Microplitis larvae (Fig. 3.52).
Methods for sampling and conserving major beneficial insects in cotton Sampling predators Visual, suction, beat sheet or sweep net can be used to sample beneficial insects and spiders and 111
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Use of parasitoids in pest management decisions In some Australian cotton crops, Trichogramma spp. are effective in reducing the number of viable Helicoverpa eggs. Thus, by incorporating levels of egg parasitism into the ‘predator to pest ratio’ to become ‘beneficial to pest ratio’ (Mensah and Scholz 2005), parasitism by Trichogramma spp. can be included by deducting parasitised Helicoverpa eggs from visual egg counts before calculating the ratio. Fig. 3.52. Larva of Microplitis demolitor (below) squeezed from a still-living H. armigera larva. (DPI&F Qld: J. Wessels)
of these the beat sheet method is the most effective and can be used all season round (Scholz et al. 2001). For good estimation of beneficial insect numbers, it is important that insects are sampled before noon or late in the afternoon. As the temperature rises throughout the day, most beneficial insects seek shelter in cracks in the soil or move lower down the plant, which can make them difficult to find and count.
Thus, the beneficial to pest ratio is calculated as: total number of predators/(Helicoverpa eggs per metre – [% egg parasitism]+ very small + small larvae) For example, if visual assessment of cotton crops is the following: total predators per metre = 20 number of Helicoverpa eggs per metre = 40 number of very small and small larvae per metre = 1.2 per cent egg parasitism = 50%
Use of predator to pest ratio in pest management decisions Mostly, pest control decisions are made on the basis of the pest numbers rather than the ratio of natural enemies to the pests. With the increasing reliance on beneficial insects to minimise pesticide use and the adoption of IPM worldwide, the use of both beneficial insects and pests thresholds is becoming increasingly important. There is always a question asked as to how many predators or total beneficial insects to pests are enough or required to maintain major pests such as Helicoverpa spp. below economic thresholds. An acceptable predator to pest ratio is 0.5 or higher; a ratio of less than 0.5 could result in a higher survival of Helicoverpa spp. larvae to exceed the recommended pest threshold of two larvae per metre in Australian cotton (CottonLogic 1999). The calculation of the predator to pest ratio was based on visual counts of a metre row of cotton of a total predators per metre divided by Helicoverpa spp. eggs and very small (VS) plus small (S) larvae. The calculation of the ratio excluded Helicoverpa spp. medium and large larvae since at this stage of the life cycle of the pest they were too large for the predators to capture as food.
Beneficial to pest ratio (including egg parasitism): = 20 predators ÷ (40 eggs per metre – [50% of 40 eggs = 20 eggs] + 1.2) = 20 ÷ (40–20 + 1.2) = 20 ÷ 21.2 = 0.94, which is higher than 0.50. This means the system is working well against Helicoverpa spp. and so there is no need for Helicoverpa control. In contrast, in another region where egg parasitism is low, parasitoids are unlikely to be recorded, and the ratio based on the above check would have been: predator to pest ratio = 20 ÷ 41.2 = 0.49 This means that it is necessary to apply appropriate pest management measures to manage Helicoverpa spp. without reducing beneficials.
Conserving major beneficial insects in cotton Australian cotton-production systems consist of large fields that do not offer refuge sites to beneficial insects. Lack of ecological or crop
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diversity at the cotton field level can result in reduced abundance and ability to maintain large populations of beneficial insects in cotton fields. Ecological diversity or diverse cropping systems support a greater abundance and diversity of beneficial insects. Cotton is grown as an annual crop and when the crop matures and is harvested, beneficial insects have few host plants to forage on and hunt for prey. As a result, beneficial insects relocate to new areas which may be far away from the cotton growing areas in search of food, overwintering sites and mates (Landis et al. 2000; Mensah 1999; Mensah and Khan 1997; Mensah and Sequeira 2004). Techniques that can be used to conserve and enhance beneficial insect activity, particularly predatory insects and spiders, on cotton farms include: • use of refuge or nursery crops within the cotton crop to conserve beneficial insects • use of beneficial insect food attractants (food sprays) • use of the ratio of predators and parasitoids to pests to account for beneficials when making crop management decisions • inundative releases of beneficial insects, particularly Trichogramma spp. • tolerating non-economic early season damage by allowing cotton plants to compensate for damage • use of insect-resistant cotton varieties • appropriate use of insecticides including reduced application frequency, use of insecticides of low predator toxicity and residual action and application at times least likely to target beneficials. Use of refuge or nursery crops within the cotton crop to conserve beneficial insects In Australia, the source of beneficial insects in cotton fields is often the trees, grasses, weeds, and crops that surround the farm or individual fields. These refuge areas provide overwintering sites, shelter, food and mates for beneficial insects after crops are harvested. However, the majority of beneficial insects that colonise these refuges may not return to recolonise cotton
crops. Thus, an area-wide management that protects trees, shrubs and catchment areas surrounding cotton fields is important in stabilising insect populations within adjacent cotton fields. Provision of artificial refuges within cotton farms Cotton intercropping provides refuges for beneficial insects. Intercropping cotton crops in commercial fields with lucerne strips (Fig. 3.53) or blocks used as trap and refuge crops can provide resources such as food, shelter and overwintering sites, thereby increasing the establishment, abundance and effectiveness of beneficial insects (Mensah 1999; Mensah and Sequeira 2004). Similarly, sorghum inter-planted with cotton attracts Helicoverpa armigera to lay eggs, which act as hosts to Trichogramma pretiosum. These egg parasitoids may be released into sorghum where they establish for a number of generations. Sorghum also acts as a nursery for red and blue beetles and the white collared ladybird, the most important beetle predators in southern Qld cotton crops (Scholz and Parker 2004). Supplementary food or beneficial insect attractants (food sprays) to conserve and enhance beneficial insects The application of supplementary food on commercial cotton crops enables beneficial insects, particularly predators, to be attracted, retained and conserved in the cotton crop. Use of supplementary food and attractants can either increase the total number of predators and/or pests or alter the ratio of predators to pests in favour of predators, particularly those effective
Fig. 3.53. Refuges for beneficials in cotton: intercrop lucerne strips (centre) with cotton either side. Also shown is a method of furrow irrigation. (R. Mensah)
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against Helicoverpa spp. (Mensah 1996; Mensah and Harris 1995). Beneficial insects are best established in cotton before the arrival of pests because most major pests of cotton, especially Helicoverpa spp. can rapidly migrate during the early season (Fitt 1994). The successful establishment of beneficial insects, especially predators in cotton crops, may vary in the degree of attraction, the type of food product used and the rate of growth of the crop when the food attractant was applied (Mensah 1997; Mensah and Singleton 2003). Tolerating non-economic early season damage by allowing cotton plants to compensate for damage Cotton plants have the ability to compensate for damage during the early cotton season without affecting yield or crop maturity. It is important that fruit damage and retention is monitored in addition to pests and beneficials for management purposes. Combining pest and fruit damage in regular crop assessments and using them in pest-management decisions assists in minimising insecticide use and in conserving beneficial insect populations. Use of insect-resistant cotton varieties Transgenic cotton varieties have become important pest management tools in Australian cotton production. Transgenic plants are used to manage Helicoverpa spp. and a number of other lepidopteran pests without the need to spray synthetic insecticides previously used to manage
Helicoverpa spp. and have provided increased opportunities to conserve beneficial insects. However, the Bt toxins in the transgenic cotton do not affect sucking pests such as plant bugs (e.g. mirids, aphids, whitefly) and these insects need to be managed to allow beneficial insect and spider activity. Appropriate use of insecticides Intervention with synthetic insecticides is a last resort when pests exceed thresholds. The rate, timing and method of application of synthetic insecticides can be used to minimise the impact of the insecticide against beneficial insects: • Synthetic insecticides, when used at a low rate, can be as effective as the full label rate but may be less disruptive against beneficial insects. • Timing of insecticide application to coincide with periods when beneficial insects are not foraging on the cotton crops also helps minimise the disruptive impact on beneficial insects. • Application of insecticides to the stems rather than the foliage of plants conserves beneficial insects on crop plants. • Spot treatment within fields instead of blanket sprays can also help to conserve beneficial insects. In this way, only those fields that actually require pest control are sprayed. This leaves other fields that are not being sprayed to serve as sources of beneficial insects to re-colonise the treated fields.
Sources of information Bournier, J.P. (1994). Thysanoptera. In: Insect Pests of Cotton (Matthews, G.A. and Tunstall, J.P., eds). CAB International, Wallingford, UK. pp 381–391. Common, I.F.B. (1953). The Australian species of Heliothis (Lepidoptera: Noctuidae) and their pest status. Australian Journal of Zoology 1: 319–344. Common, I.F.B. (1990). Moths of Australia. Melbourne University Press, Melbourne. CottonLogic (1999). A Compendium of Information on Insects in Cotton: Integrated Pest Management Guidelines for Australian Cotton Growers (compiled by Mensah, R.K. and Wilson, L.). Australian Cotton Co-operative Research Centre, Technology Resource Centre Press, Narrabri, New South Wales, Australia. Cullen, J.M. (1969). The reproduction and survival of Heliothis punctigera Wallengren in South Australia. PhD thesis, University of Adelaide.
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De Barro, P.J. (1995). Bemisia tabaci biotype B: a review of its biology, distribution and control. CSIRO Entomology Technical Paper No. 33, CSIRO Australia. Deutscher, S.A., Wilson, L.J. and Mensah, R.K. (2005). Integrated Pest Management Guidelines for Cotton Production Systems in Australia. Australian Cotton CRC Publication. ISBN 1921025018. Dillon, M.L., Fitt, G.P., Hamilton, J.G., and Rochester, W.A. (1996). A simulation model of winddriven dispersal of Helicoverpa moths. Ecological Modelling 86: 145–150. Ellsworth, P.C., Diehl, J.W., Dennehy, T.J. and Naranjo, S.E. (1995). (Rev. 11/2000). Sampling Sweetpotato Whiteflies in Cotton. IPM Series No. 2. Publication No. 194023. University of Arizona, College of Agriculture and Life Sciences, Cooperative Extension, Tucson, Arizona. Firempong, S. and Zalucki, M.P. (1990). Host plant preferences of populations of Helicoverpa armigera (Hubner) (Lepidoptera, Noctuidae) from different geographic locations. Australian Journal of Zoology 37: 665–673. Fitt, G.P. (1987). Ovipositional responses of Heliothis spp. to host plant variation in cotton (Gossypium hirsutum). In: Insects-Plants. Proceedings of the 6th International Symposium on Insect–Plant Relationships (Pau 1986) (Labeyrie, V., Fabres, G. and Lachaise, D., eds), Dr. W. Junk, Dordrecht. pp. 289–295. Fitt, G.P. (1989). The ecology of Heliothis species in relation to agro-ecosystems. Annual Review of Entomology 34: 17–52. Fitt, G.P. (1994). Cotton pest management: Part 3: An Australian perspective. Annual Review of Entomology 39: 543–562. Fitt, G.P. (2003). Deployment and impact of transgenic Bt cottons in Australia. In: The Economic and Environmental Impacts of Agbiotech: A Global Perspective (Kalaitzandonakes, N.G. ed.). Kluwer, New York. pp. 141–164. Fitt, G.P. (2004). Implementation and impact of transgenic Bt cottons in Australia. In: Cotton Production for the New Millenium. Proceedings of the 3rd World Cotton Research Conference, 9–13 March, 2003, Cape Town, South Africa. Agricultural Research Council—Institute for Industrial Crops, Pretoria, South Africa. pp. 371–381. Fitt, G.P. and Daly, J.C. (1990). Abundance of overwintering pupae and the spring generation of Helicoverpa spp. (Lepidoptera: Noctuidae) in northern New South Wales, Australia: consequences for pest management. Journal of Economic Entomology 83: 1827–1836. Forrester, N.W. and Wilson, A.G.L. (1988). Insect pests of cotton. NSW Agriculture AGFACT Agdex 151/620. ISSN 0725–7759. Gregg, P.C., Fitt, G.P., Zalucki, M.P. and Murray, D.A.H. (1995). Insect migration in an arid continent II. Helicoverpa spp. in Australia. In: Insect Migration: Tracking Resources Through Space and Time (Drake, V.A. and Gatehouse, A.G., eds). Cambridge University Press, New York. pp. 151–172. Gunning, R.V. and Easton C.S. (1994). The response of Helicoverpa punctigera (Wallengren) (Lepidoptera, Noctuidae) to DDT, endosulfan, deltamethrin and fenvalerate. Journal of the Australian Entomological Society 33: 61–64. Hardwick, D.F. (1965). The Corn Earworm Complex. Memoirs of the Entomological Society of Canada 40. 247 pp. Hearn, A.B. and Fitt G.P. (1992). Cotton Cropping Systems. In: Field Crop Ecosystems of the World (Pearson, C., ed.) Elsevier Press. pp. 85–142. Hearn, A.B., and Bange, M.P. (2002). SIRATAC and CottonLOGIC: persevering with DSSs in the Australian Cotton Industry. Agricultual Systems 74: 27–56.
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Herron, G.A., Edge, V.E., Wilson, L.J. and Rophail, J. (1998). Organophosphate resistance in spider mites (Acari: Tetranychidae) from cotton in Australia. Experimental and Applied Acarology 22: 17–30. Herron, G.A., Rophail, J. and Wilson, L.J. (2001). The development of bifenthrin resistance in twospotted spider mite (Acari: Tetranychidae) from Australian cotton. Experimental and Applied Acarology 25: 301–310. Herron, G.A., Rophail, J. and Wilson, L.J. (2004). Chlorfenapyr resistance in two-spotted spider mite (Acari: Tetranychidae) from Australian cotton. Experimental and Applied Acarology 34: 315–321. Hill, L. (1993). Colour in adult Helicoverpa punctigera (Wallengren) (Lepidoptera, Noctuidae) as an indicator of migratory origin. Journal of the Australian Entomological Society 32: 145–151. Johnson, A. and Farrell, T. (2004). Cotton pest management guide, 2004–05. NSW Department of Primary Industries publication. ISSN 1442–8792. Kelly, D., Wilson, L. and Parlato, D. (2002a, b). Silverleaf whitefly in Australian cotton. Australian Cotton CRC Research Review No. 12 & 13. Khan, M. and Bauer, R. (2002). Damage assessment, monitoring and action thresholds of stinkbugs in cotton. Proceedings of the 11th Australian Cotton Conference, Brisbane, August 2002. pp. 395–400. Khan, M., Hickman, M., Kelly, D., Mensah, R., Brier, H. and Wilson, L. (2004). Mirid ecology in Australian cotton. Australian Cotton Cooperative Research Centre Research Review Nos. 13 & 14. www.cotton.crc.org.au Landis, D.A., Wratten, S.D. and Gurr, G.M. (2000). Habitat management to conserve natural enemies of arthropod pests in agriculture. Annual Review of Entomology 45: 175–201. Maelzer, D.A., and Zalucki, M.P. (1999). Analysis of long-term light-trap data for Helicoverpa spp. (Lepidoptera: Noctuidae) in Australia: the effect of climate and crop host plants. Bulletin of Entomological Research 89: 455–463. Maelzer, D.A., and Zalucki, M.P. (2000). Long range forecasts of the numbers of Helicoverpa punctigera and H. armigera (Lepidoptera: Noctuidae) in Australia using the Southern Oscillation Index and the Seas Surface Temperature. Bulletin of Entomological Research 90: 133–146. Matthews, M. (1999). Heliothine Moths of Australia. CSIRO Publishing, Collingwood. Mensah, R.K. (1996). Suppression of Helicoverpa spp. oviposition by use of the natural enemy food supplement Envirofeast®. Australian Journal of Entomology 35: 323–329. Mensah, R.K. (1997). Local density responses of predatory insects of Helicoverpa spp. to a newly developed food supplement ‘Envirofeast® ’ in commercial cotton in Australia. International Journal of Pest Management 43: 221–225. Mensah, R.K. (1999). Habitat diversity: implications for the conservation and use of predatory insects of Helicoverpa spp. in cotton systems in Australia. International Journal of Pest Management 45: 91–100. Mensah, R.K. (2002a). Development of an integrated pest management programme for cotton. Part 1: Establishing and utilising natural enemies. International Journal of Pest Management 48: 87–94. Mensah, R.K. (2002b). Development of an integrated pest management programme for cotton. Part 2: Integration of a lucerne/cotton interplant system, food supplement sprays with biological and synthetic insecticides. International Journal of Pest Management 48: 95–105.
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Mensah, R.K. and Harris, W. (1995). Using Envirofeast® spray and refugia technology for cotton pest control. Australian Cotton Grower 16: 30–33. Mensah, R.K. and Khan, M. (1997). Use of Medicago sativa (L.) interplantings /trap crops in the management of the green mirid, Creontiades dilutus (Stål) in commercial cotton in Australia. International Journal of Pest Management 43: 197–202. Mensah, R.K. and Sequeira, R.V. (2004). Habitat manipulation for insect pest management in cotton cropping systems. In: Ecological Engineering for Pest Management (Gurr, G. M., Wratten, S. D. and A. Altieri, M., eds). CSIRO Publishing, Australia and CABI Publishing, UK. pp. 187–197. Mensah, R.K. and Singleton, A. (2003). Optimum timing and placement of a supplementary food spray Envirofeast® for the establishment of predatory insects of Helicoverpa spp. in cotton systems in Australia. International Journal of Pest Management 49: 163–168. Michael, P.J. and Woods, W.M. (1980). An entomological review of cotton growing in the Ord River Area of Western Australia. Technical Bulletin No. 48. Western Australian Department of Agriculture. Oertel, A., Zalucki, M.P., Maelzer, D.A., Fitt, G.P. and Sutherst, R. (1999). Size of the first spring generation of Helicoverpa punctigera (Wallengren) (Lepidoptera: Noctuidae) and winter rain in central Australia. Australian Journal of Entomology 38: 99–103. Pyke, B.A. and Brown, E.H. (1996). The Cotton Pest and Beneficial Guide. Cotton Research and Development Corporation and Centre for Tropical Pest Management publications, GOPRINT Press, Woolloongabba, Queensland, Australia. ISBN 0 7242 6633 X. Rochester, W.A., Dillon, M.L., Fitt, G.P. and Zalucki, M.P. (1996). A simulation model of the longdistance migration of Helicoverpa spp. moths. Ecological Modelling 86: 151–156. Room, P.M. (1979). Insects and Spiders of Australian Cotton Fields. NSW Department of Agriculture publications, Sydney. 69pp. Sadras, V.O. and Wilson, L.J. (1998). Recovery of cotton crops after early season damage by thrips (Thysanoptera). Crop Science 38: 399–409. Scholz, B., Cleary, A. and Lloyd, R. (2001). Sheet unbeatable for sampling predators in cotton. Australian Cotton Grower 22: 14–17. Sequeira, R., De Barro, P., Gunning, R., Wilson, L., Franzmann, B., Grundy, P. and Kelly, D. (2006). Silverleaf whitefly management guidelines for Australian cotton. Cottom Catchment Communities Cooperative Research Centre, Narrabri, NSW. http://www.cotton.crc.org.au Sholz, B. and Parker, N. (2004). Evaluations of different releases rates of Trichogramma pretiosum against Helicoverpa armigera eggs in sorghum and cotton. Proceedings 12th Australian Cotton Conference. pp. 649–656. Stanley, J. (1997). The seasonal abundance and impact of predatory arthropods on Helicoverpa spp. in Australian cotton fields. PhD thesis, University of New England, Armidale, Australia. Stanley, J.M. (1978). Competitive interactions between larvae of Heliothis armigera (Hübner) and Heliothis punctigera Wallengren (Lepidoptera: Noctuidae). PhD thesis, Australian National University, Canberra. Titmarsh, I.J., Storey, R.I. and Strickland, G.R. (1990). The species composition of Helicoverpa Hardwick (Heliothis Ochsenheimer) (Lepidoptera: Noctuidae) infestations on tobacco in far north Queensland. Journal of the Australian Entomological Society 29: 81–86.
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Twine, P.H. (1978). Effect of temperature on the development of larvae and pupae of the corn earworm, Heliothis armigera (Hübner) (Lepidoptera: Noctuidae). Queensland Journal of Agricultural and Animal Sciences 35: 23–28. Trowell, S.C., Forrester, N.W., Garsia, K.A., Lang, G.A., Bird, L.J., Hill, A.S., Skerritt, J.H. and Daly, J.C. (2000). Rapid antibody-based field test to distinguish between Helicoverpa armigera (Lepidoptera: Noctuidae) and Helicoverpa punctigera (Lepidoptera: Noctuidae). Journal of Economic Entomology 93: 878–891. Walden, K.J. (1995). Insect migration in an arid continent. III. The Australian plague locust Chortoicetes terminifera and the native budworm Helicoverpa punctigera in Western Australia. In: Insect Migration: Tracking Resources Through Space and Time (Drake, V.A. and Gatehouse, A.G., eds). Cambridge University Press, New York. pp. 173–190. Walter, G.H. and Benfield, M.D. (1994). Temporal host-plant use in 3 polyphagous Heliothinae, with special reference to Helicoverpa punctigera (Wallengren) (Noctuidae: Lepidoptera). Australian Journal of Ecology 19: 458–465. Wardhaugh, K.G., Room, P.M. and Greenup, L.R. (1980). The incidence of Heliothis armigera (Hubner) and H. punctigera Wallengren (Lepidoptera: Noctuidae) on cotton and other host plants in the Namoi Valley of New South Wales. Bulletin of Entomological Research 70: 113–31. Waterhouse, D.F. and Norris, K.R. (1989). Biological Control. Pacific Prospects Supplement 1. ACIAR, 123pp. Wilson, A.G.L. (1983). Abundance and mortality of overwintering Heliothis spp. Journal of the Australian Entomological Society 22: 191–199. Wilson, L.J. (1993). Spider mites (Acari: Tetranychidae) affect yield and fiber quality of cotton. Journal of Economic Entomology 86: 566–585. Wilson, L. (1994). Resistance of okra-leaf cotton genotypes to two-spotted spider mites (Acari: Tetranychidae). Journal of Economic Entomology 87: 1726–1735. Wilson, L. (1995). Habitats of two-spotted spider mites (Acari: Tetranychidae) during winter and spring in a cotton-producing region of Australia. Environmental Entomology 24: 332–340. Wilson, L.J. and Bauer, L.R. (1993). Species composition and seasonal abundance of thrips (Thysanoptera) on cotton in the Namoi Valley. Journal of the Australian Entomological Society 32: 187–192. Wilson, L.J., Bauer, L.R. and Lally, D.A. (1998). Effect of early season insecticide use on predators and outbreaks of spider mites (Acari: Tetranychidae) in cotton. Bulletin of Entomological Research 87: 477–488. Wilson, L.J., Bauer, L.R. and Lally, D.A. (1999). Insecticide-induced increases in aphid abundance in cotton. Australian Journal of Entomology 38: 242–243. Wilson, L.J., Bauer, L.R. and Walter, G.H. (1996). ‘Phytophagous’ thrips are facultative predators of two-spotted spider mites (Acari: Tetranychidae) on cotton in Australia. Bulletin of Entomological Research 86: 297–305. Wilson, L., Herron, G., Franzmann, B. and Heimoana, S. (2001). Cotton aphid management suggestions. Australian Cottongrower 22(4): 28–34. Wilson, L.J., Mensah, R.K. and Fitt, G.P. (2004). Implementing Integrated Pest Management in Australian Cotton. In: Insect Pest Management: Field and Protected Crops (Horowitz, A.R. and Ishaaya, I., eds). Springer, Berlin, Heidelberg, New York. pp. 97–118.
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Wilson, L.J. and Morton, R. (1993). Seasonal abundance and distribution of Tetranychus urticae, the two spotted spider mite (Acari: Tetranychidae), on cotton in Australia and implications for management. Bulletin of Entomological Research 83: 291–303. Wilson, L. and Spora, A. (2001). Aphids in cotton. Australian Cotton CRC Research Review No. 10, and Strategies to manage aphids in cotton. Australian Cotton CRC Research Review No. 11. Zalucki, M.P., Daglish, G., Firempong, S. and Twine, P. (1986). The biology and ecology of Heliothis armigera (Hubner) and Heliothis punctigera Wallengren (Lepidoptera, Noctuidae) in Australia— what do we know? Australian Journal of Zoology 34: 779–814. Zalucki, M.P., Murray, D.A.H., Gregg, P.C., Fitt, G.P., Twine, P.H. and Jones, C. (1994). The ecology of Helicoverpa armigera (Hübner) and H. punctigera (Wallengren) in the inland of Australia: larval sampling and host plant relationships during winter and spring. Australian Journal of Zoology 42: 329–346.
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4 MAIZE D.A.H. Murray N Maize, Zea mays Poaceae. Origin: Tropical America
(a) Irrigated maize crop (Pacific Seeds)
(b) A silking corn cob (Pacific Seeds)
germination growth tasselling seed filling maturity & harvest
Sep
Oct
Nov
Dec
Jan
Feb
Mar
Apr
May
Phenology of a maize crop.
PEST (major pests in bold) Germination black field earwig maize leafhopper maize thrips false wireworm
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PEST (major pests in bold)
PAGE
sugarcane wireworm cutworms Growth locusts corn aphid green vegetable bug African black beetle whitegrubs whitefringed weevil redshouldered leaf beetle swarming leaf beetles common armyworm dayfeeding armyworm Silking to tasselling corn earworm Seed-filling to maturity two-spotted mite green vegetable bug redbanded shield bug yellow peach moth
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The average area planted with Maize in Australia is 71 000 ha, of which 70% is grown in Qld, 29% in NSW and 1% in Vic. and WA. Maize is grown as a dryland and irrigated crop. Most of the maize crop is used by the domestic market for stock food (grain and silage) and processed for human consumption. Maize is a summer crop planted between September and January. In southern Qld, normal spring planting is in September/ October when soil temperatures are above 12°C. There is another opportunity to plant in December, to avoid flowering in the midsummer heat. Spring plantings are later in southern districts, and earlier in warmer coastal regions. The main factors influencing planting time are soil temperature for germination, avoiding summer heat during flowering, and reducing disease risk with late planting. Germination usually takes 5–7 days, vegetative growth about 60–70 days, tasselling and silking (flowering) 7–12 days, seed-filling 50–60 days (to physiological maturity), maturity and harvest 4.5–6 months after sowing, depending on variety and seasonal conditions.
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Black field earwig Nala lividipes (Dufor) Dermaptera: Labiduridae Distribution: Southern Europe, Asia, Africa, Hawaii and Australia. Widespread in Australia. Pest status: A major pest in Qld, minor in southern Australia, widespread but populations are greatest in areas with moisture-retentive soils. Regular occurrence in Qld maize. Identification: Adults are 15 mm long, shiny black with a flattened body and a pair of curved pincers at the end of the body. Nymphs resemble adults but are wingless and paler (Fig. 4.1). May be confused with: The common brown earwig Labidura truncata, which grows to a larger size (24 mm) and is lighter in colour than black field earwig. Host range: Wheat, sorghum, maize and sunflowers are the preferred hosts, but seedlings of most field crops are susceptible (Simpson and Robertson 1993).
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a
Fig. 4.1. Black field earwigs. The upper two are nymphal stages and lower is an adult (15 mm long). (DPI&F Qld:
b
D. Ironside)
Life cycle on maize: The black field earwig normally feeds on decaying stubble in cultivation with all stages (adults and immatures) present during warmer months. In common with a number of earwig species, female black field earwigs lay eggs in a burrow in the soil and remain to care for the eggs and nymphs. Eggs hatch in 6–7 days at 29°C. The developmental time for five nymphal instars is about 7 weeks in clay soils, longer in sandy soils. Nymphs develop into adult females or major or minor males. Longevity is about 20 weeks (Simpson 1991). Risk period: During warmer months, germinating seed and early seedling growth stages are most susceptible to attack. Irrigation or rainfall during spring–summer favours population increase. Damage: The black field earwig eats newly sown and germinating seed, and the roots of crops resulting in poor establishment (Fig. 4.2). Feeding on secondary roots may cause the plants to fall over as they get larger. Monitoring: Germinating seed baits and spade samples before sowing. Action level: Control if more than 50 earwigs in 20 germinating seed baits or, control is indicated if one earwig is found in 20 spade samples. Chemical control: Grain baits containing insecticide applied at sowing offer best protection. Insecticide seed dressings provide some protection. In-furrow sprays are not effective in protecting against dense populations.
Fig. 4.2. (a) Healthy maize seedlings (two on the left) and compared with the two plants to the right with secondary roots damaged by field earwigs; (b) feeding on prop roots by earwigs and other soil insects may result in lodging of maize plants. (DPI&F Qld: D.A.H. Murray)
Cultural control: The black field earwig is mainly a pest in areas having heavy, black soils. Earwigs prefer cultivated soils rather than undisturbed (zero til) soil. Use press wheels at sowing, which are set at 2–4 kg cm–1 width after planting rain, or 4–8 kg cm–1 in dry soil.
Maize leafhopper Cicadulina bimaculata (Evans) Hemiptera: Cicadellidae Distribution: Europe, Australia, in Qld and NSW. Pest status on maize: Minor, widespread in Qld, irregular Identification: Adults are 3 mm long, yellow or green–bright green, elongate, wedge-shaped and jump readily or move sideways when disturbed (Fig. 4.4). Nymphs are similar, but smaller, paler and wingless. 12 3
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Earwigs and wireworms as both pests and beneficials A number of earwig species are omnivorous. Their plant-feeding behaviour may result in plant damage but by their predatory behaviour they may also effectively control other crop pests. The pest status of the common black earwig changes in southern cropping areas where, together with the common brown earwig, Labidura truncata Kirby Labiduridae, it is an effective predator of a number of pests (Horne and Edward 1995, Kapuge et al. 1987) (Fig. 4.3). European earwigs, Forficula auriculata Linnaeus Forficulidae (Fig. 5.3), recently
found to be spreading in southern cropping areas, are known as both pests and beneficials in fruit orchards. True wireworms, Agrypnus variabilis, Elateridae are omnivores and may be pests of seedlings, but later in the season a
their larvae prey on pest species. During mid to late summer, wireworm larvae eat pupae of corn earworm and native budworm and appear to be important in reducing overwintering populations of these pests (Murray 1989). b
Fig. 4.3. Common brown earwig adults (24 mm long) attacking (a) a moth larva (M. Shepard) , and (b) a moth pupa (DPI&F Qld: J. Wessels) .
May be confused with: Other leafhoppers. Host range: Maize, sorghum, millet. Life cycle on maize: Multiple generations can occur on a crop but usually in low numbers.
their dark green colour and thickening of the veins on the underside of the leaves. Monitoring: Inspect crops weekly during the vegetative stage.
Risk period: The maize leafhopper is common on maize, particularly in coastal districts, during late summer. Damage: The maize leafhopper feeds by sucking sap. It also transmits wallaby ear mycoplasma (Fig. 4.5). In coastal areas, heavy infestations (more than 15 per plant) produce wallaby ear symptoms in susceptible maize hybrids. Affected plants can be recognised by
Fig. 4.4. Adult maize leafhoppers (3 mm long). (DPI&F Qld)
Fig. 4.5. Leafhopper-transmitted wallaby ear mycoplasma symptoms in a maize plant. (DPI&F Qld)
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Action level: Spray if more than 10 leafhoppers per plant (average) are found and wallaby ear symptoms are present. Chemical control: Chemical control options are available, but crops can be rapidly re-infested after spraying. Host-plant resistance: Hybrid varieties offer some resistance to wallaby ear.
Maize thrips Frankliniella williamsi Hood Thysanoptera: Thripidae Distribution: Americas, Australia. Pest status: Minor in Qld, widespread in Qld, irregular. Identification: Adults are 2 mm long with two pairs of narrow wings fringed with hairs. Immature thrips are wingless. May be confused with: A number of other species of thrips. Host range: Maize. Life cycle on maize: Multiple generations can occur on maize. Adults lay eggs in maize tissue. Risk period: During seedling and vegetative growth stages, particularly if plants are stressed. Damage: Thrips in the whorl can stop the growth of small plants, particularly if under water stress or if waterlogged. Monitoring: Inspect weekly during seedling and vegetative stages. Action level: Apply controls if plants develop yellowing in the throat and/or necrotic stripes on young leaves, and thrips can be found in the throat. Re-infestation may be rapid, so more than one spray may be required. Chemical control: Chemical control is costeffective.
False wireworms Gonocephalum spp., Pterohelaeus spp. Coleoptera: Tenebrionidae
N (Main entry in Chapter 5 Oilseeds.)
Pest status in maize: Minor, widespread, regular. Identification: See Chapter 5. May be confused with: Several species of false wireworms may occur in any particular crop, depending on locality, soil type, organic matter and tillage practices. Host range: All field crops. Life cycle on maize: Larvae feed on decaying vegetable matter and crop residues in the soil, as well as on newly germinated seed. They usually have a single generation per year. Risk period: Larvae cause most damage to germinating crops during spring while adults are most troublesome in summer. Damage: Larvae feed on germinating seed and chew on seedling roots and shoots, resulting in patchy stands. Adults chew on seedlings at or above ground level, ring-barking or completely cutting the stem. Monitoring: Germinating seed baits (GSB) and spade samples. Action level: Treatment is required if more than 25 wireworm larvae are found in 20 GSB. For adults, apply bait to the whole field if the soil insect rating (SIR) is more than 6. Chemical control: For larvae, use seed dressings or in-furrow sprays. For adults, use cracked grain baits. Cultural control: The false wireworm is mainly a pest in areas having heavy, black soils. It prefers cultivated soils rather than zero til. Use press wheels at sowing, which are set at 2–4 kg cm–1 width after planting rain or 4–8 kg per cm in dry soil.
Sugarcane wireworm, true wireworm, click beetle Agrypnus variabilis (Candèze) Coleoptera: Elateridae Distribution: Australia (and New Zealand as an accidental introduction). Pest status: Major, widespread, irregular. 125
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Identification: Eggs are ovoid, 0.6 × 0.5 mm Larvae grow to 35 mm long, are shiny and cream, yellow or tan, with three pairs of legs behind the head. Unlike false wireworms, they are soft-bodied, and flatter in cross-section (Fig. 2.20) with a flattened head. Adult beetles are 25 mm long, grey to brown and are known as click beetles. May be confused with: Larvae are similar to false wireworm larvae. They may also be mistaken for predatory larvae of carabid beetles (Family Carabidae) and rove beetles (Family Staphilinidae). Host range: Sugarcane wireworms are omnivorous. They originally inhabited native grasslands but have adapted to feeding on cultivated crops including field crops and pastures. They are also predatory, feeding on soil invertebrates (McDougall 1934a; Murray 1989). Life cycle on maize: Most individuals complete a single generation in a year but a small number complete two generations in a year. In Qld, adults emerge between late October and early February, with most emerging between November and early December. Adults shelter in refuges for several weeks, then move into the soil, where they may be found to a depth of 7 cm. Three to 4 weeks after emergence, females lay eggs either singly on the soil surface or in batches of 10–15 eggs in crevices to 5 cm deep in the soil. There are eight larval instars with a total average larval duration of 315 days; the last instar, the most damaging, occupies 48% of this time. Larvae pupate in cells in the soil during October to January. Adults emerge after 14 days. Adult females live for a maximum of 7 weeks in the field. In contrast to a number of species of click beetles, sugarcane wireworm adults do not fly to lights. Adults and larvae feed in the soil on vegetation, including roots, and in addition, larvae may feed on soil and invertebrates (McDougall 1934a, b). Risk period: Immediately after sowing and early seedling growth, especially if germination is delayed by cold, wet weather. Damage: Larvae bore into germinating seed and chew on seedling roots and shoots resulting in reduced vigour or seedling death.
Monitoring: Use germinating seed baits (GSB) or soil sampling to detect larvae prior to sowing. Monitor crops after sowing until establishment. Action level: Treatment is required if more than 25 wireworm larvae are found in 20 GSB. Chemical control: Seed dressings, in-furrow sprays and granular insecticides offer some control. Natural enemies: Common brown earwig.
Cutworms Agrotis spp. Lepidoptera: Noctuidae. Pest status: Minor, widespread, irregular. Identification: Larvae are up to 50 mm long, hairless with dark heads and usually darkish coloured bodies, often with longitudinal lines and/or dark spots. Larvae curl up and remain still if picked up. Moths are a dull brown–black colour (Figs 2.23–2.27). May be confused with: Armyworms and Helicoverpa larvae. Host range: All field crops. Life cycle on maize: Usually a single generation occurs during early crop growth stages. Risk period: Crops are most at risk during seedling and early vegetative stages. 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. They usually feed in the late afternoon or at night. By day they hide under debris or in the soil. Monitoring: Inspect crop twice weekly in seedling and early vegetative stage. Larvae feed late afternoons and evenings. Action level: Chemical control is warranted when there is a rapidly increasing area or proportion of crop damage. Chemical control: Insecticide application is costeffective. The whole crop may not need to be
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sprayed if distribution is patchy; spot spraying may suffice. Cultural control: Control weeds 3–4 weeks prior to sowing. Conservation of natural enemies: Cutworms are attacked by a number of predators, parasites and diseases. G ROW T H
Locusts Orthoptera: Acrididae Australian plague locust, Chortoicetes terminifera (Walker) Migratory locust, Locusta mirgratoria (Linnaeus) Spur-throated locust, Nomadacris guttulosa (Walker) N (Main entry in Chapter 18 Locusts and grasshoppers of
pastures and rangelands.)
Pest status on maize: Australian plague locusts may be major pests of maize throughout Australia, migratory locusts are major pests in Qld and spur-throated locusts are minor pests of maize in Qld. All species are irregular in occurrence. Identification: See Chapter 18. Host range: All field crops are susceptible to feeding by Australian plague locusts and spurthroated locusts. Migratory locusts are grass feeders (including maize, sorghum, millet, wheat, barley, triticale and oats), but do not feed on other crops such as sunflower and cotton.
Chemical control: Chemical control may be cost-effective. Control campaigns by the Australian Plague Locust Commission and some State Governments are undertaken during severe outbreaks.
Corn aphid Rhophalosiphon maidis (Fitch) Hemiptera: Aphididae Distribution: The corn aphid probably originated in Asia. It is now distributed worldwide and throughout Australia. Pest status: Minor, widespread, regular. Identification: Adults are 2 mm long, light green to dark olive-green with a purple area at the base of the siphunculi and may have wings. The antennae extend about a third of the body length. Nymphs are similar but smaller in size (Figs 2.9, 2.10). May be confused with: Corn aphids are fairly distinctive, and unlikely to be confused with other aphids on maize. Host range: Mainly sorghum, maize and barley, but occasionally on wheat, oats and triticale. Life cycle on maize: Multiple generations can occur on a crop. Risk period: The corn aphid an affect any crop stage. Damage: Adults and nymphs suck sap. Heavy infestations may turn plants yellow and to appear generally unthrifty. Monitoring: Inspections should be made at weekly intervals.
Risk period: Any crop stage susceptible to damage.
Action level: The incidence of damage is too low to warrant management recommendations.
Damage: Adults and hoppers chew irregular pieces from leaves and stems. They can cause complete defoliation overnight if populations are high enough.
Conservation of natural enemies: Natural enemies (ladybeetles, hoverflies, lacewings and parasitoids) are effective in controlling the pest and are worthwhile conserving.
Monitoring: Check for swarms of adults or bands of nymphs. The Australian Plague Locust Commission provides details of hopper migrations.
Green vegetable bug
Action level: Immediately when swarms and bands are present.
Nezara viridula (Linnaeus) Hemiptera: Pentatomidae N (Main entry in Chapter 7 Pulses—summer.)
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Pest status: Minor in Qld, widespread, irregular. Identification: See Figures 7.45–7.50 and page 198. May be confused with: Redbanded shield bug (Fig. 7.59, page 205). Host range: Maize, soybean, mung bean, navybean, sunflower, cotton, linseed, adzuki bean. Life cycle on maize: Adults invade crop and one generation may develop on a crop during summer. Risk period: Summer and autumn. Damage: Adults and nymphs feed by piercing and sucking on developing cobs. The cobs may be severely deformed. Monitoring: Inspect crop weekly. Action level: There are none available. It is relatively uncommon. Chemical control: Chemical control is costeffective.
in after planting. Beetles are often found near the base of damaged plants. Damaged young plants usually produce suckers. Action level: Beetle numbers in excess of 10 per square metre may result in significant crop damage but control may be warranted with densities of five per square metre or less. Control: Control should aim at preventing beetles feeding on young plants. Chemical control: Chemicals should be applied at planting in the furrow or as a surface band if numbers are less than five per square metre. Cultural control: Avoid planting into recently cultivated pasture that may contain adults. Removal of grass and weeds from headlands removes potential reservoirs. Beetles crawling into a crop can be stopped by cutting a deep furrow with a vertical side nearest the crop. Beetles that concentrate in the furrow can be killed. Sowing delayed until November– December when most beetles have died, but before the new generation of beetles emerges in January, may avoid most beetle activity.
African black beetle Heteronychus arator (Fabricius) Coleoptera: Scarabeidae N (Main entry in Chapter 15—winter pastures.)
Pest status on maize: The African black beetle is a major pest in southern Australian maize, but is not recorded as a pest in Qld. It has a widespread distribution in winter rainfall areas. Its occurrence is irregular, but an outbreak may occur several years in succession in the same location. Identification: See Chapter 13. Risk period: Spring and early summer. Dry springs and summers favour build-up in numbers. Damage: Most damage is caused by adult feeding on the underground stems of young plants, often killing growing points so that the central shoots wither and the plants become dead-hearted. Older plants usually survive, but remain weak and liable to lodging. Monitoring: Beetles may be present in the soil prior to planting, especially if the land has recently grown pasture. Soil sampling will indicate the presence of beetles. Beetles may fly
Whitegrubs Coleoptera: Scarabaeidae Heteronyx spp., Sericesthis spp. and Pseudoheteronyx spp. Distribution: Australia. Pest status: Major, restricted, irregular (Rogers et al. 1992). Identification: Larvae are up to 25 mm long, creamy white with a dark head and are bent in a C-shape. Adult beetles are 7 mm long, oval and uniformly brown. May be confused with: Peanut scarab and black sunflower scarab. Host range: Whitegrubs are native species that complete their life cycle on native hosts, including Eucalyptus spp. Larvae have adapted to feeding on pasture plants and a number of crop species. Life cycle on maize: Depending on the species, there may be a generation every 1 or 2 years. Risk period: Patchy damage signs include wilting and lodging of plants.
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Damage: Larval feeding on roots causes loss of plant vigour and lodging. Beetles may feed on leaves. Monitoring: Whitegrubs are difficult to monitor prior to planting. The presence of small larvae and adults may be an indicator of likely later crop damage. Action level: There are none available as there are no effective controls. Chemical control: None available. Cultural control: Avoid sowing new ground with maize after pasture in areas that have a known history of white grubs. Natural enemies: The fungus Metarhizium and entomopathogenic nematodes (Baker et al. 1997) occasionally cause high mortality of larvae. Birds are often seen eating larvae of whitegrubs.
There are eleven larval instars and total larval development takes between 8–20 months to complete depending on temperature. Most larval growth occurs in the spring–summer period (September–January). Adult whitefringed weevils emerge from pupae in the soil and are active from summer to early winter (December– June) (Matthiessen 1991) when they feed, usually at night, on foliage, leaving scalloped margins along the leaves. Adults are flightless and disperse by walking or being carried in hay or other hosts. Single females reproduce without mating (parthenogenesis). Risk period: Major damage by larvae occurs in autumn–winter. Damage to maize: Larvae chew into lateral roots causing death and reduced vigour. Infestations are usually in patches. Adult weevils feed on leaves but cause little damage. Monitoring: Inspect for adult weevil activity.
Whitefringed weevil Naupactus leucoloma (Boheman) Coleoptera: Curculionidae Pest status: Major in Qld, restricted, irregular. Identification: Soil-dwelling larvae are up to 9–13 mm long, are white to grey with a brown head, and are legless with a slightly curved body (Fig. 15.60). Adult weevils are up to 15 mm long, are grey–brown with a white band along the side of their body and have a short snout (Fig. 15.61). Adult weevils cannot fly and emerge from the soil in summer. May be confused with: Sitona weevil and vegetable weevil. Host range: The whitefringed weevil has a wide host range, including beans, chickpeas, clovers, lucerne, maize, medics, peanut, peas, potatoes, stylo, and tomatoes (Gough and Brown 1991). Life cycle: Usually one generation occurs per year (but may extend over two seasons in southern Australia). Eggs are laid during summer and autumn on plants or debris near the soil surface. Eggs hatch in 17–30 days or several months in the absence of water. Most eggs hatch in autumn and large numbers of firstinstar (non-feeding) larvae appear at this time. During winter, 95% of the whitefringed weevil population is composed of first-instar larvae.
Action level: Mass emergence of adults occurs after rain (November–January). Spray adults. Cultural control: Damage is often worse when two host crops are grown in sequence.
Redshouldered leaf beetle Monolepta australis (Jacoby) Coleoptera: Chrysomelidae N (Main entry in Chapter 7 Pulses—summer.)
Distribution: Australia. Pest status: Minor, restricted to Qld and the north coast of NSW, irregular. Identification: Adults are 6 mm long and are yellow with red–purple patches on the shoulders and ends of the wings. It is a distinctive beetle, unlikely to be mis-identified. (Fig. 7.78). Host range: Maize, sorghum, peanuts, soybean, cotton. Life cycle on maize: Immature stages develop outside the crop and adult swarms invade. Risk period: Any stage of crop growth during spring–autumn. Damage: Swarms of adult beetles move into a crop and feed on foliage, tassels, silks and the husk at the top of the cob. Injury to silks may 129
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reduce seed set and the tips of cobs may be exposed to other insects and fungi.
ends. Body colour can vary with longitudinal stripes (Figs 2.31, 2.34).
Monitoring: Inspect crop weekly. Thorough inspections are required as infestations are usually patchy.
May be confused with: Various armyworm species, Helicoverpa and cutworms.
Action level: Spray if 95% of plants in an area are infested and 70% of flag leaves are eaten.
Host range: Barley, wheat, oats, triticale, sorghum, maize, millet, sunflower. It is less common in other broadleaf crops.
Chemical control: Chemical control is costeffective.
Life cycle on maize: Usually only one generation occurs resulting from an influx of moths laying eggs on the crop.
Swarming leaf beetles
Risk period: Any stage of the crop.
Rhyparida spp. Coleoptera: Chrysomelidae
Damage: The common armyworm feeds on leaves on the outer leaf margins or down the throat of young plants.
Distribution: Australia. Pest status: Minor in Qld except in the north where the black leaf beetle, R. nitida Clark, may be a major pest, restricted, irregular. May be confused with: Flea beetles. Host range: Grasses, maize. Life cycle on maize: Larvae are soil-dwelling and feed on the roots of grasses including maize. Risk period: They are most likely during seedling stage, particularly in north Qld. Damage: Larvae feed on roots and cause seedling death. Severe infestations have resulted in 40% crop loss through death of young plants. Monitoring: No recommended methods available. Action level: Economic injury levels have not been established. Cultural control: Avoid planting maize immediately after grass pasture.
Monitoring: Inspect crops weekly during the seedling and vegetative phases. Action level: Seedling stage: Control is warranted if there is a rapidly increasing area or proportion of crop damage/seedling loss. The whole crop may not need to be sprayed. Vegetative stage: Examine five consecutive plants in the row at six locations throughout the crop (30 plants). Treat if more than 27 out of 30 plants are infested and more than 21 out of 30 have 75% or more flag loss. Chemical control: Chemical control is costeffective. Conservation of natural enemies: Broadspectrum sprays will disrupt activity of natural enemies.
Dayfeeding armyworm Spodoptera exempta (Walker) Lepidoptera: Noctuidae
Common armyworm
Distribution: Australia.
Leucania convecta (Walker) Lepidoptera: Noctuidae
Pest status on maize: Major, widespread in northern areas, irregular.
N (Main entry in Chapter 2 Cereals.)
Distribution: Australia. Pest status: Minor, widespread, irregular. Identification: Mature larvae are 30–35 mm long, hairless with bodies that taper at both
Identification: Fully grown larvae are up to 30 mm long with distinctive green, dark green and yellow longitudinal stripes running the full length of the body (Fig. 4.6). Moths have a 30–40 mm wingspan with speckled dark-grey to black colouration of forewings (Fig. 4.7).
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Fig. 4.6. Two fully grown dayfeeding armyworm larvae. (DPI&F Qld: D. Ironside)
May be confused with: Other armyworms, cutworms and Helicoverpa. Host range: Mostly members of the grass family and also maize, sorghum and sugarcane. Life cycle on maize: Outbreaks follow good rains after a drought period. Usually only a single generation occurs on a crop. Risk period: Any stage of crop growth. Outbreaks usually occur between late December and March, and are more important in north Qld. Damage: Leaves up to 45 cm from the ground are stripped. Damage may not be noticed until the larvae are almost fully grown (Fig. 4.8). Monitoring: Inspect crops weekly in the seedling and vegetative stages. Action level: Seedling stage: Control is warranted if there is a rapidly increasing area or proportion of crop damage/seedling loss. The whole crop may not need to be sprayed.
Fig. 4.8. Dayfeeding armyworm damaging maize. (DPI&F Qld)
Vegetative stage: Examine five consecutive plants in the row at six locations throughout the crop (30 plants). Treat if more than 27 out of 30 plants are infested and more than 21 out of 30 have 75% or more flag loss. Chemical control: Spraying is necessary on crops during outbreaks and effective control depends on early detection and prompt treatment. Conservation of natural enemies: Larvae are subject to fungal and viral diseases, but these normally act too late to prevent serious damage by the pest. Disease and parasites usually ensure there is only one, sometimes two, generations of armyworm during an outbreak.
S I L K I N G TO TA S S E L L I N G
Corn earworm Helicoverpa armigera (Hübner) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Pest status on maize: Major, widespread, regular. May be confused with: Armyworms, cutworms.
Fig. 4.7. Adult dayfeeding armyworm, wingspan 30–35 mm. (DPI&F Qld: J. Wessels)
Life cycle on maize: There can be three to four generations each year, with infestations during both vegetative stage and reproductive stage. Female moths lay eggs on the stem, leaves (both 131
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sides) tassels, silks and husks on the upper twothirds of plants. Caterpillars hatching from eggs prior to silking cause little damage by their feeding on the tassel, but when they migrate to cobs they may cause damage. Caterpillars hatching from eggs laid on silks or husks may cause significant damage. Risk period: Silking–tasselling during spring, summer and autumn. Damage: Corn earworms feed on the top 1–3 cm of cob. Occasionally, medium to large larvae develop on tassels and move onto cob silks. Reduced pollination, resulting in poor seed set and appreciable cob damage, can result. Monitoring: Leaf damage may be an indicator of subsequent cob damage. Parallel rows of holes are signs of feeding on unopened leaves. Check twice weekly during silking, but they are not normally monitored as cob tip losses are considered unpreventable. Action level: Usually corn earworms are not economical to control, except in high-value seed maize. Spray if more than two medium to large larvae per plant (average) are found eating the silks off as, or before, they emerge. Chemical control: Chemicals are registered, but are usually not cost-effective, especially if natural enemies are prevalent or larvae are larger than 5 mm. If warranted, control should be aimed at caterpillars up to 5 mm long on tassels and emerging silks. Cultural control: Maize varieties with husks extending 50–80 mm beyond the top of the cob and closing tightly around the silks restrict the entry of earworm larvae into the cob. Watering during dry weather prevents the husks from loosening. Cultivation to 100 mm (pupae busting) destroys overwintering pupae. Conservation of natural enemies: There are a wide range of natural enemies active and worth conserving.
N (Main entry in Chapter 3 Cotton.)
Distribution: Worldwide Pest status: Major, widespread, irregular. Identification: Adults are 0.5 mm long, have eight legs, are usually yellow–green and have two prominent dark spots on the body. Nymphs are similar but are smaller in size (Fig. 3.11). May be confused with: Strawberry spider mite, bean spider mite. Host range: Most horticultural and field crops, including maize, cotton, soybean, canola, lucerne, peanut, mung bean, navy beans, adzuki bean. Life cycle on maize: Multiple generations build up at the end of the crop cycle. Mite populations build up on lower leaves and move upwards. Risk period: Towards the end of the crop cycle, particularly during late summer/early autumn. Favoured by hot, dry weather. Damage: Adults and nymphs pierce and suck on lower leaf surfaces. A fine webbing is visible on the lower leaf surface. Feeding causes yellowing of the upper leaf surfaces. Heavy infestations will result in leaf desiccation, leaf drop and yield loss. Monitoring: Check the lower leaf surface of older leaves with a hand lens. Check all stages, but numbers often build up rapidly in the later stages of the crop. Initial infestations can be patchy. Action level: None are available as there are no effective controls registered in maize. Chemical control: Not cost-effective. Conservation of natural enemies: Broadspectrum sprays applied against other pests disrupt activity of natural enemies and induce mite outbreaks.
Redbanded shield bug S E E D - F I L L I N G TO M AT U R I T Y
Two-spotted mite Tetranychus urticae Koch Acarina: Tetranychidae
Piezodorus hybneri (Gmelin) Hemiptera: Pentatomidae Distribution: Asia, Australia. Pest status: Minor in Qld, widespread, irregular. Identification: See Chapter 7 Pulses—summer.
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May be confused with: Green vegetable bug.
a
Host range: Maize, soybean, mung bean, navybean, sunflower, cotton, linseed, adzuki bean. Life cycle on maize: Adults invade the crop and may produce one generation on the crop during summer. Risk period: The crop is most at risk during summer and autumn.
b
Damage: Adults and nymphs feed by piercing and sucking on developing cobs. The cobs may be severely deformed. Monitoring: Inspect crop weekly. Action level: There are none available. It is relatively uncommon. Chemical control: Shows some tolerance to insecticides used against the green vegetable bug. See p. 205.
Yellow peach moth
Fig. 4.9. (a) A mature larva of the yellow peach moth (20 mm long) tunnelling into a cob (DPI&F Qld: D. Ironside) , and (b) an adult moth (13 mm long) (DPI&F Qld: J.Wessels) .
Conogethes punctiferalis (Guenèe) Lepidoptera: Pyralidae
persimmon, pomegranate and field crops including maize, sorghum and cotton.
Distribution: Asia, Papua New Guinea, Australia.
Life cycle on maize: A single generation develops from eggs deposited during silking. Several further generations may be completed on other hosts.
Pest status: Minor in Qld, restricted, irregular. Identification: Larvae grow to 20 mm long and have a dark head and are grey–pink with darker oval spots on the body (Fig. 4.9). Moths are 13 mm long and are bright yellow–orange with black markings (Fig. 4.9). May be confused with: The larvae are similar to pink-spotted bollworm larvae. Host range: The yellow peach moth has a wide host range that includes plants such as apple, citrus, cardamom, ginger, grapes, guava, mango,
Risk period: Late summer in central Qld. Damage: Larvae tunnel in stems or cobs in maize and produce masses of webbing and excreta at the tunnel entrance. Monitoring: Inspect weekly after silking. Action level: There are none available. Chemical control: The habit of mining in the cob or stem makes spray application ineffective.
Sources of information Baker, G.L., Poinar, G.O. Jr and Campbell, A.J. (1997) . Psammomermis sericesthidis n. sp. (Nematoda: Mermithidae), a parasitoid of pasture feeding scarab larvae (Coleoptera: Scarabaeidae) in southeastern Australia. Fundamental and Applied Nematology 20: 571–580. Gough, N. and Brown, J.D. (1991). Development of larvae of the whitefringed weevil, Graphognathus leucoloma (Coleoptera: Curculionidae), in northern Queensland. Bulletin of Entomological Research 81: 385–393. 133
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Horne, P.A. and Edward, C.L. (1995). Phenology and food preferences of Labidura truncata Kirby (Dermaptera: Labiduridae) in western Victoria. Journal of the Australian Entomological Society 34: 101–104. Kapuge, S.H., Danthanarayana, W. and Hoogenraad, N. (1987). Immunological investigation of prey–predator relationships for Pieris rapae (L.) (Lepidoptera: Pieridae). Bulletin of Entomological Research 77: 247–254. McDougall, W.A. (1934 a). The determination of larval instars and stadia of some wireworms (Elateridae). Queensland Agricultural Journal 42: 43–70. McDougall, W.A. (1934 b). The wireworm pest and its control in Central Queensland sugar-cane fields. Queensland Agricultural Journal 42: 690–726. Matthiessen, J.N. (1991). Population phenology of whitefringed weevil, Graphognathus leucoloma (Coleoptera: Curculionidae), in pasture in a Mediterranean-climate region of Australia. Bulletin of Entomological Research 81: 283–289. Murray, D.A.H. (1989). Predatory role of Agrypnus spp. larvae in field crops. Proceedings of a Soil Invertebrate Workshop, Indooroopilly, Qld. Robertson, L.N. and Simpson, G.B. (1989). Using germinating-seed baits to detect soil insect pests before crop sowing. Australian Journal of Experimental Agriculture 29: 403–407. Rogers, D.J., Brier, H.B. and Houston, K.J. (1992). Scarabaeidae (Coleoptera) associated with peanuts in southern Queensland. Journal of the Australian Entomological Society 31: 177–181. Simpson, G.B. (1991). Effects of soil type and moisture on development and reproduction of Nala lividipes (Dufour) (Dermaptera: Labiduridae). Journal of the Australian Entomological Society 30: 281–287. Simpson, G.B. and Robertson, L.N. (1993). Effects of germinating seeds of the development and reproduction of Nala lividipes (Dufour) (Dermaptera: Labiduridae). Journal of the Australian Entomological Society 32: 169–175.
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5 OILSEEDS WINTER OILSEEDS F.A. Berlandier and G.J. Baker N Canola, oilseed rape and fodder brassica, Brassica napus Brassicaceae. Origin: Europe, Asia N Linseed, linola and flax, Linum usitatissimum Linaceae. Origin: unknown N Safflower, Carthamus tinctorius Asteraceae. Origin: W. Asia N Sunflower, Helianthus annuus Asteraceae. Origin: N. America
(a) Canola in flower
(b) Canola podding
The winter oilseed season seeding germination growth flowering podding maturation drying harvest
Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec Jan
Indicative phenology of a winter oilseed crop.
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PEST (major pests in bold)
PAGE
Establishment snails and slugs balaustium mite bryobia mite blue oat mite and redlegged earth mite lucerne flea European earwig false wireworms Fuller’s rose weevil small lucerne weevil spotted vegetable weevil vegetable weevil cutworms Growth diamondback moth brown pasture looper chevron cutworm and green cutworms vegetable looper Flowering to podding canola aphids: turnip, cabbage, green peach and potato aphids safflower aphids: green peach and plum aphids Rutherglen bug, grey cluster bug thrips cabbage-centre grub native budworm corn earworm cabbage white butterfly Harvest snails and Rutherglen bugs Summer oilseeds
Winter oilseeds Although winter oilseeds are from different plant families they are subject, in varying degrees, to a similar range of pests. Winter oilseeds are grown within the Australian cropping areas of winter dominant rainfall. Canola is the predominant oilseed with a planted area of about 1.5 million hectares. WA (37%) and NSW (34%) are the main producing states, with Vic. (16%), SA (11%), Qld and Tas. producing the remainder. Canola grows in winter rainfall areas with average rainfall as low as 350 mm. Safflower and linseed plantings are generally less extensive than canola and may vary between seasons. Linseed is being grown as a fibre crop (flax) in southern Qld. Recommended rotation for canola, the most extensively planted oilseed, is: legume pasture–canola–cereal–lupins/field peas–cereals.
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Crops are usually sown near or following autumn rains (April–May–June), and may take up to 34 weeks to reach maturity. Safflower crops may not be planted until June–July. Some oilseed crops may be sown in spring, maturing in as little as 20 weeks.
E S TA B L I S H M E N T
Common white snail, white Italian snail, pointed snail and small pointed snail N (Main entry in Chapter 2 Cereals.)
Pest status on canola: Major, restricted, regular.
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Damage: Snail-feeding may retard development of young plants. At harvest, snails incidentally harvested with seed require removal before sale. Monitoring: Key monitoring times are: (1) January–February to assess options for stubble management; (2) March–April to assess options for burning and/or baiting; (3) May–August to assess options for baiting, particularly along fence-lines; (4) 3–4 weeks prior to harvest to assess the need for header modifications. Monitoring at times (1), (2) and (3) can help target control to areas of high snail density. Action level: In emerging canola crops five snails per square metre is the baiting threshold. Chemical control: For baiting, snails must be active, i.e. there must be cool moist conditions for 5–7 days. Bait should be distributed evenly, and must be completed at least 8 weeks before harvest due to the risk of bait/residue contamination in grain. For five to 80 snails per square metre, apply bait at 5 kg per ha; over 80 snails per square metre, apply bait at 10 kg per ha. Cultural control: See Chapter 2 for stubble and burning management guidelines.
Black-keeled slug and reticulated slug Eupulmonata: Limacidae Milax gagates Draparnaud, and Deroceras reticulatum (Müller) Distribution: Originally from Europe, both species are now present in the Mediterranean basin, North America, South America, New Zealand and Australia. In Australia, both species are mainly pests in the high rainfall (> 500 mm pa) areas of the cropping zone. Pest status on canola: Major, restricted, regular. Identification: The reticulated slug is coloured light grey to fawn with dark brown mottling and grows to 4 cm long (Fig. 5.1) The keel of the reticulated slug is only present at the posterior end of the body. The black-keeled slug is uniform grey to black in colour and grows to 5 cm. The keel on the black-keeled slug runs from the tip of the tail to the mantle (Fig. 5.1) (Smith and Kershaw 1979).
Fig. 5.1. The reticulated slug (upper; 3 cm long), showing brown mottling and milky slime and the blackkeeled slug (lower) with uniform dark colouration. (DAFWA: P. Mangano)
May be confused with: Other pest slugs found in canola crops include the brown slug, Deroceras panormitanum (Lessona & Polomera), the striped field slug, Lehmannia nyctelia (Bourguignat) Eupulmonata: Limacidae and the hedgehog slug, Arion intermedius Normand Sigmurethra: Arionidae. Hosts: Most crop plants, pastures and weeds. Life cycle: In Europe, the life cycle of the reticulated slug is completed in less than 1 year (Lamb and Peters 2001). Slugs, which are hermaphrodites, lay eggs in the soil, probably during autumn. Heavy soils, particularly those which crack and allow movement and protection of slugs, are favourable to slug populations. When surface moisture is available, slugs move on the surface at night and shelter under litter etc. during daytime. As surface moisture dries, slugs may move to 20 cm deep in the soil. The black-keeled slug is probably more of a burrowing species than the reticulated slug. Summer rain promotes buildup of numbers for the following autumn. Slugs eat by rasping plant tissue. Slug-feeding on canola is characterised by serrated edges to cotyledons. Reduced tillage and greater stubble retention of conservation farming are allowing more slugs to survive and breed. Risk period: From emergence of the crop until about 2 months afterwards. Damage: Damage to cotyledons, death of seedlings and reduction of leaf area of establishing plants may occur. Once seedling cotyledons are destroyed, the plants will not recover. Damage often goes undetected as the crop is emerging, and the resultant poor establishment is incorrectly put down to 137
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agronomic factors. Damage may be particularly marked along weedy boundaries and may be more common in higher rainfall districts, but extensive damage can occur in drier areas and in the centre of crops. Damage from European earwig, false wireworms and even the lucerne flea can be falsely attributed to slugs. Determining the true cause of damage is essential for achieving good control. Monitoring: Pre-sowing slug occurrence can be detected by setting bait traps when the soil surface is moist. Bait traps can be made from either hessian bags or upturned 20 cm diameter flower pot bases with gaps to allow entry of slugs, under which are placed several bait pellets. Action level: If slugs are caught under bait traps, baiting of the crop is recommended prior to or at seeding. Chemical control: Either bait the affected area, which is often along fencelines from adjoining pasture, or bait the whole crop if the bait stations reveal an extensive problem. Border treating with higher rates may be necessary when slug numbers are high. Slug bait pellets may remain effective for about a week, so successive applications at a lower rate (5 kg per ha) may be more economical than higher rates, especially if rain is forecast. They should be broadcast evenly on the surface before the crop emerges, and not placed underground with the seed as the slug is less likely to find it. Crops should be monitored at 5–6 day intervals up to the fourth true leaf stage and baiting repeated if necessary.
legs. Under magnification, short stout hairs can be seen covering the body. In addition to being herbivores, balaustium mites are also predatory on other mites when food is scarce. Predatory mites previously recorded as B. murorom (Hermann) were actually B. medicagoense (Halliday 2001). May be confused with: Redlegged earth mites, but adult balaustium mites are larger. Host range: All crops and pasture. Canola and lupins are particularly susceptible. Life cycle: Several generations occur each year. Autumn rainfall is required for oversummering eggs to hatch. Newly hatched nymphs have six legs and are an orange colour. Development from egg to adult takes about 5–6 weeks. Risk conditions: Pre- and post-emergent seedlings, cotyledon and early establishment stages are vulnerable. Autumn crops sown into paddocks that were in pasture the previous year, with high levels of broadleaf weeds, especially capeweed, will be most at risk from mite damage. Damage to canola: Seedlings may be killed after emergence. Retarded and stunted growth of seedlings results in weakened plants (Fig. 5.2). Balaustium mites are recorded as economic pests in WA since 1997. Mites feed on the leaves of plants by probing into the surface cells with their mouth parts, and sucking out sap. Monitoring: Signs of mite activity are mottled or whitened cotyledons. Weeds present in
Cultural control: Cultivation. If canola is to be planted in a high-risk paddock, then early planting will help reduce damage. Biological control: Carabid beetles and other ground-dwelling predators consume slug eggs and active stages.
Balaustium mite Balaustium medicagoense Von Heyden Acarina: Erythraeidae Pest status on canola: Minor, restricted, irregular. Identification: Adult balaustium mites reach 3 mm in length with a greyish-red body and red
Fig. 5.2. Adult female balaustium mites (3 mm long) causing feeding mottling on a canola cotyledon. (DAFWA: F. Berlandier)
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paddocks prior to cropping should be checked to determine the abundance of balaustium mites. Action level: There are none presently available. Chemical control: In areas with a history of balaustium mite damage, incorporation of an insecticide with herbicide immediately prior to sowing is a more effective control strategy than spraying when the crop is emerging and has very little cover of green material. Cultural control: Early control of summer weeds in paddocks that are to be cropped may prevent the build up of mite populations before crop emergence.
Bryobia pasture mite or brown clover mite Bryobia praetiosa (C.L. Koch) Acarina: Tetranychidae N (Main entry in Chapter 15 Pastures—winter
Monitoring: Mite-feeding causes whitish grey spots, giving leaves a stippled, wilted appearance. Chemical control: Chemicals are registered for bryobia mite, but these are not usually costeffective. Cultural control: Early control of summer weeds in paddocks that are to be cropped will prevent the build up of mite populations. Weeds present in paddocks prior to cropping should be checked and if mites are found in large numbers then the incorporation of insecticide with herbicide immediately prior to sowing is a more effective control strategy than spraying when the crop is emerging and has very little cover of green material. Note that rates of insecticides commonly used to control redlegged earth mite and lucerne flea are not effective against bryobia mites. Conservation of natural enemies: Predatory mites can be effective. Predatory ladybirds (coccinellids) also feed on the mites.
rainfall.)
Pest status on canola: Minor, restricted, irregular. Reported as damaging canola in NSW and WA. May be confused with: They are easily confused with redlegged earth mite and are difficult to separate without the use of a hand lens. Bryobia mites usually occur in early autumn, whereas redlegged earth mites occur later as they need cold temperature before hatching. Risk conditions: Summer rains followed by warm mild autumns give bryobia pasture mites the best conditions for survival and increase. They do not tolerate cold wet weather but can persist into June following warm autumn conditions. Crops planted into paddocks with a history of summer/early autumn weeds and warm dry conditions after crop emergence are most at risk. The use of minimum tillage, earlier sowing times and tolerance to some insecticides have led to the increased importance of this pest. Damage: High numbers of bryobia pasture mites can damage emerging canola crops in autumn. Mites feed on the tops of leaves by stabbing into the surface cells with their sharp mouth parts, and sucking out sap.
Blue oat mite and redlegged earth mite N (Main entry in Chapter 15 Pastures—winter rainfall.)
Pest status on canola, linseed, safflower: Major, widespread, regular. Risk period: Pre- and post-emergent seedling, cotyledon stage and early establishment of crop. Damage to canola: Seedlings may be killed either before or after emergence. Retarded and stunted growth of seedlings results in weakened plants. Monitoring: Signs of mite activity are mottled or whitened cotyledons. Mites actively feed in groups from late afternoon to early morning. If spraying with a systemic insecticide at sowing, monitor carefully at 2–3 day intervals by estimating mite numbers in a 10 × 10 cm (100 cm2) ground area. Repeat at five to 10 sites in the crop. Avoid monitoring in bright light. Choose cloudy days and/or early morning or late afternoon. Spray at the first sign of mite activity. Use the TIMERITE® program. Chemical control: The previous spring: Application of a systemic insecticide to pasture during August to early September or heavy grazing by livestock reduces the numbers of 139
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oversummering eggs available to develop in the following canola crop. Control at sowing: Insecticide-treated seed protects young seedlings against low populations of mites. In high-risk paddocks, spraying the surface of the soil with a contact insecticide immediately following sowing kills newly emerged mites. Spraying the seedling foliage with a systemic insecticide is a third option. For this option to be effective, the mites must be detected early by careful monitoring. Perimeter spraying of weedy fence lines and adjacent pasture may reduce mite immigration and hence damage along the edges of canola crops.
Lucerne flea Sminthurus viridis (Linnaeus) Collembola: Sminthuridae N (Main entry in Chapter 15 Pastures—winter rainfall.)
Pest status on canola: Minor, widespread, irregular. Risk period: Seedling stage. Damage: Feeding on leaves produces a thin transparent ‘window’. Seedling growth can be retarded, and in heavy infestations seedlings can die. From a distance, heavily damaged crop areas can appear bleached. They are usually associated with heavier soils and prefer cool, moist conditions. Monitoring: Monitor crops frequently at germination and in the weeks following. Working on hands and knees, look for fleas, checking the soil surface where they may shelter, and for signs of their feeding damage. Inspect 0.5 metre of crop row at five to 10 sites. Action threshold: Spray if on average > 10 feeding holes are found per leaf. Chemical control: Spray with a registered foliar insecticide. Conservation of natural enemies: No effective natural enemies have been identified.
European earwig Forficula auricularia Linnaeus Dermaptera: Forficulidae
Fig. 5.3. Adult European earwigs. The male (left) has a straight section at the base of the forceps, distinguishing this species from other common earwig species found in Australian crops, and the female (right). (DAFWA)
Distribution: Europe, North America. The European earwig is an introduced pest that continues to expand its distribution across southern Australia. Pest status on canola: Minor, restricted, irregular. Identification: European earwigs grow to about 16 mm length, and like other earwigs are characterised by a strong pair of terminal, moveable forceps. The base of male forceps has a straight section (Fig. 5.3), which helps distinguish this species from other common earwigs. Host range: A very wide range of cultivated plants (Hely et al. 1982). Life cycle: Multple generations occur per annum. European earwigs are nocturnal, and during the day shelter under clods of soil, stones and debris. They feed on a wide range of food types including leaf litter and other organic matter, seedling plants, etc. Risk period: Seedling stage and windrowing. They are more prevalent in cool, higher rainfall districts, and appear to be favoured by minimum tillage and greater stubble retention of conservation farming. Damage: Immature and adult earwigs can occur in high densities in germinating canola, where they lop young plants off at ground level (Fig. 5.4), requiring re-seeding (Fig. 5.4).
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a
Cultural control of canola establishment pests
b
Canola crops sometimes follow a pasture phase. The risk from pasture-dwelling pests, such as redlegged earth mite, blue oat mite and false wireworms, can be reduced by a fallow period between cultivation of the pasture and the sowing of the crop (Horne and Edward 1998). Some pests can multiply on or take shelter on weeds. Maintenance of a clean fallow by occasional cultivations, and clean cultivation of headlands during summer and autumn, can help stop pests from breeding or sheltering oversummer. Burning wheat stubble for snails also controls establishment pests in the following canola crop.
Eastern false wireworm, Pterohelaeus darlingensis Carter Fig. 5.4. (a) Leaves of a rosette-stage canola plant scalloped by European earwig feeding. Earwigs are night feeders that usually hide under litter during the day (SARDI: G.J. Baker) . (b) A canola crop damaged by feeding of European earwigs on seedlings (DAFWA: A. Dore) .
If present in high densities, they can also damage mature canola by feeding on the pods, and may contaminate the harvested grain. Monitoring: Search by torchlight at night, or place out artificial refuges, such as carpet squares, for several days as outlined for bronzed field beetle monitoring. Action threshold: Unknown. Chemical control: No insecticide treatments are currently registered for European earwig control in canola. Conservation of natural enemies: No effective natural enemies have been identified.
False wireworms Coleoptera: Tenebrionidae The main species of false wireworms causing crop damage include:
Bronzed field beetle, Adelium brevicorne Blessig
Striate false wireworm, Pterohelaeus alternatus Carter Grey false wireworm, Isopteron (=Cestrinus) punctatissimus (Pascoe) Southern false wireworm, Gonocephalum macleayi (Blackburn) Distribution: Native to Australia, distributed across southern and eastern Australia. The bronzed field beetle is recorded as a pest of canola in lower Eyre Peninsula, mid-north and south-east SA (Miles 1997) and south-western WA. The eastern false wireworm is recorded as a pest in NSW, Qld and Vic., the striate false wireworm in northern NSW and western Qld (Elder et al. 1992), the grey false wireworm in NSW, the southern false wireworm in NSW and Qld, and Gonocephalum spp. in SA and Vic. In addition, some species with localised distributions are reported as being associated with crops: Gonocephalum adelaidae (Blackburn) in western Vic., Helea tuberculatus in NSW, and Saragus sp. in parts of SA (Allsopp 1979; Horne and Edward 1998). Pest status on canola, linseed, safflower: Major, restricted, irregular. The status of individual species varies. The eastern false wireworm is not regarded a damaging species in Vic. 141
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wireworms and small false wireworms, recorded as pests in northern NSW and Qld oilseed crops, are illustrated in Fig. 5.27.
a
May be confused with: False wireworm larvae resemble other soil-dwelling larvae including true wireworm (or click beetle) larvae, which are cream-coloured but soft-bodied and flatter than false wireworms (Fig. 2.19). Fully grown larvae of the bronzed field beetle are larger than grey false wireworm larvae and appear rounder in cross-section.
b
Fig. 5.5. (a) Fully grown larvae of the bronzed field beetle (12 mm long), and (b) adults. (SARDI: G.J. Baker)
Identification: The larvae of false wireworms have elongate, generally cylindrical bodies, usually hard, with three pairs of legs behind the head. Bronzed field beetle larvae are dark brown with fairly soft segments, the last of which has two upturned spines (Fig. 5.5). Larvae grow to 12 mm. Larvae of the eastern false wireworm grow to 50 mm, are light cream to tan with darker rings around each segment and do not have large protrusions on the tail. Larvae of the grey false wireworm grow to about 9 mm, are coloured greyish brown and have a pair of upturned spines on the tail (Fig. 5.6). Larvae of the southern false wireworm grow to 20 mm long, are coloured cream, yellow or tan with spinelike hairs on the tail. Adults of bronzed field beetles are 11 mm long and shiny black in colour with a bronze sheen under certain lights. Adults of eastern false wireworms are 25 mm long, ovoid shape with flanged edges on the dull black body. Adult beetles of grey false wireworms are slender, dark brown and grow to about 8 mm in length (McDonald 1995). Adult southern false wireworms are generally dark brown–grey, oval beetles with flanged edges, giving them a pie-dish shape. They sometimes have a coating of soil on the body. Striate false a
b
Fig. 5.6. (a) Grey false wireworm larvae, and (b) an adult. (SARDI: G.J. Baker)
Host range: Originally, native grasses (McDonald 1995), but larvae have now adapted to eating most winter-sown crops. Canola appears to be particularly susceptible to the bronzed field beetle, whereas cereal and legume crops appear unaffected. Life cycles: Adult beetles lay eggs in or just below the soil surface, mostly in stubble and crop litter on which the developing larvae feed. They are often found in light, dry soils with high organic content (Dunn and Miles, undated). When the soil is reasonably moist, the larvae tend to aggregate in the top 10–20 mm of plant litter. As the soil dries, the larvae move lower down, but may move to the surface to feed at night when the soil surface is dampened by dew. It is possible that drying soils may stimulate wireworm larvae to feed on germinating crops. Minimum tillage and stubble retention farming favours the build-up of false wireworm populations in field crops and pastures (McDonald 1995). The number of generations per year may vary with seasonal conditions and availability of food (G. Goodyer, unpublished). It is possible that generations of some species overlap. Adult bronzed field beetles shelter in crop residues during summer and autumn and become active after autumn rains. Egg-laying by females may commence before seeding and by the time of seeding, larvae may be 5 mm long when they are most damaging. About 15 adults per square metre produce up to 1500 larvae. During the day, larvae shelter in stubble, clods of dirt or in cracks. At night they move from shelter to feed. Depending on rainfall, eggs may be present from March to May, larvae from April to September, pupae from August to October, and adults from August to the following May, by which time most have died (Michael et al. 2002).
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representative parts of the field (five to 10 sites), and count and record the number of wireworm larvae. Eastern and southern false wireworm larvae are easily detectable, but grey false wireworm larve are small and difficult to detect. Germinating seed baits may also be used (see Glossary), but in southern Australian crops they yield estimates similar to direct counts (with the aid of a sieve). Before sowing: Adult numbers of bronzed field beetles can be estimated before sowing using artificial refuges such as carpet squares, placed for several days in areas of previous damage or abundant litter.
Fig. 5.7. Chewing by larvae of bronzed field beetles (arrowed) has felled this established canola plant. (SARDI)
The eastern false wireworm has two generations per year; the larvae of the winter generation are active between autumn and spring and the adults aestivate during spring and summer. The larvae of the summer generation are active during spring and summer, with adults aestivating during autumn and winter. The grey false wireworm has one generation per year, with larvae active between autumn and spring, with adults aestivating during late spring and summer. The southern false wireworm has two generations per year, with larvae active at similar times to those of the eastern false wireworm. Risk period: Between sowing and establishment of crop. Once the damage has become obvious it is too late to treat the crop. Damage: Larval feeding on roots and underground stems may stunt or kill seedlings (Fig. 5.7), thinning the crop or, in severe cases, leaving bare patches in the crop. Adults chew on seedlings at or above ground level. Canola sown into cereal stubble is particularly at risk. Half-grown bronzed field beetle larvae at a density of 1500 per square metre kills most canola seedlings. Monitoring: The previous spring: Carefully look in the top-soil in 30 cm × 30 cm quadrats. Look to just below the junction of loose, dry, cultivated soil and the undisturbed moist soil below in
Action threshold: Indicative action level for bronzed field beetle adults is five or more adult beetles per square metre. Indicative levels for eastern false wireworm larvae in NSW is 12 per square metre and for grey false wireworm is 35– 50 per m2. In Vic., an average of 25 eastern and southern false wireworm larvae or 50 grey false wireworms per m2 are indicative thresholds. Chemical control: Soil-incorporated insecticides used prior to sowing and seed dressings are commonly used to prevent larval attack. Broadscale use of insecticides may severely impact on non-target organisms. Cultural control: Clean cultivation of fields and headlands during summer and autumn removes
Agronomic trade-off: minimum tillage favours some soil pests Increasing the use of minimum tillage in winter field crops appears to have elevated to pest status a number of previously uncommon species. Bryobia pasture mites, true wireworms, a number of species of false wireworms, the bronzed field beetle, slugs, snails and pasture webworm appear to have benefited from undisturbed soil, together with the food and shelter offered by organic matter that accumulates on the surface. There is evidence that this environment also encourages natural enemies of these pests but there is presently no indication of their effectiveness and how to manage them (Horne and Edward 1998).
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food and shelter of adult beetles. Where threshold densities are present, increasing the seeding rate and sowing shallow may maintain crop density. Compaction of the seed bed may reduce damage.
Fuller’s rose weevil Asynonychus cervinus (Boheman) Coleoptera: Curculionidae Distribution: Originally from South America, now cosmopolitan. Pest status: A minor pest of canola in WA, with irregular occurrence. Host range: These weevils have an extensive recorded host range of broadleafed plants (Hely et al. 1982). Identification: Adult weevils are grey–brown, and up to 8 mm long. Mature larvae are yellow and up to 6 mm long. Fuller’s rose weevil may be confused with the small lucerne weevil. Life cycle: The life cycle is completed in 1 year from egg to adult. Adults emerge from pupal soil cells in autumn. Damage and control: Similar to that for small lucerne weevil.
Small lucerne weevil
Damage: Adults weevils can damage seedling and young crops by chewing the base of the plant. The edges of the crop tend to be attacked first. Monitoring: Sample for weevils at night, as they are not active during the day when they hide in surface residues or crevices in the soil. Weevils are less active when temperatures drop below 10°C. Infestations spread slowly because the weevils cannot fly, and spread by crawling instead. Action level: 50–100 weevils per square metre. Chemical control: At present, no insecticide is registered for small lucerne weevil in canola in Australia, but chemicals registered for other weevils in canola may be effective. Synthetic pyrethroid efficacy is reduced when applied in paddocks with little plant cover. Cultural control: Rotary hoeing will disrupt and kill some weevil pupae in soil cells, thereby reducing the numbers of emerging adults.
Spotted vegetable weevil Desiantha diversipes (Pascoe) Coleoptera: Curculionidae N (Main entry in Chapter 2 Cereals.)
Pest status on canola: Minor, restricted, irregular.
Atrichonotus taeniatulus (Berg) Coleoptera: Curculionidae
May be confused with: The small lucerne weevil is a lighter colour than the spotted vegetable weevil, and about twice the size.
N (Main entry in Chapter 15 Pastures—winter rainfall.)
Risk period: Immediately after sowing.
Pest status on canola: A minor pest recorded on WA canola, irregular occurrence. May be confused with: The spinetailed weevil (Fig. 2.17), the adults of which are a darker colour and smaller than the small lucerne weevil. Host range: A range of broadleafed field crops, including canola, and weeds including capeweed. Risk period: Seedlings and young crops are at greatest risk from emerging adult weevils, which can damage many plants. In some cases, paddocks need to be re-sown. In some years, high weevil numbers have been noted in paddocks with soils containing gravel.
Damage: Adults can damage some germinating canola crops. They are often distributed across much of the paddock, unlike the vegetable weevil that is usually confined to crop edge. Paddocks most at risk are those that were in pasture the previous year and had an abundance of capeweed, a preferred host.
Vegetable weevil Listroderes difficilis Germain Coleoptera: Curculionidae Distribution: Originated in South America, now in North America, South Africa, east Asia, New Zealand and Australia.
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Fig. 5.8. A larva of the vegetable weevil (13 mm long) and feeding damage on the leaf. (DAFWA: F. Berlandier)
Pest status on canola: Minor, widespread, irregular. Identification: Larvae grow up to 13 mm long, are legless, with curved bodies coloured green, yellow–green or cream, with a pale yellow to brown head and a brown plate behind the head (Fig. 5.8). Adult weevils are about 8 mm long, dull greyish brown with a V-shaped pale mark near the middle of the back. They have prominent snouts and a small pointed process towards the rear of each wing cover. May be confused with: Unlikely to be confused with other canola insects. Host range: Most vegetables, especially brassicas (including canola), and a wide range of weeds, especially capeweed. Pasture legume species and grasses are not preferred hosts. Life cycle: One generation occurs per year. Adult female weevils are capable of laying several hundred eggs. Eggs laid in the surface litter of soil in autumn hatch into larvae that feed during the night on developing canola plants and shelter in the soil during daytime. The fully grown larvae pupate in cells in the soil in early spring. Adults emerge in spring, and like the larvae, adults feed on leaves of canola plants at night, moving to the soil during the day. During summer, adults remain inactive in the soil. Vegetable weevils are flightless (Hely et al. 1982). Risk period: Seedling stage, which may be attacked by both adults and larvae. Damage: Seedlings may be retarded (Fig. 5.9) or killed by feeding. Damage can be severe if the larvae or adults are numerous and seedling growth is impaired by adverse weather.
Fig. 5.9. Vegetable weevil damage to canola seedlings. Small serrations on the leaf margin distinguish weevil damage from the scalloping caused by earwig-feeding. (Graphic Science: © Denis Crawford)
Monitoring: Inspect emerging crops at night, especially at the edges where damage may be concentrated. Check for damage and the presence of larvae and adults. Action threshold: As indicated by leaf damage and seedling death. Chemical control: Spraying damaged patches may be cost-effective. Cultural control: Alternate cropping with nonhost plants or a long weed-free fallow prior to planting minimises weevil numbers. Planting a canola crop away from previously affected land reduces the risk of invasion. Conservation of natural enemies: No effective natural enemies have been identified.
Cutworms Agrotis spp. Lepidoptera: Noctuidae N (Main entry in Chapter 2 Cereals.)
Pest status on canola, linseed, safflower: Minor (but occasionally causing major damage in restricted areas), widespread, irregular. Damage to canola: Plants are killed after being cut at the base by larval feeding, at or below ground level, and are severely damaged by larvae climbing plants and consuming the leaves. Large patches of the crop may be damaged if larvae are numerous during establishment. Risk period: Emergence and establishment. 145
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Monitoring: Look for signs of chopped stems, which may occur in patches, especially along fencelines bordering pastures. Early detection of larvae or plant damage essential. Inspect plants, soil litter and the soil surface in 0.5 metre of row (ideally in the evening or at night when larvae are active). Repeat at five to 10 sites. Spray if two or more large larvae per 0.5 metre row are found. Chemical control: Spot-spray detected patches with a contact insecticide. This is most effective at night when larvae are exposed. Cultural control: Clean-fallow and eliminate weeds around the perimeter at least 1 month before sowing.
G ROW T H
Diamondback (cabbage) moth Plutella xylostella (Linnaeus) Lepidoptera: Plutellidae Distribution: Cosmopolitan. In Australia, occurs wherever introduced brassica plants are grown. a
b
Fig. 5.10. (a) Diamondback moth larvae. The mature larva (lower) is 12 mm long. (b) The moth (10 mm long) has a characteristic pattern on its back. (UnivAd: M. Keller)
Pest status: Canola: minor, widespread, irregular. Identification: Yellow, oval-shaped eggs are laid on the underside of leaves. Larvae are pale green and up to 12 mm long (Fig. 5.10). Young larvae mine leaves; mature larvae perforate leaves from the underside. Pupae are usually found on the underside of leaves encased in a fine cocoon. The adult moth is 8–10 mm long, has grey– brown wings with a medial white stripe which, when the wings are folded, appears as a series of diamonds (Fig. 5.10). May be confused with: Larvae of the native budworm, cabbage centre grub and cabbage white butterfly, all of which have legs and may occur on canola. Larvae of the vegetable weevil, which are legless, may also be found on canola. The diamondback moth is usually the most numerous and smallest caterpillar found on canola and this, together with their uniform colour and habit of vigorous wriggling and hanging by a silken thread when disturbed, readily distinguishes them from other canola caterpillars. Host range: A wide range of introduced Cruciferae, including brassica vegetables, fodder rape, mustard, canola and many cruciferous weeds. Life cycle: Development from egg to adult may take as little as 14 days in summer, extending to 21 days or more in winter (Cook et al. 2000). The diamondback moth does not hibernate during winter but development of pupae may be prolonged. Female moths lay eggs soon after emergence and lay an average of 160 eggs during their life of some 16 days. The mines that young larvae eat in leaves offer them some protection from natural enemies and pesticides. Older larvae feed on the surface of leaves where they are more exposed. As the plant matures, larvae move from leaves to buds, flowers and developing pods. The entire life cycle may be completed on the host plant, on which several generations may develop between autumn and summer. Where host plants are locally available, adult moths do not fly far (about 40 m) (Cook et al. 2000). Cruciferous weeds (especially wild radish) or other crucifer crops near to newly planted canola crops may act as reservoirs of
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diamondback moths. Cruciferous weeds and volunteer canola induced by summer rains provide a bridge for the moth to invade autumn sown crops. Risk periods: During the bolting stage of the crop and any time the crop is water-stressed. Larval survival is favoured by dry, warm conditions. Mortality is increased following rainfall, which may wash off some larvae, and may induce disease. Damage: Reduction of leaf area of young crops and damage to floral parts of older crops affects yield (Fig. 5.11). At densities of 500 larvae per 10 sweeps, crop yield may be reduced by 0.2–0.3 t per ha and densities of below 200 larvae per 10 sweeps may reduce yield. Monitoring: By sweep net during autumn, winter and spring. Action level: For SA and Vic., 20 larvae per 10 sweeps during mid to late flowering, 50 larvae per 10 sweeps at pod maturation. For WA, 100 larvae or more per 10 sweeps. a
b
Fig. 5.12. Larva of diamondback moth infected with the fungus Zoophthera radicans. During wet periods, this fungus may contribute to controlling diamondback moth populations. (SARDI: G.J.Baker)
Chemical management: One spray may be costeffective on young, moisture-stressed crops, or during the bolting stage if densities exceed 100 larvae per 10 sweeps. Protected stages (larvae in mines, pupae) may survive a single spray application. Some diamondback moth populations have developed resistance to some insecticides. Where numbers are patchy in the crop, spot spraying these patches may suffice. Diamondback moth populations outside the crop (for example on weeds and self-sown canola) form a reservoir of insecticide susceptible genes, so spraying these should be avoided. Cultural control: Removal of all crucifers (especially weeds) at least 4 weeks prior to sowing may reduce bridging populations of diamondback moths. Reservoirs of diamondback moth may be avoided by planting the new canola crop distant from the previous year’s crops and removing cruciferous plants growing adjacent to the new canola crop. Conservation of natural enemies: Diamondback moth has a number of parasitoids, generalist predators and a fungus disease (Fig. 5.12), which together reduce diamondback moth populations. In many years, they do not prevent economic damage to canola crops.
Brown pasture looper Ciampa arietaria (Gueneé) Lepidoptera: Geometridae Fig. 5.11. (a) Damage by the diamondback moth larvae to flowers (arrowed) reduces yield, while (b) feeding on pods causes relatively minor yield loss. (SARDI: G.J. Baker)
Distribution: Native. Occurs in all Australian winter oilseeds areas. 147
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a
Fig. 5.13. Larva of brown pasture looper. (SARDI: G.J. Baker)
Pest status on Canola: Minor, restricted, irregular. Identification: Brown pasture looper larvae crawl with a looping motion and are 35–40 mm long when fully grown. They have yellow– brown heads and are grey to dark brown, with a pale stripe down each side of the back and a row of reddish spots along each side of the body (Fig. 5.13). Host range: Broadleaf pastures, lupins and canola crops. Capeweed is a favoured host. Life cycle: Brown pasture loopers have only one generation per year. Moths fly from March to June and eggs are laid at this time. These eggs hatch and caterpillars grow to full size in about 2 months, pupate and remain in the pupal stage over spring and summer, until they emerge as moths in the next season. Risk period: Vegetative stage. Damage to canola: Brown pasture loopers are early season defoliators, often prevalent around patches of capeweed and at the edge of crops, where they have moved from adjacent pasture. Spot- or perimeter-spraying is usually all that is required.
Cutworms Lepidoptera: Noctuidae Chevron cutworm, Diarsia intermixta (Guenée) Green cutworms, Neumichtis niderrima (Guenée), N. spumigera (Guenée) and N. saliaris (Guenée) Distribution: Native species, recorded in NSW, SA, Qld, Tas. and Vic. Pest status: All are minor pests of fodder brassicas in Tas.
b
Fig. 5.14. (a) A chevron cutworm (30 mm long), and (b) green cutworm (35 mm long) showing two white spots on its rear end (right). (DPI&W Tas: L. Hill)
Identification: Larvae of the chevron cutworm grow to 30 mm long, are counter-shaded; dark brown to black above, black below. The larva has a series of paired, black, short, oblique marks along the back and a pale mark traversing the rear end (Fig. 5.14). Green cutworm larvae grow to 35 mm long, are green or brown in general colour with a pair of small but distinct white spots on the top of the hind end (Fig. 5.14). These cutworm species may occur in mixed populations (McQuillan et al. 2006). Host range: Chevron cutworms feed on brassicas, including brassica weeds, docks, and crop plants including fodder beet, corn and clovers. Green cutworms feed on brassicas, broadleafed weeds and some vegetable crops (McQuillan et al. 2006). Life cycle: There are probably two generations per year. Larvae pass through six instars. Under favourable conditions, the duration from egg-lay to adult emergence is 8–11 weeks, depending on species (McQuillan et al. 2006). Risk period: Autumn and winter. Damage: Larval-feeding reduces leaf area. Unlike Agrotis spp. cutworms, these species do not characteristically cut young plants at their
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Pest status on oilseeds: Minor, widespread, irregular. Caterpillars are grass-green, smooth and slender, and move with a characteristic looping action. They are 2.5–3 cm long when fully grown (Fig. 5.15). They feed on canola foliage, but rarely if ever cause economic damage (Baker 2001).
Fig. 5.15. A larva of the vegetable looper (30 mm long). They are common foliage feeders on canola, but cause little damage. (Graphic Science: © Denis Crawford)
F LOW E R I N G TO P O D D I N G
Canola aphids
base. Dense populations may cause bare areas in the crop.
Turnip aphid, Lipaphis erysimi (Kaltenbach) Hemiptera: Aphididae
Monitoring: Larvae shelter at the base of plants during the day but may be seen feeding on exposed foliage during evenings and at night.
Cabbage aphid, Brevicoryne brassicae (Linnaeus)
Control: Control in forage crops is seldom recommended. Spot-spraying with a registered insecticide may sometimes be cost-effective. Natural enemies are thought to provide adequate control in most years.
N (Main entry in Chapter 8 Pulses—winter.)
Vegetable looper
Pest status: Canola: major, widespread, irregular.
Chrysodeixis argentifera (Guenée) Lepidoptera: Noctuidae N (Main entry in Chapter 7 Pulses—summer.)
Distribution: A native species, recorded in all states.
Green peach aphid, Myzus persicae (Sulzer)
Potato aphid, Macrosiphum euphorbiae (Thomas) Distribution: These four aphids are distributed worldwide.
Identification: It is important to distinguish if green peach aphids are in the crop, as unlike the other species mentioned here, some populations of green peach aphids are resistant to some insecticides (Berlandier 1999).
Table 5.1. Distinguishing features of green-coloured aphids found in canola crops Turnip aphid (Fig. 5.16)
Cabbage aphid (Fig. 5.16)
Green peach aphid (Fig. 5.17)
1.4–2.4 Yellowish to olive– green Body sometimes with waxy dusting, dark bars on abdomen, dusky wing veins
1.6–2.8 Greyish to mid-green
Colony habit
Dense colonies, usually around growing tips
Abundance on canola
Usually the most common aphid on canola
Dense colonies, but usually not in association with turnip aphid Common on canola, depends on season
1.2–2.3 Shiny yellow to midgreen to pink or red Black patch on abdomen of winged adults. Wingless forms are uniform in colour Sparse colonies. May occur with turnip or cabbage aphid, prefers lower leaves Common but seldom forming large colonies
Length of adult (mm) Abdomen colour Other features
Body covered with a dense white mealy wax
Potato aphid 1.7–3.6 Light yellowish to pinkish-green Red eyes
Small colonies
Rare
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a
b
Fig. 5.16. (a) Olive-coloured turnip aphids on inflorescence of canola (F. Berlandier) , and (b) nymphs and wingless adult cabbage aphids with waxy coating on a canola stem (Graphic Science: © Denis Crawford) .
Hosts: Turnip aphid and cabbage aphid: Brassicaceae, particularly the weeds wild radish, and turnip weed. Green peach aphid has a large host range that includes over 40 different plant families; common hosts in field cropping regions of Australia include capeweed, wild radish and lupins. Potato aphid is polyphagous, with over 200 hosts recorded in 20 plant families.
Life cycle: Winged aphids migrating from weeds start colonies on canola seedlings or young crops in autumn. Temperatures during autumn and spring are optimal for aphid survival and reproduction. During these times, the aphid populations may undergo several generations. Populations peak in late winter and early spring; development rates are particularly favoured when daily maximum temperatures reach 20–25°C. Risk periods: Autumn and spring but especially during bud formation and flowering. Summer rain favours increase in numbers in autumn. Damage: Feeding damage. Aphids suck the sap of canola plants and ‘moderate’ numbers can affect yield and oil quality in susceptible varieties and crops under drought stress. Yield losses of up to 33% have been recorded in WA. Dense colonies feeding on upper stems, developing flower heads and seed heads may reduce pod set, seed fill and grain quality. Canola can compensate for aphid damage but not drought stress. Aphid-vectored viruses can also contribute to yield loss (see box). Monitoring: Weekly from late winter to early spring, especially from flowering to grain fill. Aphid distribution may be patchy, so at least four sampling points of 20 plants each should be spread over the field. Action levels: Drought-stressed crops are at greatest risk; aphid colonies increase rapidly on such crops. WA: As a guide, 20% of plants infested with aphids depending on cultivar. Crops in high rainfall zones (> 450 mm) during seasons of average to above average rainfall are at least risk from aphid-feeding damage. SA and Vic.: (low rainfall) 25 mm+ of stem infested on 20% of plants. Qld: One colony (about 10 mm length) on most flower heads Cultural control: Early control of weed hosts, particularly wild radish and wild turnip, reduces the reservoir population of aphids. Sowing early allows crops to establish before aphids become numerous.
Fig. 5.17. Non-colonial green peach aphids are shiny, and wingless forms have a uniform colour. (Graphic Science: © Denis Crawford)
Conservation of natural enemies: Parasitic wasps and a number of generalist predators such as hoverflies (Fig. 5.18) (commonly Melangyna viridiceps (Maquart) and Simosyrphus grandicornis (Macquart)), ladybirds (Figs 3.46
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a
Aphid-transmitted virus diseases in Australian canola crops
b
Beet western yellows luteovirus (BWYV) is the most common virus in WA canola crops (Coutts and Jones 2004). BWYV is persistently transmitted by a range of aphid species, with green peach being the most important vector. A combination of BWYV and green peach aphids causes yield losses of up to 50% in canola (Jones and Hawkes 2002). Insecticide treatment of canola seed is the most successful method to manage this virus (Jones et al. 2003).
Safflower aphids Hemiptera: Aphididae Green peach aphid, Myzus persicae (Sulz.) Leafcurl plum aphid, Brachycaudus helichrysi (Kaltenbach) c
Thistle aphid, Capitophorus elaeagni (del Guercio) Pest status: Minor, widespread and irregular pests of safflower crops. Aphid-feeding may damage seedling plants but is more often evident during budding and flowering. Severely damaged buds and flower heads turn a mottled yellow and become distorted and shrivelled (G. Goodyer, unpublished).
Fig. 5.18. Some natural enemies of canola aphids: (a) the fawn, bloated aphid bodies (mummies) contain the wasp parasitoid Diaretiella rapae McIntosh Hymenoptera: Aphidiidae (Graphic Science: © Denis Crawford) , (b) adult hoverfly feeding on brassica flower (SARDI: P.T. Bailey) , and (c) yellow, nobbly-skinned larvae (6 mm long) of hoverflies are effective predators of aphids (SARDI: G.J. Baker) .
and 3.47) and lacewings may exert significant control of aphid populations. They may be conserved by sparing use of aphid-specific insecticides. Chemical control: Systemic insecticides may be cost-effective when action levels are exceeded. Some green peach aphid populations are resistant to some insecticides; if this is the dominant species, consider insecticides from the carbamate group.
Rutherglen bug Nysius vinitor Bergroth
Grey cluster bug Nysius clevelandensis Evans Hemiptera: Lygaeidae Pest status on winter oilseeds: Rutherglen bug: major, widespread, irregular. Grey cluster bug: major, reported mainly as a pest in Qld, regular (Elder et al. 1992). Risk period: From flowering until after seed heads have dried. Rutherglen bugs move into crops as host weeds (e.g. capeweed) dry off. Infestation is most likely during hot, dry 151
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weather and damaging populations can build up rapidly.
a
Damage: Nymphs and adults suck sap from leaves, stems, flowers and pods. Seed yield, oil quality and seed viability may be reduced by feeding (Broadley et al. 1986). Damage is aggravated if the crop is water-stressed. Monitoring in canola: Canola: inspect crops regularly from flowering to windrowing. Inspect heads visually, or by shaking into a bucket. Examine 20 heads. Repeat at five to 10 sites. Safflower (Qld): examine plants regularly before and after flowering (this can be done at the same time as monitoring for Helicoverpa spp.). Examine five consecutive plants in a row at six locations (Elder et al. 1992). Action level: Canola: 10 adults or 20 nymphs per plant. Higher numbers can be tolerated if moisture is not limiting (Colton and Sykes 1992). Safflowers can compensate for up to 40% bud loss (Colton 1988); indicative threshold (Qld) preflowering is 300 adult bugs on 30 plants or postflowering 150 adults (Elder et al. 1992). Chemical control: Insecticides may be costeffective. Spraying during flowering is avoided to allow bee activity. Spray before windrowing if numbers are high to avoid damage to windrowed seeds. Cultural control: Cultivating or herbiciding weeds near to crops should be avoided.
Safflower thrips
b
Fig. 5.19. (a) Fully grown larva of cabbage-centre grub (12 mm long) among webbed canola leaf (Graphic Science: © Denis Crawford) , and (b) moth (12 mm long) (SARDI: G.J. Baker) .
Distribution: Australia and New Zealand. Occurs in all Australian winter oilseeds areas. Pest status on canola: Minor, restricted, irregular. Sometimes damages fodder rape crops. Risk period: Flowering stage.
Tomato thrips, Frankliniella schultzi (Trybom)
Identification: Moths are 12 mm long with mottled brown wings (Fig. 5.19). The creamish larvae have dark heads, longitudinal reddishbrown stripes and when full-grown are about 12 mm long (Fig. 5.19).
Pest status: Minor, widespread, irregular.
Host range: Plants in the Family Cruciferae.
Thrips may damage foliage and flowering parts of plants. Bronzed buds are indicative of thrips feeding, particularly if the crop is waterstressed. Although plants can compensate for bud damage, thrips control just before flowering may be warranted if about 25% of buds are dead and thrips are still active.
Life cycle: Several generations each year. More common in warm, dry cropping areas.
Cabbage-centre grub
Native budworm
Hellula hydralis Gueneé Lepidoptera: Pyralidae
Heliocoverpa punctigera (Wallengren)
Thysanoptera: Thripidae Onion thrips, Thrips tabaci Lindeman
Damage to brassicas: Larvae attack foliage and developing floral parts, but numbers are seldom high enough to cause economic loss. This species may cause economic damage to fodder brassicas.
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adult budworm moths may be attracted to the crop by honeydew secreted by aphids. Caterpillars of the common armyworm, Leucania convecta, sometimes occur in large numbers in oilseed crops. They feed only on grasses and weeds and should not be mistaken for native budworms or corn earworms.
Corn earworm H. armigera (Hübner) N (Main entry in Chapter 3 Cotton.)
Pest status of native budworm: Canola: minor, widespread, irregular. Linseed: major, widespread, regular. Safflower: minor, widespread, regular. Pest status of corn earworm: Canola: not recorded as a pest. Linseed: minor, restricted, regular. Safflower: minor, restricted, regular. Canola: Small native budworm caterpillars feed mainly on foliage but older larvae (up to 3.5 cm long) may eat flowers, young pods and bore into the maturing pods (Fig. 5.20). Canola is not a preferred host for the native budworm; however,
Monitoring: Pull up plants and beat into a container. Sample 10–20 plants, and count the number of caterpillars collected. Determine density (number of plants per sq m) according to average crop density. Spray if five to 10 or greater caterpillars per square metre are found. Linseed: Young native budworm larvae enter flower buds, flowers and young bolls in which they develop, but it is the feeding of mobile larger (7–20 mm) larvae that causes the most damage. Weekly monitoring of buds and flowers for larvae and damage indicates whether control is warranted. More than six 7–20 mm long larvae per 30 terminal buds beat into a bucket from 30 widely spaced plants is an indicative control threshold in Qld. At least two sprays, one before early flowering and a second at late flowering, may be necessary to control rapidly developing populations during early flowering. Corn earworm larvae may be expected in crops grown within 30 km of summer irrigated crop areas. These should be controlled as eggs or young larvae if chemical control is indicated. Safflower: Unstressed safflower plants may compensate for pre-flowering loss of up to 40% of buds and developing flower heads.
a
Cabbage white butterfly Pieris rapae (Linnaeus) Lepidoptera: Pieridae Distribution: Cosmopolitan. Occurs in all Australian winter oilseeds areas.
b
Pest status on canola: minor, restricted, irregular. Risk period: Flowering stage. Identification: Creamy white butterflies have a wingspan of 5 cm (Fig. 5.21). Larvae grow up to 3 cm and are velvety green with a pale yellow stripe along the back (Fig. 5.21). Host range: Plants in the Family Cruciferae.
Fig. 5.20. (a) Native budworm eating into a mature pod (SARDI: T. Potter) , and (b) the results of budworm feeding on green canola pods can be seen in the chewed, seedless pods at harvest (SARDI: G.J.Baker) .
Life cycle: Females lay eggs singly on leaves, which hatch into larvae. Larvae consume leaves but seldom attack floral parts of the plant when foliage is available. There are between five to seven generations per year in southern 15 3
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Fig. 5.21. Adult cabbage white butterfly. (SARDI: G.J. Baker)
Australia, able to live continuously in subtropical and tropical parts of Australia. The creamy white butterflies are usually seen flying in the crop or visiting flowers nearby, especially on warm sunny days. Caterpillars are slow moving and blend well with the foliage, out of which they eat ragged holes. Damage to canola: Larvae consume leaves, but numbers are seldom high enough to cause serious damage to a crop. They are easily controlled by a range of pesticides; there is no insecticide resistance.
Fig. 5.23. Larva of green lacewing suck through their hollow mandibles the contents of cabbage white butterfly eggs. These larvae camouflage themselves with the emptied bodies of their prey. (SARDI: G.J. Baker)
Cotesia rubecula and Pteromalus puparum (Pteromalidae) (Fig. 5.22). Native predators including green lacewings (Chrysopa sp.) are common during spring (Fig. 5.23). Pathogens have also been recorded in Australia, including fungi, the granulosis virus Bergoldia virulenta, and a bacterium. HARVEST
Natural enemies: A range of parasites have been introduced to Australia to control P. rapae, including Apanteles glomeratus (Braconidae),
Snails (Fig. 5.24) and Rutherglen bugs are frequent contaminants of harvested oilseeds. These pests may either be collected by the header as the crop is harvested or move onto the harvested crop after it is windrowed. These pests need be controlled before harvest.
Fig. 5.22. Adult wasp parasitoids (Pteromalus puparum Linnaeus Hymenoptera: Pteromalidae) emerging from the pupa of a cabbage white butterfly. (SARDI: G.J. Baker)
Fig. 5.24. Snails in harvested seed should have been controlled earlier in the season. (SARDI: T. Potter)
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SUMMER OILSEEDS B.A. Franzmann N Sunflowers, Helianthus annuus (Asteraceae). Origin: North America
Sunflowers (P.T. Bailey) PEST (major pests in bold)
PAGE
Establishment wingless cockroaches
156
black sunflower scarab
156
false wireworms
157
cutworms
158
Growth silverleaf whitefly
158
greenhouse whitefly
158
Budding, flowering black field cricket
159
Rutherglen bug, grey cluster bug and Invermay bug
159
native budworm and corn earworm
161
The average Australian area sown to sunflowers is 111 000 ha, of which 67% is grown in Qld and 32% in NSW with a little grown in WA. In Qld sunflowers are grown as dryland crops, while in southern NSW they are irrigated. In southern Qld and NSW, sunflowers are planted in spring to summer; in central Qld, planting occurs mainly in January and February; however, in all areas planting occurs whenever planting rains arrive any time between October and January in
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southern areas and January to March in northern areas. Buds are prominent about 7–9 weeks after planting and flowering commences 1–2 weeks later. Flowering (from outside to inside of the head) takes about 2 weeks. The seeds grow to full size and are fully mature after about another 4–5 weeks, and another 4–5 weeks is required for the seeds to dry down sufficiently for harvest. Planting to maturity takes about 4–5 months.
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E S TA B L I S H M E N T
Wingless cockroaches
Damage: On small seedlings, they feed on cotyledons and stems, often severing the stem. On larger seedlings, they feed on the leaves and growing points.
Blattodea: Blaberidae
Monitoring: Use germinating seed baits.
Calolampra elegans Roth & Princis, and C. solida Roth & Princis
Action level: One or more cockroaches per two germinating seed baits.
Distribution: Both species are Australian natives.
Chemical control: Baiting with insecticidetreated cracked grain baits.
Pest status: Major, recorded as pests in the central highlands of Qld, irregular. Other native cockroaches recorded as minor pests are Cosmozosteria sp. and Platyzosteria sp. Blattidae. Identification: Adult males and females of C. elegans are large (25–35 mm long) and shiny brown with yellow stripes and margins (Fig. 5.25). The male of C. solida is fully winged, whereas the females are wingless. Nymphs are initially greyish-brown or tan, developing yellow markings when about half adult size.
Cultural control: Wingless cockroach populations reach highest densities under no tillage with stubble retained. Conservation of natural enemies: No effective natural enemies have been identified.
Black sunflower scarab Pseudoheteronyx sp. Coleoptera: Scarabaeidae
Host range: Omnivorous; food includes seedlings of all field crops.
Distribution: Qld.
Life cycle on sunflowers: One generation per year.
Identification: A black beetle, about 15 mm long (Fig. 5.26).
Risk period: When seedlings are present, in late summer and autumn.
May be confused with: false wireworm adults, and the feeding damage is similar.
Pest status: Minor, restricted, irregular.
Host range: Parthenium weed is a favoured host but the larvae can develop on the roots of a number of grasses and weeds. Life cycle on sunflowers: One generation per year, with the adults damaging sunflower crops in summer.
Fig. 5.25. Adult wingless cockroach (length 3 cm). (DPI&F Qld: D. Ironside)
Fig. 5.26. Black sunflower scarab beetles (length 15 mm). (DPI&F Qld: D. Ironside)
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Risk period: Adult beetles damage seedlings and young plants in summer.
a
Damage: Adult beetles defoliate and kill plants up to 40 cm tall. They often feed in a line across the field. Monitoring: Look for feeding beetles just before sunset. Action level: Four beetles per square metre can cause severe losses to young seedlings. Chemical control: Chemicals are registered, but are of limited effectiveness.
b
Cultural control: Removal of the host parthenium weed is advised. Damage is most prevalent where sunflowers follow wheat, sorghum or grass pasture. Conservation of natural enemies: No effective natural enemies have been identified.
False wireworms Coleoptera: Tenebrionidae
Fig. 5.27. (a) Small false wireworm, Gonocephalum sp. (adult length 7 mm) (DPI&F Qld: D. Ironside) , and (b) striate false wireworm, Pterohelaeus alternatus Pascoe (adult 16 mm). Beetles in the field often have soil stuck to their backs (DPI&F Qld: D. Ironside) .
Gonocephalum spp. (including G. macleayi) Pterohelaeus spp. (including P. alternatus) N (Main entry in this chapter, page 141.)
Identification: Fig. 5.27. See also p. 141. Pest status on sunflowers: Major, widespread, irregular. Life cycle on sunflowers: Adults emerge from the soil during spring and early summer, and larvae are found from about 2 months later until the next spring. Risk period: The risk from adults is highest in summer, whereas for larvae the risk is highest for early-planted crops. Damage may occur if early plant growth is slowed by cool, damp weather, allowing larvae to remain in the moist root zone. As the soil dries, larvae retreat below the root zone. However, if crops are sown into dry seedbeds, and the soil remains dry, damage may be significant. Damage: Larvae feed on newly germinated seeds and the growing points of plants. Adults damage young plants by surface feeding or cutting off the plant at, or near, ground level and may also feed on seed in the ground.
Monitoring: Fields should be monitored before the crop is sown. The use of germinating seed baits is an effective method of detecting both adults and larvae. Action level: Decisions on the requirement for treatment are based on the numbers of insects found at germinating seed baits. Control larvae if more than 25 larvae are found around 20 germinating seed baits, and adults if more than 30 are found around 20 baits. Chemical control: Chemical control is available. For adults use cracked-grain baits and for larvae use a seed treatment or in-furrow spray. Cultural control: Prepare the ground so that germination is as even and rapid as possible. The use of press wheels at planting, which are set at no more than 2 kg cm–1 of wheel width, provides some control. Clean cultivation during summer dries out the topsoil and eliminates weeds that provide food for the adults. Adult beetles are more damaging to sunflower seedlings where stubble is buried by cultivation compared with crops direct-drilled through surface-retained stubble. 15 7
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Conservation of natural enemies: Natural enemies provide very little control.
Cutworms Agrotis spp. Lepidoptera: Noctuidae N (Main entry in Chapter 2 Cereals.)
Pest status in sunflowers: Minor, widespread, irregular. Life cycle on sunflowers: One generation per crop. Risk period: Spring and summer. Damage: Older cutworm larvae sever the stems of young seedlings at, or near, ground level, thereby causing collapse of the plant. Sometimes the young plant is partially dragged into the soil. Larvae may also climb plants and browse on or cut off leaves. Monitoring: Examine emerging seedlings twice per week and plants up to the budding stage once per week. Action level: Treat seedlings when there is a rapidly increasing area or proportion of crop damage. Treat older plants if more than 90% of plants are infested and more than 50% have 75% or more leaf tissue loss. Chemical control: Chemical control may be cost-effective. Spot spraying of identified patches may suffice. Spraying when cutworms are active in evenings may be more effective than daytime spraying, when larvae are sheltered under soil. Cultural control: Control weed growth for 3–4 weeks prior to planting. Conservation of natural enemies: Cutworms are attacked by a wide range of parasitoids, predators and diseases.
Pest status on sunflowers: Minor, restricted, irregular. May be confused with: Greenhouse whitefly. Silverleaf whitefly adults sit with their wings in a tent-like position with a slight gap between the wings, whereas greenhouse whitefly wings are flattened and slightly overlapping (Fig. 3.2). Host range: Cotton, beans, and many broadleafed weeds. Life cycle on sunflowers: The silverleaf whitefly breeds throughout the warm months on sunflowers with a life cycle of about 5–7 weeks. It has up to three generations on a sunflower crop and seven generations per year. Risk period: Summer and autumn. Damage: Adults and nymphs feed on sap from the leaves and excrete honeydew. A secondary infection develops when a black sooty mould fungus grows on the sticky honeydew. Under very heavy infestations, plants lose vigour and damage is manifest under severe moisture stress, causing leaf-wilting and failure to set seed. Chemical control: No chemical control is available. Cultural control: Removal of alternative weed hosts may be helpful, but weed-free environments may hinder the survival of natural enemies. Conservation of natural enemies: Parasitic wasps commonly provide effective biological control.
Greenhouse whitefly Trialeurodes vaporariorum (Westwood) Hemiptera: Aleyrodidae N (Main entry in Chapter 3 Cotton.)
G ROW T H
Pest statuson sunflowers: Minor, widespread, irregular. May be confused with: Silverleaf whitefly.
Silverleaf whitefly Bemisia tabaci (Gennadius) Hemiptera: Aleyrodidae N (Main entry in Chapter 3 Cotton.)
Host range: Cotton, beans, and many broadleafed weeds. Life cycle on sunflowers: The greenhouse whitefly breeds throughout the warm months on sunflowers with a life cycle of about 5–7
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OILSEEDS
Fig. 5.28. Adult Encarsia formosa wasp (2 mm long) near three parasitised greenhouse whitefly ‘pupae’. (SARDI: G.J. Baker)
weeks. It has up to three generations on a sunflower crop and seven generations per year. Risk period: Summer and autumn. Damage: Adults and nymphs feed on sap from the leaves and excrete honeydew. A secondary infection develops when a black sooty mould fungus grows on the sticky honeydew. Under very heavy infestations, plants lose vigour and damage is manifest under severe moisture stress, causing leaf-wilting and failure to set seed. Chemical control: No chemical control is available. Cultural control: Removal of alternative weed hosts may be helpful, but weed-free environments may hinder the survival of natural enemies. Conservation of natural enemies: Parasitic wasps (Encarsia formosa Gahan. Hymenoptera: Aphelinidae) (Fig. 5.28) commonly provide effective biological control.
Fig. 5.29. Two black field crickets (3 cm long) damaging sunflower head. (PIRSA: P.R. Birks)
Guillou), Lepidogryllus comparatus (Walker) and L. parvulus (Walker) (Simpson et al. 1992). Pest status on sunflowers: Minor, widespread, irregular. Host range: Soybean, mung bean and many weeds. Life cycle on sunflowers: One generation on a sunflower crop and one generation per year. Risk period: Spring and summer. Damage: Crickets feed on the leaves and stems of seedlings, sometimes severing the stem at or above ground level. They may also attack more mature plants, feeding on the back of the heads and on maturing seeds on the face of the head (Fig. 5.29). Monitoring: Count the number of insects at germinating seed baits. Action level: One or more crickets per two germinating seed baits.
B U D D I N G , F LOW E R I N G
Black field cricket Teleogryllus commodus (Walker) Orthoptera: Gryllidae N (Main entry Chapter 15 Pastures—winter rainfall.)
Other crickets implicated in damage to summer rainfall crops include Teleogryllus oceanicus (Le
Chemical control: Field crickets are controlled using insecticide-treated cracked-grain baits. Conservation of natural enemies: Natural control agents, including diseases, parasitic insects, and predatory birds and insects, appear to have little effect.
Rutherglen bug Nysius vinitor Bergroth 15 9
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Grey cluster bug Nysius clevelandensis Evans
Invermay bug Nysius turneri Evans Hemiptera: Lygaeidae Distribution: All are Australian natives. The Rutherglen bug is common in NSW, Qld, SA, Vic. and southern WA. The grey cluster bug is common in NSW and Qld and has been recorded in northern NT and WA. The Invermay bug has been recorded on sunflowers in Tas. Pest status on sunflowers: Rutherglen bug: major, widespread, irregular. Grey cluster bug: minor, widespread in northern Australia, irregular. Identification: Adults are small (about 6 mm long), dull-grey in colour, narrow-bodied and possess two pairs of silverygrey wings (Fig. 5.30). Nymphs are reddish in colour with wing buds (Fig. 5.30). Eggs are 1 mm long, and cream when laid. May be confused with: Both the Rutherglen bug and grey cluster bug may occur in the same crop. They may be separated in the field using a hand lens. The forewing of Rutherglen bug is smooth, while that of grey cluster bug is covered with small hairs (Evans 1929). These bugs may also occur with the brown mirid and apple dimpling bug. Host range: Rutherglen bug: sunflower, safflower, sorghum and many broadleaf weeds. Grey cluster bug: sunflower, safflower and other plants in the Family Asteraceae. Life cycle: The Rutherglen bug and grey cluster bug have about eight generations per year. The
duration from egg to adult is about 4 weeks in summer with the adult living a further 4 weeks. On sunflowers, there is one generation per crop. Adults and late-stage nymphs overwinter in plant debris on the soil surface and move during spring to weeds, on which eggs (up to 400 eggs per female), singly or in small groups are laid and nymphs develop. As the host plants dry during summer, bugs move to green plants, including crops. Flightless nymphs move by walking and adults by flying, generally at night, into the crop by either low-level local flights, or are carried in by winds from long distances (McDonald and Farrow 1988). Depending on time of planting of crop, adults may be present during budding and flowering and nymphs post flowering. During summer, several overlapping generations develop in which all stages may be present (McDonald and Smith 1988). Risk conditions: Winter and spring conditions favouring prolific weed growth followed by a dry late spring, forcing bugs off their host plants. Damage: Adults congregate on the stems during budding and cause the head to either wilt, become malformed or die. After flowering, adults lay eggs in flower heads and both adults and nymphs feed on the seed and reduce grain yield, oil content, oil quality and reduce seed germination. Monitoring: Count adults on buds and heads at weekly intervals. Action level: Pre-flowering heads: August– December, more than 10 adults per plant; January –April, more than 20 adults per plant. Postflowering heads: before heads turn down, August– December, more than 25 adults per plant; January–April, more than 50 adults per plant. Cultural control: Rutherglen bugs can be controlled by removing host weeds and by ploughing a deep furrow around the crop, preventing wingless bugs from migrating from weeds. Chemical control: Chemical controls may be cost-effective. Several applications may be necessary.
Fig. 5.30. Rutherglen bugs. Two dark, winged adults (6 mm) (left); three reddish-brown, wingless nymphs (right). (DPI&F Qld: D. Ironside)
Conservation of natural enemies: Egg parasitoids are very important in hindering or preventing nymph infestations and reducing bug populations.
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OILSEEDS
Native budworm and corn earworm Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Pest status on sunflowers: Minor, widespread, irregular. May be confused with: Loopers. Life cycle on sunflowers: One generation per crop. Risk period: Spring and summer. Damage: Larvae feed on the leaves, buds, petals or on the small green bracts surrounding the head. They also burrow into the back of the head or feed on the tops of the developing seeds on the face of the head. Larvae feeding on the back of the head can predispose the crop to secondary head rots. Damage is most severe when heavy
infestations coincide with budding. Feeding can cause deformation of the head and sometimes loss of the head by the larvae chewing into its connection with the stem. Monitoring: Examine plants for larvae weekly around budding. Action level: An average of more than one medium-sized larva per terminal bud at early bud formation. Damage to the developing seeds is usually of little consequence unless infestations are particularly heavy. Chemical control: Chemical control may be cost-effective; corn earworm is resistant to a number of insecticides. Conservation of natural enemies: There are very many natural enemies such as egg and larval parasitoids, predaceous insects and spiders, and various diseases.
Sources of information Allsopp, P. (1979). Identification of false wireworms (Coleoptera: Tenebrionidae) from southern Queensland and northern New South Wales. Journal of the Australian Entomological Society 18: 277–286. Baker, G. (2001). Monitoring canola crops for cabbage moth. PIRSA Fact Sheet 144/620. Berlandier, F. (1999). Managing aphids in canola. Farmnote 45/99. Agriculture WA. Broadley, R.H., Simpson, B.W. and Beavis C.H.S. (1986). Damage by Nysius spp. (Hemiptera: Lygaeidae) in non-stressed sunflower (Helianthus annus L.) crops. General and Applied Entomology 18: 17–24. Colton, R.T. (1988). Safflower growing. Agfacts. NSW Agriculture. Colton, R.T. and Sykes, J.D. (1992). Canola. Agfacts. NSW Agriculture. Cook, D., Mangano, P., Berlandier, F., Hardie, D. and Cousins, D. (2000). Managing diamond back moth in canola. Farmnote 140/2000. WA Department of Agriculture. Coutts, B.A. and Jones, R.A.C. (2004). Viruses infecting canola (Bassica napus) in south-west Australia: incidence, distribution, spread, and infection reservoir in wild radish (Raphanus raphinistrum). Australian Journal of Agricultural Research 51: 925–936. Dunn, C. and Miles, M. Insectopedia. An electronic insect pest management manual for south eastern Australian grain and pasture pests. CD. Agriculture Victoria. ISBN 0 7311 4706 5. Elder, R.J., Brough, E.L. and Beavis, C.H.S. (1992). Managing insects and mites in field crops, forage crops and pastures. Department of Primary Industries, Queensland. 143pp. Evans, J.W. (1929). A new species of Nysius (Hem: Lygaeidae) from Australia. Bulletin of Entomological Research 19: 351–354. Halliday, B. (2001). Systematics and biology of the Australian species of Balaustium von Heyden (Acari: Erythraeidae). Australian Journal of Entomology 40: 326–330.
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Hely, P.C., Passfield, G. and Gellatley, J.G. (1982). Insect Pests of Fruit and Vegetables in NSW. Inkata Press. Horne, P.A. and Edward, C.L. (1998). Effects of tillage on pest and beneficial beetles in the Wimmera region of Victoria, Australia. Australian Journal of Entomology 37: 60–63. Jones, R. and Hawkes, J.R. (2002). Yield losses caused when BWYV infects canola. Agribusiness Crop Updates. WA Department of Agriculture. Jones, R.A.C., Coutts, B.A., Smith, L.J. and Hawkes, J.R. (2003). Benefits provided by treating canola seed with imidacloprid seed dressing. Agribusiness Crop Updates. WA Department of Agriculture. Lamb, J. and Peters, C. (2001). Slugs: the next high rainfall pest. GRDC Grains Research Advice, December. McDonald, G. (1995). Wireworms and false wireworms in field crops. Agriculture Victoria Notes Series No AG0411 June, 1995. McDonald, G. and Farrow, R.A. (1988). Migration and dispersal of the Rutherglen bug, Nysius vinitor Bergroth (Hemiptera: Lygaeidae), in eastern Australia. Bulletin of Entomological Research 78: 493–509. McDonald, G. and Smith, A.M. (1988). Phenological development and seasonal distribution of the Rutherglen bug, Nysius vinitor Bergroth (Hemiptera: Lygaeidae), on various hosts in Victoria, south-eastern Australia. Bulletin of Entomological Research 78: 673–682. McQuillan, P.B., Ireson, J.E., Hill, L. and Young, C. (2006). Tasmanian pasture and forage pests. Department of Primary Industries & Water, Tasmania. Michael, P., Dore, T. and Henry, K. (2002). Bronzed field beetle attacking canola seedlings in southern Australia. Department of Agriculture, Western Australia Farmnote 32/2002. Miles, M. (1997). False wireworms: an emerging pest of canola in Victoria’s Wimmera. In: Soil invertebrates. Proceedings of the 3rd Brisbane workshop on soil invertebrates. (Allsopp, P.G, Rogers, D.J. and Robertson, L.N., eds). BSES, Brisbane. pp. 138–139. Simpson, G.B. Mayer, D.G. and Robertson, L.N. (1992). Daily trap catches of two earwig (Dermaptera) and three cricket (Orthoptera) species in Central Queensland. Journal of the Australian Entomological Society 31: 255–262. Smith, B.J. and Kershaw, R.C. (1979). Field Guide to the Non-Marine Molluscs of South Eastern Australia. Australian National University Press, Canberra.
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6 POPPIES L. Hill
(a) Flowering crop (Tasmanian Alkaloids)
(b) Maturing capsules (Tasmanian Alkaloids)
planting to 2–8 leaf rosette run up hook & flowering capsule harvest
Jul Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun
Phenology of a Tasmanian poppy crop.
PEST (major pests in bold) Planting to eight-leaf stage slugs redlegged earth mite blue oat mite rootfeeding springtail lucerne flea common garden springtail and garden springtail earwigs Rosette and run-up cutworms Flowering and capsule native budworm
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Poppies, Papaver somniferum, Papaveraceae. Origin: West Asia, Europe Area grown in Australia: 15 000 ha. All in Tasmania.
P L A N T I N G TO E I G H T- L E A F S TAG E
Reticulated slug and brown field slug Eupulmonata: Limacidae
Fig. 6.1. Poor seedling establishment and distorted leaves result from feeding by redlegged earth mite. (DPI&W Tas: J. Dennis)
Deroceras reticulatum (Müller) and D. parnormitanum (Lessona & Pollonera) N (Main entry in Chapter 5 Oilseeds.)
Pest status on poppies: Both species: major, widespread, regular occurrence. Life cycle on poppies: Both species may have life cycles of more than 1 year. Eggs may be laid in soil during autumn by slugs that have oversummered as adults. Risk period: Between emergence and eight-leaf stage.
Pest status: Earth mites are major, widespread and frequent pests of poppy seedlings. Damage: Symptoms are silvery spots on the leaf surface, distorted leaves (Fig. 6.1) and missing plants (Fig. 6.1). Damage may be first noticed in spring. During warm weather, mites may be seen active on the soil surface and mitefeeding on plants such as clover, thistle and brassica seedlings is indicative of the potential for crop damage.
Damage: Slug-feeding may kill or severely defoliate young poppy plants.
Chemical control: Control is by spraying bare earth with insecticide at planting or by foliar applications to seedlings.
Monitoring: Placement of cover such as tiles or hessian bags along the perimeter of the crop may indicate presence or absence.
Rootfeeding springtails
Action level: None determined. Chemical control: Both species may be controlled by bait pellets impregnated with a molluscicide. These are placed near seedlings where adult slugs are detected.
Onychiurus spp. Collembola: Onychiuridae. Pest status: Minor, restricted distribution in poppy fields and irregular occurrence. They primarily feed on soil fungi and in so doing may damage the roots of seedlings (Fig. 6.2).
Natural enemies: Ground beetles (Carabidae) and earwigs include slug eggs in their diet.
Redlegged earth mite and blue oat mite Acarina: Penthaleidae Halotydeus destructor (Tucker) and Penthaleus major (Dugès) N (Main entry in Chapter 13 Pastures—summer
rainfall.)
Fig. 6.2. Patchy establishment of poppy seedlings caused by rootfeeding springtails. (GlaxoSmithKline: P. Cotterill)
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POPPIES
Lucerne flea Sminthurus viridis Linnaeus Collembola: Sminthuridae N (Main entry in Chapter 15 Pastures—winter rainfall.)
Common garden springtail and garden springtail Bourletiella hortensis (Fitch), Bourletiella viridescens Stach Collembola: Bourletiellidae. Pest status: Lucerne fleas are major, widespread and frequent pests of poppy seedlings, while the often more numerous garden springtails are minor pests, widespread and frequent in occurrence. Identification: Lucerne fleas (Figs 6.4, 15.40, 15.41) are larger (to 3 mm) than garden springtails (Fig. 6.3), which grow to 1 mm long. Both are globular-shaped and jump when disturbed. Damage: Feeding by lucerne fleas damages and distorts cotyledons (Fig. 6.4), and their chewing of young leaves produces small pits that may coalesce into irregular holes in the middle and edges of seedling leaves (Fig. 6.5). Garden springtails produce similar damage (Ireson 1993).
Fig. 6.4. Common garden springtail damage to cotyledons of poppy seedling. (TIAR: J.E. Ireson)
Chemical control: Insecticides may be costeffective when leaf damage is frequent. Cultural control: Long fallows between crops and thorough cultivation following pasture reduces damage in the poppy crop that follows.
European earwigs Forficula auricularia Linnaeus Dermaptera: Forficulidae Pest status: Earwigs are major, restricted and irregular pests of poppy seedlings.
Monitoring: Distribution in a crop may be patchy, so monitoring for activity should include representative areas of the crop. Look for activity and damage between emergence and two-leaf stage.
Fig. 6.3. Top view of the common garden springtail (length 1 mm). (TIAR: J.E. Ireson)
Fig. 6.5. Lucerne flea damage to young leaves of poppy seedling. (TIAR: J.E. Ireson)
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Identification and biology: See p. 140. Monitoring: Earwigs feed on poppies during evenings and at night, when monitoring is best undertaken.
RO S E T T E A N D RU N - U P
Cutworms Lepidoptera: Noctuidae
Fig. 6.6. Newly emerged first-instar larva of native budworm (1.5 mm long) before entering the capsule. (DPI&W Tas: L. Hill)
Common cutworm, Agrotis infusa (Boisduval) Brown cutworm, Agrotis munda Walker N (Main entry in Chapter 2 Cereals.)
Pest status: Minor, widespread and irregular pests, a consequence of flights from mainland Australia. Risk period: The spring–summer generation of cutworm larvae are most likely to damage poppies during November and December, when the first signs of larval activity are leaves or whole plants cut and laying on the ground. Migrants from the mainland lay eggs on young crops and weeds (Dennis 1998). Monitoring: Since cutworm larvae feed during dusk and at nights, their presence is best confirmed by night monitoring.
F LOW E R I N G A N D C A P S U L E
Fig. 6.7. Native budworm larva emerging from poppy capsule. Larvae feed on seeds. (Glaxo SmithKline)
Monitoring: Damage potential is indicated by caterpillars on ‘indicator’ weeds such as storksbill, Erodium spp. (Geraniaceae), and by small larvae feeding on poppy foliage. Eggs may be found along the rays of the capsule. Control: For most years, control is not costeffective, but when heavy early populations are evident, insecticide application may be warranted.
Native budworm Helicoverpa punctigera (Wallengren) Lepidoptera: Noctuidae N (Main entry in Chapter 3.)
Pest status: Minor, widespread and irregular in occurrence, a consequence of flights across Bass Strait from mainland Australia. Damage: Young larvae (Fig. 6.6) of the summer generation of native budworm larvae chew the skin from pedicels causing the capsule to wither, while older larvae chew holes in the capsule from which seed falls.
Fig. 6.8. Native budworm feeding may reduce capsule size. (DPI&W Tas: J.E. Dennis)
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POPPIES
Sources of information Dennis, J. (Ed.) (1998). Field Guide to Poppy Disorders and Their Control. Tasmania. Department of Primary Industries and Fisheries. Ireson, J.E. (1993). Activity and pest status of surface feeding Collembola in Tasmanian field crops and pastures. Journal of the Australian Entomological Society 32: 155–167.
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7 PULSES—SUMMER (INCLUDING PEANUTS) H. Brier N Adzuki beans, Vigna angularis. Origin: Asia. Australian area 2000 ha. N Cowpeas, Vigna unguiculata. Origin: Africa. Australian area: 3500 ha. N Lima beans, Phaseolus lunatus. Origin: tropical America. Australian area: 600 ha. N Mung beans or green gram, Vigna radiata. Origin: India. Australian area: 40 000 ha. N Navy beans, Phaseolus vulgaris. Origin: tropical America. Australian area: 1700 ha. N Peanuts, Arachis hypogaea. Origin: tropical South America. Australian area: 30 000 ha. N Pigeon peas or red gram, Cajanus cajan. Origin: Africa. Australian area: 2000 ha. N Regur mung beans or black gram, Vigna mungo. Origin: India. Australian area: 2000 ha. N Soy beans, Glycine max. Origin: Asia. Australian area: 45–50 000 ha.
(a) Peanuts
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Summer pulse crop phenology for the main summer pulses and for different planting dates (horizontal bars). Soybean cultivars in southern Australia are indeterminate and have overlapping flowering and podding. These phenologies are based on ‘summer’ planting times. In the north Australian tropics, pulses may be grown during the dry season (southern winter). Crop Soy beans
July
Aug.
Sept.
Oct.
Nov.
Dec.
Jan.
Feb.
Mar.
Apr.
May
June
Mung beans
Adzuki beans
Navy beans
Peanuts
Crop stages: Vegetative (different shades of green for different crops) Flowering (overlaps with podding in indeterminate cultivars) Overlapping flowering and podding (indeterminate cultivars) Early podding (pod set and mid podfill, determinate cultivars) Late podding (mid podfill to harvest)
FEEDING HABIT Seedling stage soil insects
stem and leaf feeders
PEST (major pests in bold)
HOST(S)
amnemus weevil black field earwig black field cricket false wireworm peanut scarabs true wireworms whitefringed weevil beanfly
soy beans all summer pulses all summer pulses all summer pulses peanuts all summer pulses peanuts navy beans, adzuki beans, mungs mung beans (spring crops) soy beans all summer pulses soy beans soy beans, navy beans soy beans
cotton seedling thrips cotton webspinner cutworms Helicoverpa spp. silverleaf whitefly sugarcane armyworm
PAGE 218 176 175 211 208 210 218 219 206 227 242 243 176 240
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PULSES —SUMMER
FEEDING HABIT
PEST (major pests in bold)
Vegetative stage sapsuckers lucerne leafhopper mirid bugs silverleaf whitefly
stem borers
leaf-feeding beetles leaf-feeding caterpillars
Caterpillars: leaf miners, leaf rollers, webworms and web spinners
soybean aphid vegetable leafhopper beanfly lucerne crownborer redshouldered leaf beetle plague soldier beetles Anticarsia irrorata Armyworms: common, northern, sugarcane Athetis tenuis bean looper Mocis alterna castor oil looper common grass blue Ectropis despicata eight spots moth Ericeia inangulata Eubrostis sp. Helicoverpa spp. Hydrilloides lentalis Mocis trifassiata Nodaria externalis painted pine moth Pantydia spp. Scopula perlata Simplicia caeneusalis soybean looper sugarcane looper tobacco and vegetable loopers Triginoides hyppasia Utethiesa lotrix beet webworm Lecithocera sp legume webspinner
lucerne leafroller soybean leafminer soybean moth Flowering to mid-podding soil insects peanut scarabs sciarids true wireworms whitefringed weevil
HOST(S) all but most severe in peanuts all summer pulses soy beans, navy beans, peanuts soy beans all summer pulses navy beans, adzuki beans, mung beans soy beans soy beans, mung beans, navy beans soy beans all except peanuts soy beans, peanuts soy beans, peanuts all except peanuts mostly soy beans soy beans soy beans mung beans, soy beans soy beans soy beans, peanuts all summer pulses peanuts all except peanuts soy beans, peanuts soy beans, peanuts mostly soy beans soy beans, peanuts soy beans all except peanuts all except peanuts all except peanuts soy beans soy beans all except peanuts peanuts soy beans, mung beans, navy beans soy beans soy beans soy beans peanuts peanuts peanuts peanuts
PAGE 181 182 176 179 181 219 211 216 211 239 240 242 248 246 234 232 247 243 236 243 246 249 238 236 250 233 238 251 250 254 255 237 226 223 230 223 221 221 208 219 210 218
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FEEDING HABIT leaf sapsuckers
PEST (major pests in bold)
HOST(S)
Australian crop mirid brokenbacked bug brown mirid cowpea aphid
all summer pulses all summer pulses all summer pulses all, but most common in mungs all summer pulses peanuts, soy beans, navy beans peanuts soy beans, navy beans, peanuts all summer pulses all summer pulses all summer pulses all summer pulses soy beans all except peanuts soy beans, mung beans soy beans, navy beans peanuts all except peanuts all except peanuts all except peanuts mung beans, soy beans cowpeas, mung beans, navy beans all except peanuts mung beans, soy beans all except peanuts mung beans, cowpeas, soy beans mung beans, cowpeas, soy beans peanuts all summer pulses all summer pulses mung beans, adzuki beans, cowpeas all except peanuts adzuki beans, mung beans, navy beans all summer pulses peanuts peanuts, soys, mungs, adzuki beans mung beans
green mirid peanut mealybug peanut mite silverleaf whitefly
podsucking bugs
tomato and plague thrips two-spotted mite tomato and plague thrips western flower thrips Acroelytrum muricatum brown shield bug Cletus sp. crusader bug cucurbit shield bug green stink bug green vegetable bug large brown bean bug Oncocoris spp. passionvine bug redbanded shield bug ricespotting bugs
seed feeding bugs
small brown bean bug Graptostethus servus Melanerythrus mactans
pod, flower and leaf eaters
peanut trash bug Rutherglen bug flower thrips Apion sp. beetle bean flower caterpillar bean podborer cluster caterpillar Endotricha puncticostalis etiella
Eublemma dimidialis
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243
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PULSES —SUMMER
FEEDING HABIT pod, flower and leaf eaters
PEST (major pests in bold)
HOST(S)
Gymnoscelis lophopus
soy beans, mungs, navy beans all summer pulses mungs, adzuki beans, navy beans all except peanuts all except peanuts soy beans, mungs, navy beans adzuki beans mungs, adzuki beans, navy beans soy beans soy beans, mungs, navy beans soy beans, peanuts mungs, adzuki beans, navy beans
Helicoverpa spp. Mocis spp. loopers pale pea blue pea blue redshouldered leaf beetle sorghum head caterpillar soybean loopers soybean podfly Sphenarches spp. tussock moths tobacco and vegetable loopers
Late podding to pod ripening soil insects scarab beetles whitefringed weevil podsucking brown bean bugs bugsU brown shield bug green vegetable bug redbanded shield bug pod and seed Bruchidius mackenziei eaters Etiella
Helicoverpa spp.% lucerne crownborer Post-harvest seed damageV
bean bruchid Bruchidius mackenziei cowpea bruchid Etiella #
PAGE 233 243 248 235 236 216 231 251 220 223 236 254
soy beans soy beans all except peanuts all except peanuts all except peanuts all except peanuts soy beans peanuts, soy beans, mung beans, adzuki beans all except peanuts soy beans
208 218 193 196 198 203 214 228
navy beans, lima beans soy beans cowpeas, mung beans, soy beans peanuts, soy beans, mung beans, adzuki beans
213
243 211
214 214 228
% Only late-instar Helicoverpa larvae present by this stage. Crops are no longer attractive to ovipositing moths. U Podsucking bug populations typically peak during late podding. Pods are at risk until very close to harvest. V Unlike Bruchidius mackenziei, cowpeas and bean bruchids are usually not detected until well into storage (2–3 months). However, all species can infest crops in the field prior to harvest. # Etiella does not breed in storage but live larvae often emerge from pods soon after harvest.
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Summer pulses are presently grown over a smaller area than winter pulses but they support a more diverse arthropod fauna and the number of pest species is greater. The potential for expansion of summer pulse crops further into tropical and subtropical areas of northern Australia suggests that management of these pests needs be anticipated. Summer pulses are frost-sensitive but it is possible to grow them as winter crops in northern Australia. Here, summer legume pests are active throughout the ‘winter’ or dry season. Most summer pulses in Australia are grown as dryland crops, but areas under irrigation are increasing to stabilise production.
Mites and insects on summer pulses
Peanuts suffer less from insect pests than other summer pulses. However, peanuts are susceptible to insect damage to foliage and to peanut pegs (connecting the underground pods to the plant’s branches). Depending on soil and locality, peanut pods may be attacked by Coleoptera and some Lepidoptera. Insect damage increases the risk of aflatoxin production by Aspergillus fungi.
Distribution: Found in most Qld peanut growing regions.
Soy beans are relatively more susceptible to foliage-feeding pests than the other summer pulses because their leaves are more attractive to caterpillars and their pods have thicker, less succulent walls than mung bean and navy bean pods. Soy beans can compensate for considerable insect damage during early podding as they set a large number of ‘reserve’ pods.
In the table on pages 170–173, entries are arranged in alphabetical order of common name (where these exist) within their feeding habit. In the text following, entries are arranged by Order and Family. Within families, entries are generally arranged in alphabetical order.
M I T E S ( AC A R I N A )
Peanut mite Paraplonobia sp. Acarina: Tetranychidae
Pest status: Minor, widespread in peanut crops, irregular. Identification: A relatively large mite (1.5 mm long) with a very dark green body and pale legs. These mites drop off leaves if disturbed. Leaffeeding by peanut mites is characterised by fine silver stippling. May be confused with: Similar to two-spotted mite but much larger. Peanut mite damage to leaves (stippling) may be confused with stippling caused by vegetable jassids but is much finer (Fig. 7.1).
Mung beans, adzuki beans and navy beans share similar pests to soy beans but in addition they experience beanfly attack during the seedling stage, and are very susceptible to bean podborer. Pesticide resistance in helicoverpa, the risk of flaring silverleaf whitefly and two-spotted mites, market pressure for lower pesticide residues and de-registration of older pesticides have all increased the adoption of integrated pest management (IPM) in Australian pulses. While ‘new’ and effective pesticides and biopesticides with IPM compatibility have recently been registered in pulses against caterpillar pests, the quest continues for truly effective but ‘soft’ options for bug pests in pulses.
Fig. 7.1. Stippling on a peanut leaf caused by peanut mite. (DPI&F Qld: J. Wessels)
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the adoption of IPM. Often a major pest of summer pulses in glasshouses. Risk conditions: The two-spotted mite (TSM) is most abundant in crops where non-selective ‘hard’ pesticides are extensively used against other pests. Proximity to other susceptible crops and extended periods of hot dry weather increase the risk of damage.
Fig. 7.2. Peanut mite damage near Kingaroy, Qld. Leaves of heavily infested plants are bleached. (H. Brier)
Host range: Peanuts. Risk conditions: During periods of extended hot dry weather. Populations crash after heavy rain. Damage: Peanut mite-feeding causes a yellowing and silvering of leaves. Damage by high populations results in the shedding of lower leaves and eventual plant death (Fig. 7.2). Monitoring: Check the underside of leaves for symptomatic leaf discolouration. Mites drop from leaves at the slightest disturbance. Action level: Take action if more than 30% of plants are showing damage symptoms. Chemical control: This species is readily controlled with registered pesticides. Cultural control: Regular irrigations reduce the risk of peanut mite damage. Conservation of natural enemies: Apple dimpling bug, lacewings, ladybirds, smudge bugs and predatory thrips are key mite predators.
Damage: Mites make fine webbing on the underside of the leaves, and feed by rasping and sucking plant tissue. Infested leaves appear speckled. In severe cases the leaves turn a yellow–brown before they wither and drop from the plant. Heavy infestations during flowering and early pod formation result in early leaf senescence and may significantly reduce seed size and yield. Yield loss from mites can be as high as 30%, with late-maturing, longer-season types most at risk. Monitoring: Check the lower surface of older leaves with a hand lens. Early infestations may be patchy. Action level: Indicative level of 30% of plants are infested. Chemical control: TSM is resistant to most current pesticides in pulses. The only effective miticide currently registered (as at mid 2007) in Australian pulses is abamectin, which is only registered in soybeans. Cultural control: Avoid successive plantings, which allow a build-up of mites. Proximity to earlier maturing hosts, particularly cotton, is a major risk factor. Conservation of natural enemies: Apple dimpling bug, lacewings, ladybirds, smudge bugs and predatory thrips are key mite predators. Conserve by using ‘soft’ options where possible against other pulse pests.
Two-spotted mite Tetranychus urticae Koch Acarina: Tetranychidae Also known as red spider mite or two-spotted spider mite. N (Main entry in Chapter 3 Cotton.)
Pest status on summer pulses: Major, widespread, and regular in some regions but decreasing in importance in many regions with
CRICKETS ( ORTHOPTER A)
Black field cricket Teleogryllus commodus (Walker) Orthoptera: Gryllidae N (Main entry in Chapter 13 Pastures—summer
rainfall.)
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Distribution: Widespread in Australia, common in cracking soils. Pest status: Minor, widespread, irregular. Identification: See Chapter 13. Host range: Many field crops, including most pulses. Risk conditions: Crops can be attacked at any stage. Crops in heavier soils are at greatest risk. Most damage is caused by crickets already in the crop area at planting or by adults flying into crops. Damage: Significant damage may be caused by adults and large nymphs feeding on leaves, stems and pods. When black field crickets are present in plague numbers, seedling crops can be thinned to the point where replanting is necessary. At podding, adults chew into pods to reach the seeds. Monitoring: Crickets feed at night, so inspect crops at dusk when crickets are most active. Black field cricket activity can also be monitored with light traps. Action level: Take action if significant damage or significant cricket populations are present.
Fig. 7.3. Black field earwig damage to peanut pods, Katherine NT, showing small entry hole. (NTDPI: M. Hoskins)
Risk period: Seedling stage. Peanuts are also at risk during podding. Damage: Black field earwigs usually feed on decaying stubble, but may attack newly sown and germinating seeds and seedlings. Seedlings damaged by black field earwigs often have their tap root ringbarked leading to plant death. Black field earwigs also attack peanuts, boring numerous holes in the pods (Fig. 7.3).
Chemical control: Check for the latest Australian Pesticides and Veterinary Medicines Authority (APVMA) registrations.
Monitoring: Look for adult activity in the soil at the time of planting. Seedling deaths may indicate earwig attack. Place moistened bags on the soil surface and inspect after 1–2 days for earwigs sheltering underneath.
Cultural control: Weedy cultivation prior to planting may encourage crickets.
Action level: Take action if earwigs are readily noticeable when preparing land for planting. Chemical control: Check for the latest APVMA registrations.
E ARWIGS ( DER M APTER A)
Black field earwig Nala lividipes (Dufour) Dermaptera: Labiduridae N (Main entry in Chapter 4 Maize.)
Pest status on summer pulses: Mostly minor, except in favoured soil types where it can cause significant damage; widespread. Identification: See p. 122. Host range: A polyphagous pest that can attack seedling pulses including soy beans, mung beans, navy beans and peanuts.
Cultural control: Avoid paddocks with a previous earwig history and avoid successive plantings of susceptible crops. There have been recent earwig outbreaks in peanuts in the NT in paddocks previously planted to cotton. Planting peanuts in lighter soils will also reduce the risk of attack.
W H I T E F L I E S ( A L E Y RO D I DA E )
Silverleaf whitefly Bemisia tabaci (Gennadius) biotype B Hemiptera: Aleyrodidae
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Frequently abbreviated to SLW and also known as the poinsettia whitefly. N (Main entry in Chapter 3 Cotton.)
Distribution, identification and life cycle: See Chapter 3. Pest status on summer pulses: Major, widespread, regular. SLW has the potential to invade summer pulse crops in extremely large numbers and rapidly increase during summer months. Host range: See Chapter 3. Of the summer pulses, soy beans and navy beans are preferred SLW hosts; however, in SLW plague years the pest may spill over into less-preferred legume hosts such as peanuts. Significant populations of SLW adults are frequently seen in mung beans but nymphal development on this crop is very poor. Life cycle: See Chapter 3. Risk period: Summer and autumn. Crops remain attractive to SLW until mid podfill. Once photosynthetic assimilates are redirected from leaves to fill the pods, leaves become unattractive to SLW and adults leave the crop to find more attractive hosts. Damage: SLW can reduce plant vigour and yield by removing plant photosynthate from the leaves. Severe infestations in young plants can stunt plant growth and greatly reduce a crop’s yield potential. Later infestations can reduce the number of pods set, seed size, and seed size uniformity, thus reducing yield and quality. As a rule, the impact of SLW is worst in droughtstressed crops. Pod and seed discolouration (Fig. 7.4) caused by SLW may reduce
Fig. 7.4. Pale pods symptomatic of silverleaf whitefly attack in soy beans at Coominya, Qld. (H. Brier)
Fig. 7.5. Severe sooty mould in soy beans heavily infested with silverleaf whitefly, Grafton, NSW. (N. Moore)
marketability, where vegetable soy beans (edamame), green beans and field peas are picked for the fresh market. While SLW are capable of transmitting a number of geminiviruses that infect soy beans, none have yet been detected in Australia. SLW secrete large amounts of sticky honeydew. Adult females produce more honeydew than other stages and nymphs produce more honeydew when feeding on stressed plants. The sooty mould that develops on honeydew reduces photosynthesis (Fig. 7.5). Sooty mould damage is greatest during early to mid podfill when SLW activity is greatest at the top of the canopy. Rain and irrigation wash honeydew off leaves, lessening the risk of sooty mould. Monitoring: As SLW eggs, nymphs and resting adults are found on the underside of leaves, leaves must be turned over to assess SLW activity (Fig. 7.6). Flying SLW adults are readily
Fig. 7.6. Silverleaf whitefly eggs on underside of a leaf of weed host. (CSIRO)
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observed when crops with high populations are disturbed. The presence of honeydew and sooty mould may also indicate SLW attack, but can be due to aphid-feeding.
summer pulses in Australia. SLW has resistance to most pesticides registered against other pests in summer pulses. Chemical control is not presently cost-effective.
SLW eggs are laid on young leaves; however, the production of new growth in vegetative crops means that by the time eggs develop to large nymphs, leaves with the greatest visible SLW nymphal activity may be five to seven nodes below the top node. As vegetative growth slows, however, plant nodes with greatest nymphal activity move progressively upwards to the canopy top.
Cultural control: Where possible, avoid successive plantings of summer pulses and avoid planting summer pulses in close proximity to earlier maturing SLW hosts such as cotton and cucurbits. Where damaging SLW populations are evident in other crops early in summer, or in regions with a history of damaging widespread SLW activity, consider planting mung beans or adzuki beans (Vigna sp.), which are less attractive to SLW, rather than a high-risk types such as soy beans. Control SLW weed hosts such as rattlepod and sowthistle. Irrigate crops to reduce moisture. Overhead irrigation also washes off sooty mould and drowns adult SLW. Narrow-leafed and smooth-leafed (less hairy) cultivars may be less attractive to SLW, but hairless leaves may predispose crops to aphid attack.
The following sampling guidelines are the best available to date: sample leaves from the nodes with the greatest SLW activity. Move at least 10 rows into the crop. Examine a leaf from each of 15 separate plants from each of at least two widely separated locations in a row, a total of 30 leaves. Turn the central leaflet over slowly so as to minimise disturbance of SLW adults and record the density of adults, then count the nymphs (Fig. 7.7) by placing a sampling card with a 4 cm2 circular hole (10-cent sized) over the central part of each leaflet and count the number of nymphs in the sampling hole, Be careful not to count empty pupal skins (Fig. 7.7). Action level: There are no validated SLW thresholds in summer pulses; well-watered soy beans can tolerate nymphal populations of five per square centimetre on the leaves at the peak activity nodes, but densities of 15–20 per square centimentre on peak node leaflets have a marked effect on yield and pod colour. Chemical control: There are no pesticides specifically registered for SLW control in
Conservation of natural enemies: SLW nymphs are parasitised by native species of Encarsia spp. and Eretmoceros spp. (Aphelinidae). In 2005 CSIRO first released the exotic parasite Eretmoceros hayati in south-eastern Qld. This species has successfully established in release areas, and together with native parasites, has seemingly stabilised SLW populations. Parasitism by Eretmocerus sp. in excess of 80% was observed in SLW at Bundaberg in 2007 with very little SLW activity evident in most soybean crops.
A PH I D S ( A PH I D I DA E )
Cowpea aphid Aphis craccivora Koch Hemiptera: Aphididae Distribution: A cosmopolitan species found throughout Australia. Heavy infestations have been observed in the South Burnett region of south-east Qld. Pest status: Moderate, widespread, irregular. Fig. 7.7. Silverleaf whitefly redeye pupa on peanut leaves together with empty white skins from the previous moult. (DPI&F Qld; J. Wessels)
Identification: Cowpea aphids are relatively small (up to 2.5 mm long). Adults are shiny black, while the nymphs are slate grey (Fig. 8.4).
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May be confused with: Cowpea aphid is readily distinguishable from other aphids in summer pulses by its black colouration. Brown smudge bug nymphs look superficially like cowpea aphid nymphs. Host range: A polyphagous pest attacking many grain legumes, lucerne, cotton and lettuce. Mung beans (and presumably cowpeas) are particularly attractive hosts. Life cycle: Early aphid colonisation may be overlooked but numbers can explode from flowering onwards in warm weather. Females can reproduce asexually producing over 80 nymphs per female. In summer, nymphs can complete their development in 5–7 days. High aphid populations can smother stems, pods and leaves. Risk period: Flowering onwards. Later-planted crops are at greater risk than spring-planted crops. Damage: Cowpea aphids inject toxins into the plant while feeding. Cowpea aphids most likely reduce mung bean vigour and yields. Aphidfeeding also produces honeydew, which grows sooty mould that reduces photosynthesis and makes harvesting difficult. Monitoring: The presence of ladybirds is an indication that aphids are present. Look for aphid colonies on plant stems by parting the canopy. Heavy infestations will become readily visible when they spread to the upper leaves and pods. Action level: There are no set thresholds for cowpea aphid in mung beans.
Fig. 7.8. Soybean aphids up to 2 mm long. Aphids have a bright green body, black siphunculi and pale cauda. (J. Hughes)
recently USA, Canada and Australia. In Australia, first reported in the Northern Rivers district of NSW and is now most abundant in coastal NSW and Qld and has been reported from all major production areas. Pest status: Moderate to major, widespread and regular. Populations have stabilised somewhat since the pest’s first arrival in Australia. Identification: The soybean aphid is a small bright green aphid (up to 2 mm long), with black siphunculi and pale cauda (Fig. 7.8). No other aphid on soy beans has the same size and colour combination. May be confused with: Cotton aphid (Aphis gossypii), distinguished by a shorter, dark cauda, and the cowpea aphid (Aphis craccivora) that has a dark body.
Cultural control: Not applicable.
Host range: Mainly soybean and other Glycine species. In Asia, the soybean aphid alternates between Rhamnus species (Family Rhamnaceae) and host legumes. It has also been recorded from Pueraria and Desmodium species.
Conservation of natural enemies: Pesticides that kill aphid predators (ladybirds, predatory bugs and hoverfly larvae) should be avoided.
Life cycle: Aphid densities peak between flowering to early podding and decline rapidly thereafter.
Chemical control: This pest is easily controlled with systemic pesticides.
Soybean aphid Aphis glycines Matsumura Hemiptera: Aphididae Distribution: South-East Asia, China, Japan, far eastern Russia, Korea, the Philippines, and most
Risk period: Late vegetative, flowering and early podding stages. Damage: Soybean aphids can reduce plant vigour, height and yield (the latter by up to 30% in Chinese studies). Heavily infested plants may be covered in sooty mould growing on honeydew secreted by the aphids. This further 17 9
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reduces photosynthesis. Infested plants may also have distorted leaves. In Asia, the soybean aphid is a known vector of a number of plant virus diseases, including abaca mosaic, soybean mosaic, soybean stunt, beet mosaic, millet red leaf, mung bean mosaic, bean yellow mosaic and Indonesian soybean dwarf. To date, no virus transmission by soybean aphid has been observed in Australia. Monitoring: Look for aphid colonies on the upper stems, leaflets and terminal leaves. In heavily infested crops, cast off aphid skins, sooty mould, and large ladybird populations are indicative of soybean aphids; however, the latter two symptoms are also indicative of significant whitefly populations. Chemical control: Soybean aphids are controlled with systemic pesticides but no products are specifically registered for their control in soy beans. Cultural control: Avoid water stress to plants. Conservation of natural enemies: Hoverfly larvae (Syrphidae) and ladybird larvae (Coccinellidae) play an important role in suppression of A. glycines populations in soy beans. In Asia, several species of parasitic wasps are important soybean aphid biocontrol agents but these are not presently in Australia. Avoid using non-selective pesticides against other pests that also harm aphid predators.
mealybugs are usually obscured by short, white, waxy filaments and form colonies on the lower and underground parts of host plants. Adult males are smaller than females and are reddishbrown with one pair of wings and two long waxy ‘tails’. When adults are crushed they exude pink body fluids. May be confused with: Peanut mealybugs are easily confused with larvae of Cryptolaemus montrouzieri (Fig. 3.48), which are also covered with white waxy filaments. Cryptolaemus is an important mealybug predator and should not be accidentally destroyed. Host range: A polyphagous pest attacking a wide range of crops and ornamentals. In Australia, summer pulses attacked include peanuts, soy beans, navy beans and pigeon peas. The pest is most frequently encountered in the first two mentioned crops. Other crops attacked include cotton, cucurbits, avocados, cucurbits, mangoes, and citrus. Life cycle: Female mealybugs die shortly after laying their eggs in egg sacs. Freshly laid eggs are orange but turn to pink before hatching. First-instar nymphs (crawlers) disperse by walking and by wind, and can travel considerable distances to find a suitable host. The life cycle takes about 23 to 30 days. The peanut mealybug lays up to 600 eggs and can produce up to 15 generations per year. They overwinter as eggs in the soil and in leaf trash. Risk period: Young plants are at greatest risk.
M E A LY B U G S ( P S EU D O C O C C I DA E )
Peanut mealybug Maconellicoccus hirsutus (Green) Hemiptera: Pseudococcidae Also known as the pink mealybug or the hibiscus mealybug. Distribution: Recorded worldwide in tropical and subtropical regions, including Africa, SouthEast Asia, the Caribbean (1994), Florida, USA (2002) and northern Australia.
Damage: On peanuts, mealybugs can feed on the underground parts of the roots, pods, and pegs of the plant, resulting in stunted growth and poorly developed pods. Heavily infested pods can collapse and turn black. Heavy mealy bug infestations may reduce the vigour of other hosts, such as soy beans, and have been observed feeding on both stems and roots. Monitoring: Look for the distinctive adults and nymphs on the lower stems (at or below ground level), pegs and pods.
Pest status: Minor, restricted, irregular.
Action level and chemical control: No set thresholds as chemical control is not feasible.
Identification: Adult mealybugs are small (about 3 mm long) with a pink body colour covered in a waxy secretion (Fig. 7.9). Female
Cultural control: Ensure paddocks are welldrained as infestations are heaviest in poorly drained soils.
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Fig. 7.9. Peanut mealybug on soybean stem, Childers, Qld. (J. Wessels)
Fig. 7.11. Peanut crop with severe hopper burn symptoms (yellowing and necrosis of leaf tips). (H. Brier)
May be confused with: Vegetable leafhopper. Conservation of natural enemies: The ladybird Cryptolaemus montrouzieri is an important predator. Other natural enemies include lacewings and parasitic wasps (Leptomastix sp.).
L E A F H O P P E R S ( C I C A D E L L I DA E )
Host range: All summer pulses including peanuts. Also in lucerne. Damage symptoms are most severe in peanuts. Life cycle: Eggs are laid in slits made in soft plant tissue. In hot weather, a life cycle can be completed in less than 2 weeks. This species is usually not as abundant in summer pulses as the vegetable leafhopper.
Lucerne leafhopper (jassid)
Risk period: Crops are at risk at any stage.
Austroasca alfalfae (Evans) Homoptera: Cicadellidae
Damage: Lucerne leafhoppers are phloemfeeders. Toxins injected by this species cause hopper burn (Fig. 7.11), a yellowing and then a necrosis (burning or browning) of the leaf tips. Peanuts in particular are sensitive to this damage. Severely damaged crops appear to be in full bloom, the yellowing leaves being mistaken for flowers from a distance.
Distribution: Native to Australia and found in all states. Pest status: Moderate, widespread, irregular. Identification: Similar in size (3 mm long) and appearance to vegetable leafhoppers but yellow– green (as opposed to blue–green) in colour (Fig. 7.10).
Monitoring: As for vegetable leafhoppers. Also, look for the distinctive hopper burn symptoms. Action level: Spray if more than 20% of peanut leaves have hopper burn symptoms. Chemical control: Lucerne leafhoppers are easily controlled with systemic pesticides. Cultural control and conservation of natural enemies: As for vegetable leafhoppers.
Vegetable leafhopper (jassid) Austroasca viridigrisea (Paoli) Homoptera: Cicadellidae Fig. 7.10. Lucerne leafhopper (3 mm long). (DPI&F Qld)
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Pest status: Minor, widespread, regular. Identification: Adults are 3 mm long and a bright slightly bluish-green. They have a broad rounded head and a tapering body with clear wings extending beyond the abdomen. Their eyes are widely separated and the antennae short. Nymphs are similar in shape but are smaller and are wingless. May be confused with: The white spotting on the vertex and front half of the pronotum enables this species to be readily differentiated from other Austroasca species in Australia. The cotton leafhopper Amrasca terraereginae also has white spotting on the head but also bears a brown (sometimes yellowish) spot on the tegmen, which is lacking in A. viridigrisea. The lucerne leafhopper, Austroasca alfalfae, is similar in outline and size but is yellowish green in colour. Mirid nymphs are easily distinguished by their long antennae. Host range: All summer pulses including peanuts and also leafy vegetables, carrots, potatoes and tomatoes. Life cycle: Eggs are laid in slits made in soft plant tissue. During hot weather, a life cycle can be completed in less than 2 weeks. Huge populations (q 100 per square metre) are frequently encountered in summer pulses. Risk conditions: Drought-stressed poorly growing crops.
Cultural control: Well-watered, vigorously growing crops can tolerate damage. Conservation of natural enemies: Generalist predators such as predatory bugs and spiders will attack jassids. Unnecessary sprays for leafhoppers will adversely affect these and other beneficial insects and may flare other pests such as Helicoverpa.
M I R I D S ( M I R I DA E )
Australian crop mirid Sidnia kingbergii (Stål) Hemiptera: Miridae Eurystylus australis Poppius is a junior synonym. Distribution: Australia and New Zealand. Reported from all Australian states but not (as yet) the NT. Pest status: Minor, widespread, irregular. Description: The nymphs (Fig. 7.12) are similar to brown mirid nymphs, being green with striped antennae. However, they are stouter than brown mirid nymphs, have relatively shorter antennae and have a small dark spot on their back. Adults are 4–5 mm long and have a shorter (more truncated) body than green mirids (Fig. 7.13). They vary in colour, ranging (dorsally) from mid-green to dark grey–green to reddish
Damage: Vegetable leafhoppers are xylemfeeders. Vegetable leafhopper feeding kills leaf cells and results in small white dots or stippling on the leaves. Trials in drought-stressed peanuts and navy beans have shown that even high vegetable leafhopper populations have little to no effect on yield in these crops. Monitoring: Leafhoppers are most easily sampled with a sweep net. Empty captured jassids into a container with 70% alcohol (or methylated spirits), and express counts as lefhoppers per sweep (one sweep per row). Action level: 25 vegetable leafhoppers per sweep over a single row (with 90–100 m row spacing). Trials in drought-stressed peanuts and navy beans suggest control is rarely warranted. Chemical control: Vegetable leafhoppers are easily controlled with systemic pesticides.
Fig. 7.12. Australian crop mirid final-instar nymph (4 mm), from mung beans, Kingaroy, Qld. (J. Wessels)
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parasitising Australian crop mirid eggs in New Zealand.
Brokenbacked bug Taylorilygus pallidulus (Blanchard) Hemiptera: Miridae Lygus apicalis is a junior synonym of T. pallidulus. Also referred to as Taylorilygus apicalis (Fieber) in some recent publications.
Fig. 7.13. Australian crop mirid (4 mm) from mung beans, Kingaroy, Qld. (DPI&F Qld: J. Wessels)
brown, often with dark spotting or mottling. Some specimens have a pronounced dark ‘V’ on their back pointing backwards from the shoulders. Australian crop mirid adults are bright green underneath. May be confused with: Green mirid (Creontiades dilutus), brown mirid (Creontiades pacificus), and brokenbacked bug (Taylorilygus pallidulus). Hosts: A polyphagous insect attacking mung beans, adzuki beans, and navy beans. Also reported in asparagus, carrots, clover, lucerne, passionfruit, strawberries and wheat. Life cycle: Five nymphal instars. The pale yellow eggs are 0.9 mm long and are inserted into host tissue. Risk period: Budding, flowering and early podding. Crop mirids are reported as preferring hot dry conditions. Damage: Similar to that caused by green and brown mirids, and as least as damaging as these species; however, rarely the dominant mirid species in pulse crops.
Distribution: A cosmopolitan pest recorded from Africa, Asia, Europe, North and South America, New Zealand and Australia. In Australia, it is recorded from NSW, Qld and SA, but most likely present throughout Australia. Pest status in pulses: minor, widespread, irregular. Identification: The nymphs are a pale yellow– green with pale antennae (Fig. 7.14) that are much shorter than those of green mirid. They are stouter than green mirid nymphs and are closer in outline to apple dimpling bug nymphs, having (proportionally) a much wider head than green mirid nymphs. Nymphs are larger and greener than those of apple dimpling bug (Campylomma liebknechti). Adults are smaller (4–5 mm long) but stouter than green mirids and are light green with brown flecks on the inner wings, with the outer wings (corium) predominantly brown. Wing tips are bent down at 45° giving the distinctive ‘broken back’ appearance (Fig. 7.15), a characteristic shared with the brown smudge bug (which is brown all over). May be confused with: Adult green mirid (Creontiades dilutus), brown mirid (C. pacificus), brown smudge bug (Deraeocoris signatus) and
Monitoring and action levels: As for green mirids. Chemical control: As for green mirids. Chemicals with little residual activity give poor control of nymphs hatching from eggs laid prior to spraying. Cultural control: As for green mirids. Conservation of natural enemies: As for green mirids. Chalcid wasps have been reported as
Fig. 7.14. Large brokenbacked bug nymph, Kingaroy, Qld (3.5 mm). (DPI&F Qld: J. Wessels)
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Fig. 7.15. Brokenbacked bug adult (5 mm). (DPI&F Qld:
Fig. 7.16. Brown mirid third-instar nymph (2.5 mm). (DPI&F
J. Wessels)
Qld: J. Wessels)
crop mirid (Sidnia kingbergii), and nymphs of apple dimpling bug which are smaller and are pale yellow rather than green.
Pest status: Major, widespread, regular. Mixed populations of green and brown mirids are common in many crops, and on occasions brown mirids are the dominant species.
Host range: A polyphagous insect, attacking mung beans and adzuki beans as well as avocados, commercial and wild Compositae, cotton, lettuce, lucerne, sunflowers, safflower and pine seedlings. While very common in Australian sunflowers, brokenbacked bug is less common in mung beans. It is thought to prey on lepidopteran eggs and small larvae. Life cycle: There are probably five nymphal stages. Risk period: Budding, flowering and early podding. Present throughout summer. Damage: Brokenbacked bugs attack mung bean buds, flowers and small pods causing them to abort and may be more damaging than green mirids. Monitoring, action levels, chemical control, cultural control and conservation of natural enemies: As for green mirids.
Identification: Brown mirid nymphs are similar to those of the green mirid. Brown mirid nymphs may be distinguished by the distinctive reddish (brown) and white banding of the antennae (Fig. 7.16), a characteristic they share with nymphs of the crop mirid, Sidnia kingbergii. However, nymphs of the latter species have shorter antennae and a stouter body than brown mirid nymphs, and also have a small dark spot in the middle of their back. The brown mirid adult is very similar in shape and size (6–7 mm long) to the green mirid but is slightly more linear and elongated. Adults have two distinct colour forms, the brown form being predominantly light brown with darker pigmentation on the hind legs (Fig. 7.17), and the green form being mostly bright green but with dark red (or purple–brown) pigmentation of the head, pronotum, scutellum and hind coxae.
Brown mirid Creontiades pacificus (Stål). Hemiptera: Miridae Creontiades pallidifer (Walker) is a synonym for C. pacificus. Distribution: Eastern Asia including China and Micronesia, where it is recorded as C. palildifer, in Tahiti (as C. pacificus), and in Australia (where formerly referred to as C. pallidifer). In Australia, brown mirids have been recorded from northern NSW and Qld.
Fig. 7.17. Brown mirid adult, brown form (7 mm). (DPI&F Qld: J. Wessels)
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Adults have dense setae on their scutellum and pronotum. The first antennal segment is as long as, or slightly wider than, the head width across the eyes. May be confused with: Adults may be confused with green mirid (Creontiades dilutus), brokenbacked bug (Taylorilygus pallidulus), Australian crop mirid (Sidnia kingbergii), and damsel bugs (Nabis kingbergii). Nymphs may be confused with nymphs of all the above species, as well as with those of the small predatory mirid Tytthus chiensis. Hosts: A polyphagous insect attacking a wide range of crops including pulses (mung beans, navy beans, soy beans, and peanuts), cotton, sunflower, and lucerne. It is likely that brown mirids share all the summer pulse hosts of green mirid hosts. Life cycle on grain legumes: The life cycle on grain legumes is similar to that of the green mirid, with five nymphal instars. As for the green mirid, brown mirid populations can increase very rapidly in flowering crops, and the majority of brown mirids present in crops are often nymphs. Risk period: Budding, flowering and early podding. High populations can also cause significant damage to larger pods. Damage: Brown mirids cause damage similar to that of green mirids. They attack buds, flowers and small pods causing them to abort. Damage to larger pods reduces seed size and quality. Brown and green mirids are equally damaging to mung beans and nymphs are as damaging as adults.
Fig. 7.18. Green mirid eggs inserted into mung bean pod. (DPI&F Qld: J. Wessels)
Identification: Green mirid eggs are pale and elongated and inserted in plant tissue (Fig. 7.18). The nymphs are elliptical in shape and lack wings; however, wing buds are evident on finalinstar nymphs (Fig. 7.19). Young nymphs have antennae much longer than their body. Firstinstar nymphs are pale brown–orange in colour but later instars are pale green. Green mirid nymphs have pale antennae, usually with no distinct banding, but sometimes with banding on their outer (distal) end. The green mirid adult is an elongated pale green bug 6–7 mm long with long legs (especially the hind legs) and long antennae (Fig. 7.20). Some specimens may have reddish flecking on the body and the legs. Adults have sparse setae on their scutellum and pronotum. The first antennal segment is distinctly shorter than the head width across the eyes.
Monitoring, action level, chemical and cultural control and natural enemies: As for green mirids.
Green mirid Creontiades dilutus (Stål) Hemiptera: Miridae Formerly classified as Megacoelum modestum Distant. Distribution: Endemic to and widespread throughout Australia, including Tas. Pest status on summer pulses: Major, widespread, regular.
Fig. 7.19. Green mirid fifth-instar nymph (5 mm). (DPI&F Qld: J. Wessels)
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but a small percentage of nymphs may complete their development in 4 or 6 instars. Mirid development (egg to adult) is very rapid at higher temperatures, taking only 15 days at a mean temperature of 31°C (e.g. minimum of 22°C and maximum of 40°C); however, at a mean temperature of 22°C (minimum of 15°C and maximum of 29°C) development time is extended to 29 days. Egg and nymphal development times at 31°C and 22°C are 10 and 19 days and 6 and 9 days, respectively. On average, egg development is 37% of the total development time. Development times are based on mirids reared on green beans. Fig. 7.20. Adult green mirid (7 mm). (DPI&F Qld: J. Wessels)
May be confused with: Adults may be confused with brown mirid (Creontiades pacificus), brokenbacked bug (Taylorilygus pallidulus), and Australian crop mirid (Sidnia kingbergi). Nymphs may be confused with nymphs of all the above species, as well as with those of the small predatory mirid Tytthus chiensis (Fig. 7.23). Nymphs of the latter species are small (a 3 mm long, maximum) and green but are more elongated, have shorter antennae (in relation to their body), and have a more prostrate posture than Creontiades nymphs. Hosts: The green mirid is a polyphagous insect attacking a wide range of legumes, including adzuki beans, cowpeas, lima beans, mung beans, navy beans, peanuts, pigeon peas, and soy beans (summer pulses), and field peas, lupins and faba beans (winter pulses). It is also recorded from many other field crops (cotton, sunflower), horticultural crops (e.g. apples, cucurbits, tomatoes and citrus), pasture crops (e.g. clover and lucerne) and weeds (e.g. rattle pods, phasey bean and Paterson’s curse). Life cycle on summer pulses: Green mirids may be present at any crop stage from seedlings to podding. Mirid populations are typically low during the vegetative phase, but can increase rapidly after budding as a result of in-crop breeding. Over 80% of mirids in flowering legumes may be nymphs and populations in excess of 10 per square metre are not uncommon. Populations frequently decline once flowering ceases. Eggs are inserted singly into plant tissue with a small area of the egg exposed (Fig. 7.18). There are usually five nymphal stages,
Risk period: During budding and flowering, but mirids may be more damaging during late flowering and mid podding, when plants are less able to compensate for damage. Green mirids are active from spring to late summer and early autumn. Influxes of adults often follow north-west winds. Low populations (a 1 per square metre) of green mirids are often present in vegetative crops (sometimes as early as the seedling stage), but there is no evidence they cause ‘tipping’ of vegetative terminals or yield loss. Very high mirid populations (equivalent to 30 mirids per square metre) may significantly slow vegetative growth (reduced plant height and shorter internode length). Damage: Green mirids can cause significant yield losses to summer pulses. Yield reductions of 25–50% are common, where high populations (e.g. 5–10 per square metre) are left uncontrolled in mung beans. Green and brown mirids (Creontiades spp.) can reduce mung bean yields by 3 kg ha–1 mirid-day–1 m–2 (averaged over a 4–5-week flowering period). Medium and large nymphs (instars 3–5) are as damaging as adults. The damage potential of small mirid nymphs (instars 1–2) is nearly as great as later nymphal stages, as they progress very rapidly to later instars and adulthood (within 3–6 days, respectively, at 30°C). Green mirids attack buds, flowers and small pods that subsequently abort. The risk of pod abortion decreases as pods develop; however, crop stage may be important in determining overall pod loss, as pulse crops are most likely better able to compensate for early damage than late damage. Mirid damage to larger mung bean pods is characterised by watery depressions on the pod surface, and in more severe cases by the
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browning and distortion of pods resulting in reduced seed size and seed staining (Fig. 7.22). Similar pod damage has been reported in navy beans and soy beans. Green mirids appear to prefer mung bean buds, flowers and young pods to other pulses. Severe mirid damage in mung bean results in low pod set (1–3 pods per raceme), and exposed nectaries, normally hidden on racemes with a full complement of pods (usually six to seven pods) (Fig. 7.21). However, thrips, high temperatures and moisture stress can produce similar symptoms of bud, flower and small pod-shedding. The uppermost and lowest mung bean racemes often put on fewer pods, even in the absence of mirids. Mirids also attack other summer legumes, including adzuki, cowpeas, navy beans, soy beans and peanuts. In general, determinate legumes (e.g. group VII–VIII soy beans in northern Australia) have a shorter flowering period and are at lesser risk from mirid damage than indeterminate cultivar types such as mung beans, and group III–IV soy beans in southern Australia. Recent DPI&F trials show mirid populations as high as 4/m2 have no impact on pod set in well-grown peanuts and group VII soy bean cultivars A6785 and Bunya. Monitoring: Mirids are very mobile pests and in-crop populations can increase very rapidly. Crops should be inspected twice weekly from
Fig. 7.22. Mirid damage to well-developed mung bean seeds. Note the pale discoloured marks on damaged seeds. (DPI&F Qld: J. Wessels)
budding onwards until post flowering. In row crops, the preferred method is beat-sheeting, as this method is the most effective for Helicoverpa and podsucking bugs. Use a standard-sized cloth beat sheet, 1.3–1.5 m wide × 1.5 m deep, sampling the central 1 m of row. Having a sufficiently deep sheet is important as the far side of the sheet can be draped over plants in the adjacent row, thus catching any mirids (or other insects) that might otherwise escape. The standard sample unit at each sampling location in a crop is five 1-m lengths of row (not consecutive) within a 20 m radius. Convert all mirid counts per row metre to mirids per square metre by dividing counts per row metre by the row spacing in metres. At least six sites should be sampled throughout a crop to accurately determine mirid populations. Avoid sampling during very windy weather. In broadcast or narrow-row bean crops, sweepnetting is easier but captures only 33% as many mirids as a beat sheet, but only 10% as many podsucking bugs. Take 20 sweeps in 20 m, each sweep traversing 1 m of crop from crop entry to crop exit. Convert the catch to beat sheet equivalents by multiplying by 2.5. Visual sampling is too imprecise to be recommended. Suction sampling of prostrate peanuts may be the only viable mirid sampling option. While this method captures an acceptable 40% as many mirid adults as the beat sheet, it only captures 5% as many nymphs.
Fig. 7.21. Severe mirid damage: one mature mung bean pod per raceme compared with five or more where mirids were absent. These symptoms may be confused with thrips damage, heat stress and moisture stress. The yellow ‘bumps’ are nectaries and are not damage symptoms. (DPI&F Qld: J. Wessels)
Action level: Thresholds are based on beat-sheet sampling with a standard-sized beat sheet, and are expressed as mirids per square metre. Thresholds for mung beans and navy beans are 0.3–0.5 per square metre for dimethoate (a relatively cheap insecticide) at 500 mL ha–1 applied via ground rigs and aerially, 187
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respectively. Thresholds for mirids controlled with indoxacarb are considerably higher (1.1–1.3 per square metre) because this product is more expensive than dimethoate. Indicative thresholds for soy beans and peanuts are three to four per square metre in order to manage resistance of other pests. Chemical control: Dimethoate at 500 mL ha–1 (all summer pulses) or indoxacarb at 400 mL ha–1 (mung beans and soy beans only). Dimethoate is frequently applied at lower than label rates (e.g. 200–250 mL ha–1), these rates giving excellent mirid control but having far less impact on many beneficials. The addition of salt (0.5% NaCl) as an adjuvant, improves dimethoate’s effectiveness at lower rates, and the salt rate used (0.5%), has no phytotoxic effect on summer pulse crops. ‘Hard’ water can markedly lower dimethoate’s effectiveness and should be countered by adding a buffering agent such as LI700. Indoxacarb is not recommended where there is high mirid pressure (more than two per square metre) because it has less residual activity against hatching nymphs. Cultural control: Shortening a crop’s flowering period can reduce the risk of mirid damage. The chance of a shorter flowering period can be improved 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.
observed attacking mirids in the field. Naturally occurring fungi (e.g. Beauvaria) may also infect and kill mirids, but are rarely observed in the field.
S E E D B U G S ( LYG A EI DA E )
Graptostethus servus Fabricius Hemiptera: Lygaeidae Distribution: An exotic species occurring in Europe, Asia as well as Australia. In Australia, reported from northern NSW, NT, Qld, and WA (Ord). Pest status: Minor, widespread, irregular. Identification: A small, elliptically shaped bug reaching 8 mm in length. The base colour is a pale earthy orange. The pronotum and chorium are dark grey with pale edging, the latter forming a pale cross on the bug’s back (Fig. 7.24). May be confused with: This species is similar in outline to the slightly larger and more brightly coloured Lygaeid Melanerythrus mactans Stål. Host range: Mung beans, cowpeas, soy beans, cotton and sorghum and sunflowers. Risk period: Podding. Damage: Damage to small seeds interferes with their development and reduces yield. Damage to
Natural enemies: Spiders, ants, predatory bugs (Fig. 7.23) and predatory wasps have been
Fig. 7.23. Chinese black mirid, Tytthus chinensis (Stål) (3 mm), a small predatory mirid that feeds on eggs of other mirid species. (DPI&F Qld: J. Wessels)
Fig. 7.24. Graptostethus servus (10 mm) on mung beans at Kingaroy, Qld. (J. Wessels)
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older seeds reduces the quality of harvestable seeds. Monitoring: Use a beat sheet. Chemical control: No pesticides are registered for this pest. It is most likely controlled with pesticides targeting podsucking bugs and mirids. Cultural control and conservation of natural enemies: As for podsucking bugs.
Melanerythrus mactans Stål. Hemiptera: Lygaeidae Distribution: In Australia, reported from northern NSW, NT, Qld and WA (Ord). Pest status: Minor, widespread, irregular. Identification: A small elliptically-shaped bug reaching 10 mm in length. The base colour is bright red with black markings on the pronotum and chorium. There are two black spots across the middle of the back, and centrally aligned pale spots at the front and the middle of the chorium (Fig. 7.25). May be confused with: This species is similar in outline to the slightly smaller and less brightly coloured Lygaeid Graptostethus servus. Host range: Mung beans, cowpeas, soy beans and cotton. Risk period: Podding.
Damage: Damage to small seeds reduces their viability and yield. Damage to older seeds reduces the quality of harvestable seeds. Monitoring: Use a beat sheet. Chemical control: No pesticides are registered specifically for this pest. It is most likely controlled with pesticides targeting podsucking bugs and mirids. Cultural control and conservation of natural enemies: As for podsucking bugs.
Peanut trash bug Elasmolomus pallens (Dallas) Hemiptera: Lygaeidae Previously classified as Elasmolomus sordidus (Fabricius). Also known as the sesame pod bug. Distribution: An exotic species widespread throughout the old world tropics, which has now spread to South America and Australia, where it has been recorded from northern NSW, NT and Qld. Pest status: Minor, widespread, irregular. A major pest in Africa and Asia but not a problem in Australia because of mechanised peanut harvesting and bulk handling. Identification: Adults are slender bugs reaching 10 mm in length and are mottled brown, with a dark chocolate brown head and anterior pronotum (front thorax). They have banded antennae and long legs. Young nymphs have red abdomens that get progressively darker with age. May be confused with: The adults are similar to Rutherglen bugs but are slightly larger and are brown rather than grey. Host range: Peanuts. Life cycle: In the field, eggs are laid in the soil or in peanut haulms. In storage, eggs are laid loosely on pods or on sacks. There are five nymphal stages and the relatively long life cycle takes about 2 months. Trash bugs are mainly active at night.
Fig. 7.25. Melanerythrus maclans (12 mm) on mung beans, Kingaroy, Qld. (J. Wessels)
Risk period: Peanuts are attacked after plants are pulled and are drying on the soil surface prior to threshing. Peanuts in bag stacks are vulnerable to attack but bag-stacking is no longer practised in Australia. 18 9
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Damage: Adults and nymphs damage peanut kernels by piercing the pods. Damage causes immature kernels to shrivel and increases the fatty acid content of peanut oil, resulting in a rancid flavour. Monitoring: Look for any unusual bug activity in bag stacks. Adults are readily captured in light traps. Cultural control: Thresh pulled peanuts as quickly as possible if significant trash bug activity is observed in the field. Bulk storage of shelled peanuts virtually eliminates the risk of attack in Australia.
Cultural control: Earlier maturing sunflower crops are potential infestation sources. Conservation of natural enemies: As for podsucking bugs.
S Q UA S H B U G S ( C O R EI DA E )
Acroelytrum muricatum Mayr Hemiptera: Coreidae Distribution: This tropical species occurs from the Ord in WA to southern Qld. Pest status: Minor, patchy, irregular.
Rutherglen bug Nysius vinitor Bergroth Hemiptera: Lygaeidae N (Main entry in Chapter 5 Oilseeds.)
Distribution: Throughout Australia. Pest status on summer pulses: Minor, widespread, irregular (pulses are not favoured hosts). Identification: A small grey elongated bug only 3.5–4 mm in length. Its legs and antennae are pale and its wings are clear. The nymphs are pear shaped and are often brown and pinkish in colour (Fig. 5.30). May be confused with: This species is very similar to the larger grey cluster bug, N. clevelandensis Evans (5 mm long). The head of the Rutherglen bug is broader than mirids and its antennae are proportionally shorter.
Identification: A large, stout, distinctively shaped brown bug reaching 17 mm in length (Fig. 7.26). This species has spined shoulders and a wide abdomen which is 1.4 times the width across the shoulders. The nymphs are similar in outline to the adults but lack wings. May be confused with: This bug’s distinctively wide abdomen distinguishes it from most other bug pests likely to be encountered in field crops. The only bug superficially similar is the cucurbit shield bug, Megymenum affine Boisduval. However, this species has a brassy coloured wing membrane and short antennae. Host range: Soy beans and mung beans, but most likely other summer pulses also.
Host range: Most summer pulses but only rarely in large numbers. Risk period: Budding and podding. Damage: Feeding on buds and flowers aborts these structures and reduces pod set. Feeding on developing pods damages and shrivels seeds. Monitoring: Use a beat sheet. Be ready to count bugs as soon as they hit the sheet as they are extremely flighty. Chemical control: No pesticides are registered specifically for this pest. It is most likely controlled with pesticides targeting podsucking bugs and mirids.
Fig. 7.26. Adult Acroelytrum muricatum (16 mm) on soy beans, Bundaberg, Qld. (DPI&F Qld: J. Wessels)
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Fig. 7.28. Adult crusader bug (22 mm) on cowpeas, Ord River, WA. (M. Shepard)
Crusader bug Mictis profana (Fabricius) Hemiptera: Coreidae Fig. 7.27. Cletus sp. adult (6 mm) on mung beans, Kingaroy, Qld. (J. Wessels)
Risk period: Podding. Damage: This species most likely causes damage similar to that caused by other podsucking bugs. Monitoring: Use a beat sheet.
Cletus sp. Hemiptera: Coreidae Distribution: Reported from northern Australia. Similar Cletus spp. are reported in Asia. Pest status: Minor, irregular. Identification: A small but stout brown bug reaching only 7 mm in length. May be confused with: Cletus sp. has a similar outline but is much smaller than Acroelytrum muricatum. Host range: Legume hosts include soy beans, mung beans, and Dolichos. Other hosts include cashews, squash and sorghum. Risk period: Podding. Damage: Cletus sp. most likely causes damage similar to that caused by other podsucking bugs. It is most likely less damaging than larger species. Monitoring: Use a beat sheet.
Distribution: Moluccas to Fiji. Widespread in Australia. Pest status: Minor, widespread, irregular. Identification: A large (25 mm long), robust and elongated dark brown bug with a large pale diagonal cross on its back (like a crusader’s shield) (Fig. 7.28). Its hind legs are thick and strong and it has spined shoulders and relatively long, pale-tipped antennae. Crusader bugs have small vents above the base of the middle legs, used to emit foul smells to repel predators. The nymphs are flatter and more tapered towards the head than adults and have long antennae. They are brown with two creamy yellow spots on the middle of their backs. May be confused with: The distinctive cross distinguishes this bug from other coreid species. Host range: Soy beans, navy beans and lablab bean (Lablab purpureus). Life cycle: The bug has five nymphal stages. Risk period: Podding. Damage: Potentially very damaging because of its large size. Monitoring: Use a beat sheet. Action level: Use green vegetable bug thresholds. 191
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Passionvine bug Fabrictilis gonagra (Fabricius) Hemiptera: Coreidae Also referred to as the leaf-footed plant bug. Previously classified as Leptoglossus australis (Fabricius) and sometimes referred to as Fabrictilis australis. Distribution: An exotic pest reported from northern NSW, NT and Qld.
after emergence but older instars tend to disperse. Nymphal development takes about 50 days. Adults can live for several weeks. Risk period: Podding. Monitoring: Use a beat sheet. Action level: As for the green vegetable bug.
D I N I D O R I DA E
Pest status: Minor, mostly coastal, irregular. Identification: This elongated but robust bug is 18 mm long and 6 mm wide, and is dull purple black with a transverse red/orange band behind the head and several red spots on the underside of the body (Fig. 7.29). The hind-leg tibia (lower leg segment) is swollen and flat. The eggs are laid in chains of 16–17, and are brownish in colour, cylindrical in shape and 1.5 mm long. Nymphs are similar in shape than the adults but lack wings and are reddish in colour when young. There are some black spines on the head and thorax. May be confused with: Adults may be confused with the large brown bean bug, but are larger and more robust and have orange spots underneath, as opposed to the latter’s yellow lateral bands. Host range: Cowpeas, mung beans, navy beans, passionfruit, cashew, citrus, cucurbits and pomegranate. Life cycle: The eggs are laid in chains of 16–17 and hatch in 6–7 days. Nymphs cluster soon
Fig. 7.29. Passionvine bug, Fabrictilis gonagra (Fabricius), (18 mm) on cowpeas, Katherine, NT. (M. Shepard)
Cucurbit shield bug Megymenum affine Boisduval Hemiptera: Dinidoridae Also known as the pumpkin bug. Distribution: A native species reported from northern NSW, NT and Qld. Pest status: Minor, irregular. Identification: A large stout brown bug (Fig. 7.30) with a distinctive brassy coloured wing membrane, a ‘cape-like’ pronotum (front thorax segment), and short antennae. Adults are about 14 mm long and 9 mm wide. May be confused with: The cucurbit shield bug is similar in outline to Acroelytrum muricatum (Family Coreidae), but has much shorter antennae and a brassy coloured wing membrane. Host range: This species is recorded in peanuts in the south Burnett region of Qld. Other hosts include cucurbits.
Fig. 7.30. Cucurbit shield bug (14 mm) on peanuts, Kingaroy, Qld. (J. Wessels)
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Risk period: Podding. Monitoring: Use a beat sheet.
A LY D I DA E
Large brown bean bug Riptortus serripes (Fabricius) Hemiptera: Alydidae Frequently referred to as the brown bean bug (BBB) or the podsucking bug. The preferred common name is the large BBB to avoid confusion with the small BBB (Melanacanthus scutellaris). Distribution: Native to Australia. Reported from NSW, NT and Qld, and likely tropical WA. Similar Riptortus species (e.g. R. clavatus, R. dentipes and R. linearis) occur in Asia, India and Africa. Pest status: Major and widespread. Irregular in some regions and crops but more frequent on the coast and with a preference for the Vigna legumes, e.g. adzuki beans, cowpeas and mung beans. Identification: Nymphs are ant-like (Figs 7.31 and 7.32), while the adult is an elongated dark brown bug, 16–18 mm in body length, with long antennae and with a bright yellow stripe along each side (Fig. 7.33). This stripe is more pronounced in males. Females are also ‘rounder’ in the body than males and have a broad pale band on their rear tibia. Riptortus serripes’s body narrows in the middle and it has a spine on each ‘shoulder’. It also has large, robust and spiny hind legs. When flying, the bright orange top of
Fig. 7.31. Large brown bean bug, first-instar nymph, 2 days old (2.5 mm long). (J. Wessels)
Fig. 7.32. Large brown bean bug. fouth-instar nymph (9 mm). (DPI&F Qld: J. Wessels)
its abdomen is strikingly revealed. Riptortus eggs are a dark purple–brown in colour and are laid singly or in small clusters. They are slightly elliptical with a flattened top and rounded base and are 1.5 mm long. May be confused with: Riptortus obscuricornis Dallas, which is slightly smaller (15 mm long) than the large brown bean bug, is a lighter brown, and has pale cream (as opposed to yellow) lateral stripes extending the whole length of the body. This species also may be confused with the small brown bean bug, Melanacanthus scutellaris, which is considerably smaller (10–12 mm long) and is not as robust, and with the rice or paddy bug, Leptocorisa acuta (Thunberg). Rice bugs are pale green or brown, reach 15 mm in length, and are very slender with thin hind legs (Fig. 7.34). Riptortus and Melanacanthus nymphs are difficult to distinguish. However, later-instar Melanacanthus nymphs can be distinguished by their more
Fig. 7.33. Large brown bean bug, female (17 mm). (DPI&F Qld: J. Wessels)
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Damage: R. serripes is as damaging as green vegetable bug. Damage is similar to that caused by green vegetable bug, with early damage reducing yield while later damage reduces the quality of harvested seed. Monitoring: The beat-sheet method can be used but may underestimate numbers as adults are very flighty, particularly during the hotter parts of the day. Crops should be sampled during the early morning. Flying and escaping Riptortus adults should be included in these counts.
Fig. 7.34. Paddy bug, Leptocorisa acuta (15 mm), from grassy soy beans, Kingaroy, Qld. The thin hind legs differentiate this species from brown bean bug adults. (DPI&F Qld: J. Wessels)
elongated abdomens that have six or more small dark spots on their dorsal surface. Riptortus serripes nymphs are dark brown and are similar in outline to ants (Figs 7.31 and 7.32), but they lack the very narrow waist and biting mouthparts (jaws) of ants. Host range: Riptortus spp. attack all summer pulses including adzuki beans, cowpeas, mung beans, navy beans, soy beans and pigeon peas, and also guar. Mung beans (Vigna radiata) and other Vigna legumes in particular are favoured hosts. Riptortus serripes has also been reported in lucerne, siratro, citrus and choko (Sechium edule), as well as a range of leguminous weeds including phasey bean, rattle pods (Crotalaria spp.), sesbania (Sesbania cannabina) and Senna spp. Life cycle: Riptortus serripes typically invade summer legumes at flowering and commence feeding and egg-laying. Females lay scattered single eggs, not is rafts. Riptortus serripes has five nymphal stages that usually reach a damaging size during mid to late podfill. Development times for Riptortus serripes eggs and nymphs are about 8 and 17 days, respectively, (25 days total) at 26°C. Under laboratory conditions, individual females have laid up to 600 eggs over a 94-day period. In Qld, R. serripes most likely has four generations per year. Overwintering R. serripes may shelter in curled up dead leaves.
Chemical control: Current (2006) registered insecticides (only in guar) give unreliable control in densely canopied crops. Riptortus serripes is likely to be controlled by synthetic pyrethroids that are registered against the green vegetable bug. Cultural control: Where possible, avoid sequential plantings of summer legumes as this allows successive generations of podsucking bugs to build up. Spring plantings (where feasible, e.g. with mung beans) are at lesser risk than summer-planted crops. Control of weed hosts such as phasey bean, sesbania and rattle pods bean is also recommended. Conservation of natural enemies: Spiders, ants, and predatory bugs are likely predators of R. serripes eggs and nymphs.
Small brown bean bug Melanacanthus scutellaris (Dallas) Hemiptera: Alydidae Distribution: Native to Australia. Reported from NSW, Qld, and WA and probably the NT. A similar species, Melanacanthus margineguttatus, has been reported in pulses in the Ord River Irrigation area in WA. Other Melanacanthus species, e.g. M. jaculus and M. torridus, are reported in Africa. Pest status: Major, widespread, irregular. Identification: Small brown bean bug eggs are laid in small clusters, are shiny olive-green in colour, slightly elliptical in shape, with a flat top and a rounded base and are 1.0 mm long (Fig. 7.35). The adult is an elongated brown bug 10–12 mm in length (not including legs and antennae), with long antennae and with a cream stripe along each side (Fig. 7.36). This stripe is
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Fig. 7.35. Eggs (1.0 mm diameter) of the small brown bean bug are laid in small, loose clusters. (DPI&F Qld: J. Wessels)
Fig. 7.37. Female small brown bean bug (12 mm). The female has a thicker abdomen than the male and lacks a pale patch on its scutellum. (DPI&F Qld: J. Wessels)
often less distinct in females, which are ‘rounder’ in the body than males (Fig. 7.37). Males also have a prominent pale patch in the scutellum and a short spine on each ‘shoulder’ (less pronounced than on Riptortus spp.), and have moderately robust and spiny hind legs (thinner than those of Riptortus spp.).
be confused with rice or paddy bugs, Leptocorisa spp. Melanacanthus and Riptortus nymphs are difficult to distinguish; however, later-instar Melanacanthus nymphs can be distinguished by their more elongated abdomens, which have six or more small dark spots on their dorsal surface.
The nymphs are dark brown to black and are similar in outline to ants; however, close inspection shows they lack the very narrow gaster (waist) that is typical of ants.
Host range: Melanacanthus scutellaris attacks all summer pulses including adzuki beans, cowpeas, mung beans, navy beans, soy beans and pigeon peas, and also guar (Cyamopsis tetragonoloba). Mung beans (Vigna radiata) and other Vigna legumes in particular are favoured hosts. Melanacanthus scutellaris has also been reported in lucerne and siratro, as well as a range of leguminous weeds including phasey bean and rattle pods (Crotalaria spp.).
May be confused with: Riptortus obscuricornis Dallas, which is larger and more robust (15 mm long) but which also has pale cream lateral stripes. It is less likely to be confused with the much larger and robust large brown bean bug, Riptortus serripes (16–18 mm long), which has yellow lateral stripes. Melanacanthus may also
Fig. 7.36. Male Melanacanthus scutellaris (12 mm). Note the prominent pale patch in the scutellum (the triangular area between the front of the wings). (J. Wessels)
Life cycle: Melanacanthus scutellaris typically invade summer legumes at flowering and commence feeding and egg-laying. Melanacanthus lay scattered single eggs. There are five nymphal stages and nymphs usually reach a damaging size to coincide with mid to late podfill. Development times for M. scutellaris eggs and nymphs are about 6 and 20 days, respectively, at 26°C. Under laboratory conditions, individual females have laid up to 300 eggs over a 58-day period (about five eggs per day). Potentially two M. scutellaris generations could develop per summer pulse crop. Risk period, action level, cultural control, and conservation of natural enemies: Refer to the green vegetable bug and large brown bean bug. 19 5
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Damage: M. scutellaris is as damaging as the green vegetable bug and the large brown bean bug (Riptortus serripes). Refer to these species. Monitoring: Beat-sheet sampling may underestimate numbers of M. scutellaris, as they quickly fly away when disturbed. Crops should be sampled during the early morning and crop scouts should familiarise themselves with the appearance of flying (and escaping) M. scutellaris and include these in sampling counts. Chemical control: As for Riptortus serripes. Fig. 7.39. Brown shield bug fourth-instar nymphs (5 mm). (H. Brier)
S H I E L D B U G S ( PE N TATO M I DA E )
Brown shield bug Dictyotus caenosus (Westwood) Hemiptera: Pentatomidae Also known as the brown stink bug and (less frequently) as the brown ground bug. Distribution: Native to Australia but introduced to New Zealand and New Caledonia. Reported from NSW and Qld, but most likely more widely spread. Pest status: Minor, widespread, irregular. Identification: Brown shield bugs lay eggs in either small twin rows or small irregular rafts containing 10–16 eggs (Fig. 7.38). The eggs are pale cream and similar in shape to eggs of the green vegetable bug. Newly hatched nymphs are orange with black markings and are very similar to newly hatched nymphs of many other shield bugs. Larger nymphs have dark brown
Fig. 7.38. Brown shield bug egg rafts, twin-row form, each with 14 eggs. (H. Brier)
(sometimes almost black) heads and thoraxes, and a pale brown abdomen with transverse dark brown and pale (almost white) markings at its centre (Fig. 7.39). There is also a transverse pale band at the front of the abdomen. Brown shield bug adults are shield-shaped, matt mid-brown (Fig. 7.40), and are noticeably smaller (8 mm long) than green vegetable bug adults. May be confused with: Adults may be confused with those of the glossy shield bug, Cermatulus nasalis (Westwood), which is slightly larger and is a predatory species. Eggs and nymphs of this species are distinct from brown shield bug eggs and nymphs. Host range: All summer pulses but usually in low numbers. Sometimes in large numbers in
Fig. 7.40. Adult brown shield bug (8 mm). (DPI&F Qld: J. Wessels)
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soy beans and other summer pulses in the western Darling Downs of Qld, and in northwest NSW. Also reported in lucerne, clover, horehound and emu foot (Psoralea tenax). Life cycle: Brown shield bugs typically invade summer legumes at flowering and commence feeding and egg-laying. Nymphs usually reach a damaging size during mid to late podfill. There are five nymphal stages. Usually only one generation develops per summer legume crop but more than one generation is possible if temperatures are high. Damage: The brown shield bug damages only 20% as many seeds as the green vegetable bug (soybean data). Refer to the green vegetable bug and the box.
Fig. 7.41. Egg raft of the green stink bug (13 eggs). (DPI&F Qld: J. Wessels)
Risk period, action level, chemical and cultural control: Refer to the green vegetable bug and the box. Monitoring: As for green vegetable bug. Beatsheeting is the preferred sampling method. Sample crops in the early to mid morning when bugs are more likely to be at the top of the crop. Look for the distinctive egg rafts, which indicate the presence of brown shield bug. Conservation of natural enemies: Spiders, ants, and predatory bugs are major predators of the brown shield bug, particularly of eggs and young nymphs, with mortality of these stages sometimes exceeding 90%. Eggs may be parasitised by the tiny wasp, Trissolcus basalis.
only 8 mm long and have a green shield-shaped body with brown wing covers (Fig. 7.43). May be confused with: The two-tone colouring of adults distinguishes the green stink bug from other stink bug (pentatomid) pests in Australia. The nymphs and eggs are similarly distinctive. Host range: All summer legumes. Other hosts include sorghum, horehound and raspberry. Life cycle: Green stink bugs typically invade summer legumes at flowering and commence feeding and egg-laying. Females can lay over 400 eggs. Nymphs usually reach a damaging size during mid to late podfill. There are five nymphal stages. Usually only one generation develops per summer legume crop, but more
Green stink bug Plautia affinis (Dallas) Hemiptera: Pentatomidae Distribution: Native to Australia, where it is reported from NSW and Qld but is most likely more widely spread. Pest status: Minor, widespread, very irregular. More commonly encountered in sorghum. Identification: The eggs are similar in shape to those of green vegetable bug but are olive-green (as opposed to cream). They are laid in small loose rafts with only 5–15 eggs (Fig. 7.41). Nymphs are cream and yellow with prominent dark markings, or are mossy green with dark markings on their back (Fig. 7.42). Adults are
Fig. 7.42. Green stink bug, fifth-instar nymph, green form (6 mm). (J. Wessels)
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Fig. 7.44. Fertile green vegetable bug egg raft (113 eggs) with typical pink–orange colouration. (DPI&F Qld: J. Wessels) Fig. 7.43. Green stink bug adult (8 mm). (DPI&F Qld: J. Wessels)
than one generation is possible if temperatures are high. Risk period: Podding. Damage: The green sting bug is the least damaging of the podsucking bugs, attacking summer legumes in Australia, damaging only 10% as many seeds as the green vegetable bug (soybean data).
frequently referred to by its acronym, GVB, or colloquially as vegie bug. Distribution: A cosmopolitan pest that originated from the Mediterranean area or north-east Africa. Now found in tropical, subtropical and warmer temperate regions of the world. The GVB was first recorded in Australia in 1916 (an accidental introduction) and is now found in all Australia states and territories.
Monitoring: As for green vegetable bugs. Beatsheeting is the preferred sampling method. Sample crops in the early to mid morning when bugs are more likely to be at the top of the crop. Look for the distinctive small egg rafts, which indicate the presence of green stink bugs.
Pest status: Major, widespread and regular. This species is the most damaging podsucking bug in pulses, by virtue of its abundance, widespread distribution, rate of damage and fecundity. It is one of the most recognised agricultural pests in Australia.
Action level, chemical and cultural control: Refer to the green vegetable bug.
Identification: Newly laid eggs are cream in colour but turn bright orange just prior to hatching (Fig. 7.44). Eggs are laid in rafts (50–100 eggs per raft) and are circular in cross-section. Parasitised GVB eggs are black. Nymphs are variable in colour. Newly hatched nymphs (1.5 mm long) are initially orange and brown (Fig. 7. 45), later turning black. Later-instars are either green or black, with white, cream, orange and red markings, and are round or oval rather than shield-shaped. (Figs 7.46 and 7.47). Final (fifth) instar nymphs are also either green or black, but are not as prominently patterned and have prominent wing buds (Fig. 7.48). Adults are bright green and shield-shaped, and are 13– 15 mm long (Fig. 7.49). Adult GVB have three small white spots at the front of the scutellum (i.e. between their shoulders). Overwintering
Conservation of natural enemies: Spiders, ants and predatory bugs are major predators, particularly of eggs and young nymphs with mortality of these stages sometimes exceeding 90%. Eggs may be parasitised by the tiny wasps Trissolcus basalis, T. oenone and Telenomus cyrus (Scelionidae).
Green vegetable bug Nezara viridula (Linnaeus) Hemiptera: Pentatomidae Referred to as southern green stink bug in most countries except Australia. In Australia, it is 19 8
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Fig. 7.45. Green vegetable bug egg raft, with newly emerged orange nymphs. (H. Brier)
Fig. 7.48. Fifth-instar green vegetable bug nymph, green form (12 mm), with an egg of the Trichpoda fly parasite behind its head on the right. (DPI&F Qld: J. Wessels)
adults are usually purple–brown in colour (Fig. 7.50). Occasionally, yellow and orange GVB variants are seen in summer populations. All stages emit a foul smell when handled, a defense against predators.
Fig. 7.46. Third-instar green vegetable bug nymph (5 mm). (DPI&F Qld: J. Wessels)
Fig. 7.47. Fourth-instar green vegetable bug nymph (8 mm). (DPI&F Qld: J. Wessels)
May be confused with: Adults are very similar to Glaucias amyoti (Dallas), which attacks pecans and ornamentals but not pulses. Glaucias amyoti is slightly larger and more rounded, and is a more yellow–green than N. viridula. Glaucius amyoti also has a yellow edging to its pronotum (the first thoracic segment), which is smooth as opposed to GVB’s, which is dentate. Smaller podsucking bugs that GVB adults may be confused with include the redbanded shield bug (Piezodorus oceanicus), which has a pink or white
Fig. 7.49. Adult green vegetable bug (15 mm) with six eggs of the parasitoid Trichopoda giacomellii on its head and anterior thorax. (K. Power)
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seeds to complete their development, and the podding phase of most summer legumes is only slightly longer in duration than GVB’s life cycle. The eggs are laid in rafts containing 50–100 eggs and take 6 days to hatch at 25°C. There are five nymphal instars, instars 1–5 being of approximately 5, 5, 5, 6 and 9 days duration, respectively, at 25°C, a total development time (eggs to adult) of 36 days at the optimum temperature of 25°C. Development is slower at temperatures lower than 25°C. GVB overwinter as adults, often sheltering in yet-to-be harvested maize crops, under bark on trees, or in farm buildings. Fig. 7.50. Adult green vegetable bug in purple overwintering colours (14 mm). Note the three small white spots at the front of the scutellum. (DPI&F Qld: J. Wessels)
band across its ‘shoulders’, the green stink bug (Plautia affinis), which is green and brown in colour, and the green potato bug (Cupsicona simplex), which is green with short spine on each shoulder. The latter two species are of minor importance in pulses. Nymphs of these species are quite different in colour to GVB nymphs. Parasitised GVB eggs may be confused with eggs of the predatory shield bugs Cermatulus nasalis and Oechalia schellenbergii, but lack the spines that ring the top of the eggs of these species. Host range: A polyphagous insect. Attacks all summer and winter pulses (except chickpea). Summer pulse hosts include adzuki beans, cowpeas, mung beans, navy beans, soy beans, pigeon peas and guar. Soy beans are a favoured host. GVB is also a pest in cotton, sesame and many horticultural crops including tomatoes, silverbeet, cucurbits, maize, capsicums and pecans. Many weeds are hosts, including wild radish (a favoured spring host), rattle pods (Crotalaria spp.), wild turnip, noogoora burr Xanthium pungens, marshmallow, castor oil Ricinus communis and wild tobaccos (Nicotiana spp.). Life cycle: GVB typically invade summer legumes at flowering and commence feeding and egg-laying. Usually only one generation develops per summer legume crop. Nymphs usually don’t reach a damaging size until mid to late podfill. Nymphs require pods containing
Risk period: GVB typically invade summer legumes at flowering. Summer legumes remain at risk until pods are too hard to damage (i.e. very close to harvest). Damaging populations are typically highest in late summer crops during late podfill (when nymphs have reached or are near adulthood). Damage: GVB are primarily pod-feeders with a preference for pods containing well-developed seeds, but they also damage buds and flowers. Damage to young pods produces deformed and shrivelled seeds (Fig. 7.51), reducing yield. Some pulse cultivars can compensate for early damage, but seeds damaged in older pods are blemished and difficult to grade out, reducing harvested seed quality, particularly that destined for human consumption. GVB and other podsucking bugs can damage ‘close-toharvest’ black mung bean pods (i.e. pods that have blackened and hardened just prior to harvest). Bug-damaged seeds have increased
Fig. 7.51. Green vegetable bug damage to soy beans during early podfill. (DPI&F Qld: J. Wessels)
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on soybean yield. However, in high rainfall regions, increased weathering of bug damage may reduce seed quality to unacceptably low levels, even for stockfeed beans. To compensate for the different damage potential of immature stages, immature bug counts should be converted to adult GVB equivalents (see box). Fig. 7.52. Bug-damaged navy bean seeds (left) compare to undamaged seeds (right). Bug damage is very noticeable in white-seeded (or pale-seeded) pulses with large areas of disfiguring staining around the feeding area. (DPI&F Qld: J. Wessels)
protein content but a shorter storage life (due to increased rancidity). Bug damage to soy beans also reduces seed oil content. Bug-damaged seeds are frequently discoloured, either directly as a result of tissue breakdown, or because of diseases such as Cercospora (purple seed stain), which may gain entry where pods are pierced by bugs (Fig. 7.52). High GVB populations (more than 10 per sqare metre) in coastal regions of Qld have been responsible for damaging soybean crops to the extent that they were not even acceptable for the crushing market. As bug-damaged seeds are more prone to weathering than undamaged seeds, crops in higher rainfall regions are likely to suffer greater reductions in quality for a given bug population. Action level: Thresholds are determined by quality requirements. A critical level of bug damage for edible pulses is about 2%, which may be produced by bug densities ranging from 0.25 to 0.7 per square metre. In a well-grown crop of edible soy beans (with 1500 seeds per sqaure metre yielding 2.5 t ha–1), 0.5 GVB per square metre produce 2% damage of harvestable seeds, so an action threshold of about 0.33 bugs per square metre is necessary to prevent damage. For a low-yielding 1.25 t ha–1 soybean crop with only 750 seeds per square metre, 0.25 GVB per square metre would produce 2% damage, so the economics of control may be marginal. Soy beans for stockfeed have higher tolerances: an indicative threshold is one adult GVB per metre at the start of podfill up to 2 weeks into podfill, when the soybean seed cavity is covered, after which bug damage has little to no impact
Monitoring for all podsucking bugs on summer pulses: Crops should be inspected twice weekly from budding until close to harvest. Sample for bugs in the early morning when most bug stages are visible at the top of the canopy. Beat-sheet sampling is the most efficient monitoring method for podsucking bugs including GVB. Use a beat sheet (cloth), 1.3–1.5 m wide × 1.5–2.0 m deep, sampling the central 1 m of row. Having a sufficiently deep sheet is important as the far side of the sheet can be draped over plants in the adjacent row, thus catching any bugs (or other insects) that might otherwise escape. The standard sample unit consists of five 1-m lengths of row (not consecutive) within a 20 m radius. In very tall soy beans, use a shallow tray to beat into. Convert all bug counts per row metre to bugs per square metre by dividing counts per row metre by the row spacing in metres. At least six sites should be sampled throughout a crop to accurately determine adult podsucking bug populations. Nymphal GVB populations are more difficult to sample accurately as their distribution is extremely clumped, particularly during the early nymphal stages (1–3). At least 10 sites (with five non-consecutive row metres sampled per site) should be sampled to adequately assess nymphal populations. In practice, it is unlikely that scouts will have the time to take so many samples. However, inadequate sampling is a major reason that sizeable populations of large GVB nymphs are frequently not ‘discovered’ in many crops until late pod development. Chemical control for all podsucking bugs on summer pulses: Synthetic pyrethroids give effective control of GVB and other podsucking bugs, but these insecticides are incompatible with IPM and on the management of easily flared pests such as silverleaf whitefly. Not all registered products give effective GVB control. Dimethoate is only effective against small 201
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Fig. 7.53. Trissolcus basalis wasps emerging from parasitised green vegetable bug egg raft. (DPI&F Qld: J. Wessels)
nymphs, and methomyl is less effective in densely canopied crops because of its short residual activity. The organophosphate (OP) trichlorfon gives only moderately effective control. Spraying is best done in the morning when GVB and many other podsucking bugs are basking at the top of the canopy. Cultural control for podsucking bugs on summer pulses: Where possible avoid sequential plantings of summer legumes, and avoid cultivar and planting time combinations that are more likely to lengthen the duration of flowering and podding (e.g. early plantings of the soybean cultivar Melrose). Spring plantings (where feasible, e.g. with mung beans) are at lesser risk than summer planted crops. Conservation of natural enemies: Spiders, ants, and predatory bugs are major predators of GVB, particularly of eggs and young nymphs with mortality of these stages sometimes exceeding 90%. GVB eggs are frequently parasitised by the tiny introduced wasp Trissolcus basalis (Wollaston) (Fig. 7.53), particularly in warmer coastal regions. Adults and fifth-instar GVB nymphs may be parasitised by the tachinid fly, Trichopoda giamocellii (Blanchard), recently introduced from South America (Figs 7.54 and 7.55). Naturally occurring fungi (e.g. Beauvaria) may also infect and kill GVB, but are rarely observed in the field.
Fig. 7.54. Trichopoda larva (5 mm) inside dissected green vegetable bug. (DPI&F Qld: J. Wessels)
Pest status: Minor, restricted, irregular in tropical regions. Identification: Small, dark brown, mottled shield bugs reaching 7–10 mm long (Fig. 7.56). Nymphs are also mottled and have banded legs. May be confused with: Oncocoris spp. may be confused with the brown stink bug, Dictyotus caenosus, but are darker and more mottled. Host range: Cowpeas, mung beans and soy beans. Life cycle: Probably similar to other Pentatomids.
Oncocoris hackeri McDonald and O. coelebs (Fabricius) Hemiptera: Pentatomidae Distribution: A tropical species recorded from the NT, the Ord in WA, and coastal Qld. Nonpest Oncocoris species are found throughout Australia.
Fig. 7.55. Female Trichopoda giacomellii (Blanchard) (8 mm). Female Trichopoda are distinguished from the brighter orange males by their dark abdomen. (DPI&F Qld: J. Wessels)
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Damage potential of podsucking bug populations computers) that are user friendly, and which factor in all the above variables. Illustrated below are threshold and bug-damage potential models for a soybean crop with 2000 seeds/m2, and 4 mainly small (2nd instar) GVB nymphs/m2. In all crops, control is warranted where the damage
potential exceeds the threshold. In the crop in question, control is warranted if bugs of the specified age and density are detected at more than 15 days from harvest. However, spraying is not warranted if the same bug population is detected within 15 days from harvest.
Bug damage potential in GVBAEQ vs bug threshold
2
3.5
GVBAEQ/m
The damage potential of podsucking bug populations in pulse crops is determined by the bug species and nymphal stages present, time to harvest, and nymphal mortality. As most bug thresholds are based on maximum tolerabe % damage (usually 2%), crop size (seeds per unit area) is also a major factor determining whether a given bug population is likely to cause economic damage. Obviously, the closer a crop is to harvest, the more bugs are required to inflict a given level of damage, and the higher the threshold. Conversely, the closer to harvest, the lower the damage potential of small nymphs as they will not have enough time to reach a damaging size. The Queensland DPI&F is currently (2007) refining bug threshold models (for palm-top
3.0
Your bugs #
2.5
Threshold *
2.0
# Damage potential of GVB population with 4 mostly small 2 nymphs/m
1.5 1.0 0.5
* for a crop with 2000 seeds/m
0.0 0
Damage: This species attacks seeds but is far less damaging than Nezara viridula and Piezodorus oceanicus (Strickland, pers. comm.).
5
10
15
20
25
30
35
40
2
45
Days to harvest maturity
Risk period, action level, monitoring, and chemical and cultural control: Refer to the green vegetable bug.
Redbanded shield bug Piezodorus oceanicus (Montrouzier) Hemiptera: Pentatomidae Redbanded shield bugs in Australia were previously classified as Piezodorus hybneri (Gmelin), and more recently as P. grossi Ahmad or P. grossi Staddon.
Fig. 7.56. Male and female Oncocoris hackeri (8 mm) on cowpeas, Ord River, WA. (M. Shepard)
Distribution: Piezodorus spp. have been reported from Africa, India, Asia and Japan, as well as the Americas. In Australia, P. oceanicus has been reported (previously as P. hybneri) from NSW, Qld, SA and WA, but is most likely present throughout mainland Australia. Some Asian specimens, previously identified as P. hybneri, 203
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Fig. 7.57. The distinctive twin-row egg raft (30 eggs) of the redbanded shield bug, Piezodorus oceanicus (Montrouzier), with emerging nymphs. (DPI&F Qld: J. Wessels)
may also be P. oceanicus. Piezodorus oceanicus and P. hybneri are very similar in appearance to Piezodorus guildinii (the small green stink bug), which occurs in the Americas. Pest status: Major, widespread, regular. Piezodorus oceanicus is 75% as damaging as GVB in summer pulses, and is sometimes the dominant podsucking bug in the tropics. Identification: Newly hatched redbanded shield bug (RBSB) nymphs are orange with black markings and are very similar to newly hatched nymphs of many other shield bugs. However, newly hatched RBSB nymphs (and nymphs of other pentatomids) are readily identified by examining the egg raft from which they have just emerged. RBSB has a very distinctive twin-row raft with dark elliptical (in cross-section) eggs ringed by small spines (Fig. 7.57). Egg rafts contain 15–40 eggs. Close examination shows that eggs have a central pale band. Larger nymphs are pale green with dark red and brown markings in the centre of their back (Fig. 7.58). Late autumn nymphs may turn a pale pinky brown.
Fig. 7.58. Fifth-instar nymph of the redbanded shield bug, (8 mm). (DPI&F Qld: J. Wessels)
Fig. 7.59. Female redbanded shield bug (10 mm long) showing female colouration with a pink transverse band and a pink perimeter band, which in most males is offwhite in colour. (DPI&F Qld: J. Wessels)
Adult RBSBs are shield-shaped and pale green, with a noticeable band across their shoulder (Fig. 7.59). They are noticeably smaller than the similarly shaped green vegetable bug, being only 8–10 mm long. Most female RBSBs (> 90%) have a pink (not red) band across their shoulders and pink bands along their sides. These bands vary in intensity between specimens from a deep rich pink to very pale pink. In contrast, most males (> 90%) have an off-white band across the shoulders and pale yellow bands along their flanks. In both sexes, the transverse shoulder band is flanked by a dark grey–purple area that doesn’t quite extend the full width of the shoulder. May be confused with: RBSB adults are similar in shape to GVB, but are smaller and paler, with pink, white or yellow bands absent on green vegetable bug adults. The eggs and most nymphal stages of RBSBs are distinctive from those of other shield bugs in pulse crops. Host range: A polyphagous insect, attacking all summer and winter pulses except chickpeas. Soy beans in particular are a favoured host. RBSB has also been reported in cotton, citrus, clover, lucerne, siratro, sunflowers, tobacco, tomatoes and maize. RBSB attack a wide range of weeds, including wild radish, rattle pods (Crotalaria spp.), Rhynchosia and phasey bean (Macroptilium lathyroides). Life cycle: In Qld, RBSB most likely has four generations per year. RBSB typically invade summer legumes at flowering and commence feeding and egg-laying. Usually only one RBSB generation develops per summer legume crop, but more than one generation is possible if temperatures are high. Nymphs usually reach a
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Fig. 7.60. Redbanded shield bug damage to mung bean seeds. (DPI&F Qld: J. Wessels)
damaging size during mid to late podfill. Adults and nymphs probably require pods containing developing seeds (i.e. pods > R5 stage) to reproduce and to complete their development, respectively. Eggs are laid in rafts containing 20–40 eggs and take 4–5 days to hatch. RBSB has five nymphal stages. The total development time (egg to adult) is 18–35 days depending on temperature. Damage: Damage is similar to that caused by the green vegetable bug, with early damage reducing yields, and later damage reducing the quality of harvested seeds (Fig. 7.60). In mung beans, cowpeas, adzuki beans and navybeans, Piezodorus are rated as 75% as damaging as GVB, but are rated as only 67% as damaging in soy beans. Risk period, monitoring and cultural control: Refer to the green vegetable bug. Action level: Refer to p. 203. RBSBs are equivalent to 0.75 of a GVB. Chemical control: None of the pesticides registered in pulses against GVB (deltamethrin, trichlorfon, methomyl or dimethoate) give any control of RBSB. However, recent trials show the addition of 0.5% salt (NaCl) to deltamethrin increases control of RBSB to a mediocre 40%. Conservation of natural enemies: Spiders, ants and predatory bugs are major predators of RBSB, particularly of eggs and young nymphs, with mortality of these stages sometimes exceeding 90%. Eggs may be parasitised by the tiny wasp, Trissolcus basalis. Adults are infrequently parasitised by the recently introduced tachinid fly, Trichopoda giamocellii.
Fig. 7.61. The ricespotting bug, Eysarcoris distinctus, on soy beans, Childers, Qld. (6 mm). (J. Wessels)
Ricespotting bugs Hemiptera: Pentatomidae Eysarcoris trimaculatus (Distant) and E. distinctus Schouteden Also known as rice stink bugs. Distribution: Native to Australia. Eysarcoris species have been and reported from NSW, NT, Qld and WA. Eysarcoris distinctus is more common in southern Qld. Pest status: Minor, widespread, irregular. Identification: The dark elliptical eggs are either laid in single row rafts of up to 15 eggs (E. trimaculatus) or in small loose rafts (E. distinctus). Nymphs have a dark head and thorax and a light brown abdomen. Adults of both species are small (5–6 mm) but broad dark brown shield bugs, with two pale elongated marks at the anterior extremities of the scutellum. On E. distinctus, these marks are nearly at 90° to the body axis (Fig. 7.61), whereas in E. trimaculatus they are strongly angled. May be confused with: Eysarcoris spp. adults and nymphs may be confused with brown stink bugs; however, Eysarcoris spp. adults are smaller, stouter and darker than the brown stink bug. The eggs of E. trimaculatus are similar to those of Piezodorus oceanicus but are laid in a single row as opposed to twin-row rafts. Note that the ricespotting bugs are totally different to the rice bugs Leptocorisa sp. (Alydidae) (Fig. 7.34), which are grass-crop feeders. 205
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Host range: Soy beans, mung beans and cowpeas. Rice is the major host for these bugs. Life cycle: Eggs are laid on leaves. There are five nymphal instars. Risk period: Podding. Damage: Early damage shrivels seeds and reduces yields, while later damage reduces seed quality. Monitoring: Use a beat sheet.
THRIPS ( THYSANAPTER A)
Cotton seedling thrips Thrips tabaci Lindeman Thysanoptera: Thripidae Distribution: A cosmopolitan species present throughout Australia. Pest status: Minor, widespread, regular. Identification: Adults are 2 mm long and are dark, cigar-shaped and have narrow wings folded along their back. Nymphs are smaller, lack wings and are pale. Thrips species can only be determined microscopically, but cotton seedling thrips are the most likely cause of seedling damage. May be confused with: This is the only species that damages seedling crops, but for positive identification a microscope is necessary. Host range: Mung beans, navy beans, cotton and cereals.
Life cycle: Adult thrips can infest a seedling’s growing point as soon as it emerges from the ground. In cracking soils, seedlings may be infested before they emerge. Nymphs feed inside vegetative terminals. Populations typically peak within 4 weeks of plant emergence. Risk period: Spring-planted crops are at greatest risk, especially those in close proximity to maturing cereal crops. Seedlings can be infested at emergence, and sometimes even before emergence in cracking soils. Damage: Thrips attack the growing point and damage the embryonic leaves; however, damage is not manifested until the first trifoliate leaves open and is not evident in the unifoliate leaves, which may be severely distorted and discoloured. Damaged plants are stunted and look as though they have herbicide damage (Fig. 7.62). Vigorously growing crops may quickly outgrow the symptoms. Poor seedling growth caused by cold temperatures is sometimes wrongly attributed to seedling thrips. Seedling thrips damage appears to have no effect on yield or plant maturity (i.e. on time to flowering and harvest). Monitoring: Open and microscopically examine the plant’s growing point for thrips. Plucked growing points can also be dunked in alcohol to dislodge thrips. Action level: There are no thresholds for seedling thrips. Chemical control: Seedling thrips damage can be greatly reduced in mung beans, provided a systemic pesticide is applied soon after emergence (within 3–4 days). Cultural control: If possible, do not plant mung beans immediately adjacent to winter cereals. Avoid spring mung bean plantings in regions where cool spring weather is likely, as this has greater impact on mung bean growth than seedling thrips. Conservation of natural enemies: If a decision is made to control thrips, apply a narrow band spray over the seedlings to preserve predators such as spiders in the inter-row.
Fig. 7.62. Feeding damage caused by cotton seedling thrips to spring mung bean seedlings, Oakey, Qld. Note the severely distorted trifoliates contrasting with the unaffected unifoliate leaves. (H. Brier)
Flower thrips Thysanoptera: Thripidae Various species including:
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Tomato thrips, Frankliniella schultzei (Trybom) Plague thrips, Thrips imaginis Bagnall Distribution: Cosmopolitan species present in all Australian states. Pest status: Major, widespread and regular.
Chemical control: Except for western flower thrips (see below), flower thrips are easily controlled with current systemic pesticides registered in pulses (e.g. dimethoate). Cultural control: Vigorously growing crops can usually compensate for flower abortion.
Identification: Adults are 2 mm in length, cigarshaped and have narrow wings folded along their back. Nymphs are smaller, lack wings and are pale. The thrips species present can only be determined microscopically.
Conservation of natural enemies: Pirate bugs, lacewing larvae and ladybirds prey on thrips.
May be confused with: Thrips species can only be determined microscopically.
Frankliniella occidentalis (Pergande) Thysanoptera: Thripidae
Host range: Flower thrips are common in the flowers of all summer pulses, as well as other crops, ornamentals and weeds.
Distribution: A cosmopolitan species that has recently established in and spread throughout Australia. Western flower thrips (WFT) is indigenous to North America but began to spread internationally around 1980, and is now established in all continents.
Life cycle: Flower thrips feed and breed inside flowers. Risk period: Crops are at greatest risk during flowering and podset. Damage: Nymphs and adults feed in growing points and inside flowers. Thrips damage can result in flower abortion and pod distortion (Fig. 7.63). Deformed pods may be difficult to thresh, resulting in further yield losses. Thrips may also partly vector (with pollen) the damaging exotic Tobacco Streak virus recently reported in mung beans at Emerald (CQ). Monitoring: Open and examine flowers for thrips. If flowers cannot be assessed immediately, store in 70% alcohol to dislodge thrips and prevent thrips escaping. Action level: Control thrips if more than four to six thrips are found per flower.
Western flower thrips
Pest status: Potentially a devastating pest in peanuts. WFT is now widespread and regular in Australia but has not yet (as at mid-2005) severely affected peanuts. Peanuts are susceptible to tomato spotted wilt virus (TSWV), and since WFT’s detection in Australian peanuts the incidence of TSWV in that crop has increased, but not yet to alarming levels. While other thrips species also vector TSWV in Australia (e.g. Frankliniella schultzei), it is thought that WFT vectors TSWV more efficiently than these species. Identification: Frankliniella occidentalis is similar to other thrips with the naked eye and must be examined microscopically to be correctly identified. The presence of WFT in peanuts is evidenced by TSWV symptoms, such as ring spots on the leaves of infected plants and stunted plant growth. May be confused with: Thrips species can only be determined microscopically. Thrips damage can be distinguished from spider mite damage by the appearance of liquid faecal deposits, which cause dark green speckling, whereas spider mites produce black granules.
Fig. 7.63. Thrips damage to mung bean pod, showing characteristic distortion. (DPI&F Qld)
Host range: Over 200 plant species from 62 families, including many field, horticultural crops and weeds are hosts. Summer pulse hosts include peanuts, soy beans, mung beans and navy beans. Other hosts include capsicum, carnations cotton, cucumbers, lettuce, olives, 207
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roses, safflower, stone fruit, sweet peas and tomatoes.
Chemical control: Chemical control of WFT in summer pulses is ulikely to be cost-effective.
Life cycle: Eggs are laid in slits made in leaves and growing points. There are two larval stages as well as a pre-pupal and pupal stage. Nymphs and adults feed in growing points and inside flowers. Pupation occurs in the soil. The life cycle can be completed in as little as 10 days at 20°C. Populations decline when temperatures exceed 30°C.
Cultural control: Late peanut plantings may help avoid WFT damage.
Risk period: Crops can be infested during the early vegetative stage but severe TSWV symptoms may not occur until much later in the crop’s development. As a rule, the earlier the thrips attack, the greater the risk of severe TSWV damage. Damage: Symptoms of F. occidentalis feeding include discoloration and indentation of the leaf surface. The pattern of damage is coarser than damage by Thrips tabaci. WFT damage to buds and flowers can result in deformed pods. TSWV symptoms include distortion and mottling of leaves and stunted plants (Fig. 7.64). Affected plants may have halo rings on leaves. In peanut crops, TSWV can potentially lead to plant collapse and death; however, TSWV symptoms of this severity have not been reported to date in Australia. Monitoring: Inspect vegetative terminals and flowers for thrips. Severe and widespread tomato spotted wilt virus (TSWV) symptoms in peanuts may indicate the presence of WFT. Action level: Very few WFT, which are resistant to most insecticides, are required to spread TSWV.
Fig. 7.64. Two peanut plants showing symptoms of tomato spotted wilt: stunting of the plants and the yellowing and distortion of the leaves compared with the surrounding healthy plants. (H. Brier)
Conservation of natural enemies: Using soft pesticides against other pests may conserve other thrips species which, by competing with WFT, may slow the latter’s population growth.
Scarab beetles (Scarabaeidae) Peanut scarabs, Heteronyx spp. and Sericestihis spp. Black sunflower scarab, Pseudoheteronyx basicollis Linnaeus Distribution: Peanut-growing regions with Ferrosol soils in north-eastern Australia including the South Burnett, Atherton Tableland, and the Clifton/Nobby region of the eastern Darling Downs. The major species in the south Burnett in order of abundance are Heteronyx piceus Blanchard, H. rugosipennis Macleay, Sericestihis ino (Balckburn) and S. suturalis (Macleay). Pseudoheteronyx basicollis L. is the dominant species on the Darling Downs but Heteronyx sp. near rugosipennis also occurs in significant numbers in this region. Pest status: Major, widespread and regular (in their preferred regions). Identification: Peanut scarab eggs are cream and spherical and relatively large (1.5 mm diameter, Fig. 7.65) Larvae are creamy white and C-shaped, with well-developed thoracic legs, a brown head capsule, and well-developed mandibles (Fig. 7.67). Fully grown extended larvae reach 25 mm in length and should be handled with care as they can inflict a very painful bite. Larvae within regions can be distinguished by their palidial patterns. The
Fig. 7.65. Heteronyx piceus eggs laid under peanut bush, Tingoora, Qld. (2 × 1.5 mm). (DPI&F Qld: J. Wessels)
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Life cycle: These are native 1-year life cycle species that have adapted to peanuts. The beetles lay their eggs under peanut seedlings. Young larvae feed on peanut roots, but older larvae attack the pods. Larvae overwinter in the soil and pupate in the spring. Beetles emerge in the late spring and early summer after heavy storm rains. Risk conditions: Peanut seedlings are at greatest risk after storm rains, which can trigger beetle emergence.
Fig. 7.66. Peanut scarab Heteronyx rugosipennis (17 mm long), Tingoora, Qld. (DPI&F Qld: J. Wessels)
palidium is a group of short, flattened spines on the larvae’s rear ends. Heteronyx piceus larvae have a distinctive Y-shaped palidium, while that of H. rugosipennis is V shaped with stout spines. Sericestihis spp. larvae also have V-shaped palidia but their spines are short. Pseudoheteronyx basicollis and Heteronyx sp. nr. rugosipennis larvae have Y- and V-shaped palidia, respectively. Heteronyx and Sericestihis adults are small brown scarab beetles about 10–15 m long. Heteronyx piceus has wing covers covered with fine short pale prostrate hairs and are 12 mm long. Heteronyx rugosipennis beetles (Fig. 7.66) are slightly larger than H. piceus and have elytra (wing covers) covered with more visible and erect pale hairs. Sericestihis adults are sparsely haired and often have a purple sheen on the wing covers. Sericestihis ino is relatively small at 10 mm long, while S. suturalis reaches 15 mm. Pseudoheteronyx basicollis beetles are 14 mm long and are shiny black with very short hairs. Heteronyx sp. near rugosipennis adults are dull black with numerous pale hairs.
Damage: Adults feed on peanut leaves, feeding inwards from the leaf edge. Fresh dark sap from feeding wounds indicates very recent damage. Young larvae feed on peanut roots, but older larvae attack the pods. Young pods may be totally consumed, while kernels in older pods will be partially eaten out. Larvae typically make a large hole in the distal end of pods (Fig. 7.67). Maturing pods with hardening shells may only suffer surface scarification (sometimes called scarabification). Heavily infested crops may suffer over 30% yield loss. Damaged pods are also prone to invasion by species of Aspergillus fungi, which can produce the highly carcinogenic aflatoxin. Monitoring: Look for freshly damaged leaves with the typical scarab-feeding pattern. Look for scarabs and eggs under peanut seedlings. In crops approaching maturity, look for damaged pods and scarab larvae feeding in the podding zone.
May be confused with: Larvae and adults are easily confused with other non-peanut-attacking scarabs that may have similar palidial patterns. Host range: Peanuts, and also sunflowers for P. basicollis. Peanut scarab species occur in pasture.
Fig. 7.67. Heteronyx piceus larva and typical damage to peanut pod. Such damage allows the growth of aflotoxin-producing fungi in the peanut pod. (DPI&F Qld: J. Wessels)
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Distribution: Found in most regions but more common in regions with heavy cracking soils. Pest status: Major, restricted to heavy soils, irregular. Identification: Adults are dark slender beetles up to 25 mm long with a spine at the rear of the pronotum. Adults make a loud clicking sound when handled and the spines can pinch unwary fingers. The larvae are soft-bodied and cream and can reach 35 mm in length. They have a distinctive chisel-shaped head. Fig. 7.68. Peanut scarab parasitoid Anthobosca sp. (Hymenoptera: Tiphiidae) (10 mm). This specimen emerged in spring from an overwintering Heteronyx piceus larva, Tingoora, Qld. (J. Wessels)
Action level: No thresholds are set because control is not feasible once a crop is infested with larvae. If damage is anticipated, apply soilincorporated pesticides at planting to control egg-laying adults and newly hatched larvae. Chemical control: A granular pesticide is registered for peanut scarabs in peanuts and gives cost-effective control. Cultural control: Avoid successive peanut crops and eliminate peanut volunteers from rotation crops. Plant peanut crops as far away as possible from where the previous year’s crop was grown. Peanut crops on the upper slopes and hill tops are at greater risk from Heteronyx piceus attack than those on lower slopes and valleys. Conservation of natural enemies: Peanut scarab larvae are sometimes parasitised by wasps (Fig. 7.68), but parasitism levels are usually low (< 1%). Larvae are sometimes killed by the Cordiceps fungus, which produces spectacular ‘horn like’ growths from their hosts (Fig. 15.22). Such infections, however, are not common.
May be confused with: Larvae can be confused with false wireworm larvae but the latter are hard-bodied with a rounded head. Holes bored into peanut pods may be confused with those made by etiella caterpillars, but the latter usually only make one hole per pod, and leave characteristic webbing and pale caterpillar frass. Host range: All field crops including summer pulses. Risk period: Crops are at greatest risk during the seedling stage but larvae may attack peanuts at podding. Damage: Larvae attack germinating seeds and attack seedling roots and shoots causing plant death. Larvae may also attack peanut pods, often boring numerous holes per pod (Fig. 7.69). Monitoring: Use germinating seed baits to detect adults, and dig and sieve soil to find larvae. Also monitor crops after planting for poor emergence and seedling deaths. Action level: Take action if significant damage or significant wireworm populations are present.
C L I C K B E E T L E S ( E L AT E R I DA E )
Sugarcane wireworm Agrypnus variabilis (Candèze) Coleoptera: Elateridae N (Main entry in Chapter 2 Cereals.)
Fig. 7.69. A true wireworm larva in peanut pod (12 mm). (DPI&F Qld: J. Wessels)
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Chemical control: Check for the latest APVMA registrations.
T E N E B R I O N I D S ( T E N E B R I O N I DA E )
Cultural control: Replant badly affected crops with treated seed.
False wireworms
Conservation of Natural enemies: Brown predatory earwigs attack wireworm larvae.
Gonocephalum sp. and Pterohelaeus sp.
Coleoptera: Tenebrionidae N (Main entry in Chapter 5 Oilseeds.)
S O L D I E R B E E T L E S ( C A N T H A R I DA E )
Plague soldier beetles Coleoptera: Cantharidae Chauliognathus lugubris (Fabricius) and other Chauliognathus species. Distribution: Probably in most summer pulse regions. Pest status: Minor, widespread, irregular. Identification: Adults are elongated flattened beetles with moderately long antennae and reaching 15 mm in length. They are frequently dark metallic green with yellow undersides and have a yellow or red band behind the head (Fig. 7.70). The soil-dwelling larvae are flat and dark-coloured and covered with hairs that give them a ‘velvety’ appearance. Soldier beetle larvae are similar to ladybird larvae but are larger and more slender. Life cycle: The larvae are soil-dwelling. Adults often congregate in large swarms. Damage: Large swarms of adults occasionally invade soy beans, feeding on the leaves. However, the soil-dwelling larvae are predacious, reputedly feeding on Lepidoptera eggs, caterpillars and adults.
Distribution: Common in summer pulses planted on heavy cracking soils. Pest status: Minor, restricted, irregular. Host range: All field crops including summer pulses. Risk period: Crops are at greatest risk during the seedling stage. Larvae and adults in the crop area prior to planting cause the damage. Damage: Larvae attack germinating seeds and attack seedling roots and shoots. Adults chewing on stems at ground level can ringbark plants or cut through them completely. Badly affected crops may have to be re-planted. Monitoring: Use germinating seed baits to detect adults, and dig and sieve soil to find larvae. Also monitor crops after planting for poor emergence and seedling deaths. Action level: Take action if significant damage or significant cricket populations are present. Chemical control: Check for the latest APVMA registrations. Cultural control: Re-plant badly affected crops with treated seed. Conservation of Natural enemies: Brown predatory earwigs attack false wireworm larvae.
Monitoring: Use a beat sheet. Chemical control: No pesticides are registered.
LO N G I C O R N B E E T L E S ( C E R A M B YC I DA E )
Lucerne crownborer Zygrita diva Thomson Coleoptera: Cerambycidae Fig. 7.70. Soldier beetle, Chauliognathus sp., (15 mm long) on soy beans, Kingaroy, Qld. (H. Brier)
Distribution: A tropical–subtropical species native to Australia. Reported from all Australian states except SA, Vic. and Tas. 2 11
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Fig. 7.71. A lucerne crownborer larva (12 mm) tunnelling in the pith of a soybean stem. (DPI&F Qld: J. Wessels)
Fig. 7.73. Corrhenes stigmatica (Pascoe) (10 mm) on soy beans. (DPI&F Qld: J. Wessels)
Pest status: Major in the tropics, minor in more southern regions, widespread, regular.
similarly sized but the less common Corrhenes stigmatica (Pascoe) (Fig. 7.73). Adults of C. stigmatica are pale mottled brown and lack the dark wing cover spots of Z. diva.
Identification: The cream larvae can reach 25 mm in length and have a wide head. They have marked constrictions between the body segments (Fig. 7.71). Adults are 15 mm long and are typically bright orange with black legs and long black antennae with white bands. They usually have a pair of prominent large black spots on their wing covers but may have up to three pairs of spots or bands (Fig. 7.72). Some specimens may have the black bands edged with white. Specimens with no spots or with totally black wing covers are infrequently encountered. The orange scales often rub off the wing covers of older specimens leaving a base colour of grey brown. May be confused with: Zygrita adults and larvae may be confused with those of the
Fig. 7.72. A lucerne crownborer with two black spots on the bright orange elytra (10 mm). (DPI&F Qld: J. Wessels)
Host range: Soy beans and lucerne. Also found in phasey bean and sesbania. Life cycle: Adults lay the eggs in the stems of young soy beans. Larvae tunnel through the pith, but usually pupate in the taproot. Risk conditions: Soybean crops in the tropics, or growing in abnormally ‘hot’ summers, or in close proximity to lucerne are at greatest risk. Proximity to lucerne increases the risk of early infestation and hence earlier pupation and girdling. Damage: In some crops, over 80% of plants may be infested. Both Z. diva and C. stigmatica cause similar damage. Larval feeding has little impact on yield, but prior to pupating larvae ‘girdle’ or ringbark stems internally in order to plug the stem cavity above the pupal chamber. This cuts the vascular tissue and causes plant death above the girdle. Crops with thin plant stands may lodge before harvest as they are not supported by adjoining plants. In southern Qld, this usually occurs after seeds are fully developed (physiological maturity) with no yield loss. In tropical regions, larval development is more rapid, with considerable crop losses. Crownborers are very damaging to CV ‘edamae’ soy beans where green immature pods are harvested by mechanical pod-pluckers. The
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Fig. 7.74. Iphiaulax sp. (7 mm), a braconid parasitoid of lucerne crownborer Zygrita diva. (J. Wessels)
stems of infested plants are sufficiently weakened to snap off and contaminate the harvested product. Monitoring: Break open stems and look for larvae and eaten out and brown discoloured pith. Action level: There are no specified thresholds. Chemical control: Chemical control is incompatible with IPM and is not cost-effective, as larvae in the stems are protected from insecticide. Cultural control: Avoid planting susceptible crops close to lucerne. If in an at-risk region, consider later plantings to shorten crop development. In the tropics, consider winter plantings. Avoid thin plant stands to reduce the lodging of damaged plants. Conservation of natural enemies: Because of their sheltered feeding sites, crownborer larvae are largely immune from attack by predators and parasites. However, the small braconid wasp (Iphiaulax sp., Fig. 7.74) has been recorded as parasitising Z. diva.
LE AF BEETLES, SEED WEE VILS ( C H R Y S O M E L I DA E )
Bean bruchid Acanthoscelides obtectus (Say) Coleoptera: Chrysomelidae Distribution: Acanthoscelides obtectus is a South American species that is now a cosmopolitan
Fig. 7.75. Bean bruchid (3.5 mm long) on navy bean seed. (DPI&F Qld: P. Collins)
pest throughout the tropics, subtropics and warmer temperate regions. In Australia, A. obtectus is established in Australia as far south at least as central NSW. Pest status: Acanthoscelides obtectus is a major, widelydistributed and regularly occurring pest of stored navy beans and lima beans. While predominantly a problem in storage, outbreaks are often the result of infestation of crops prior to harvest. Identification: Acanthoscelides obtectus adults are of typical bruchid shape. They are 3.5 mm long and are grey–brown with faint speckled markings (Fig. 7.75). The loosely laid eggs are small and white, and are not glued to seeds or pods as are the eggs of Callosobruchus spp. bruchids. Larvae are pale cream and are scarabaeiform with reduced legs. Seeds with pupae inside have translucent windows through which emerging adults will escape. May be confused with: Bean bruchid adults are similar in outline to many other bruchids; however, they may be distinguished by having a single row of three to four closelylocated spines (one large, two to three small) on the outer end of each hind femur. Host range: Navy beans and lima beans. Also reported in field peas. Life cycle: Adults can infest seeds in the field or in storage. Eggs are laid on maturing pods in the field, but in storage are laid loosely among seeds. Hatching larvae burrow into the seed and 213
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complete their development (including pupation) within a seed before emerging as adults. Adults do not feed on seeds, but may feed on pollen and nectar in flowers. Bean bruchids breed rapidly in storage and can complete a life cycle in as little as 23 days; however, they can continue to breed slowly at temperatures as low as 18°C. Risk period: As for C. maculatus. Damage: As for C. maculatus, except that A. obtectus eggs are not conspicuous in storage as they are laid loosely among seeds. Monitoring and action level: As for C. maculatus, except that even lower aeration temperatures (15°C) would be required because of A. obtectus’s greater cold tolerance. Chemical control: As for C. maculatus. Phosphine fumigation of harvested seed is the preferred option.
Bruchidius mackenziei Kingsolver. Coleoptera: Chrysomelidae
Fig. 7.76. Bruchidius mackenziei (2.5 mm long) on soybean seed. (DPI&F Qld: J. Wessels)
of maturing soybean crops. Larvae and pupae are often found in seeds after harvest’ however, overseas Bruchidius spp. adults do not re-infest stored grain and B. mackenziei eggs have not yet been found in stored soybean seed. The emergence of adults as late as 2–3 months post harvest may be due to slowed larval development in harvested seed with low moisture content.
In view of its restricted host range, ‘soybean bruchid’ would be an appropriate common name.
Risk period: Late maturing crops are at greatest risk.
Distribution: Bruchidius mackenziei is a native bruchid recorded from soy beans in the Burnett and Darling Downs regions in Qld.
Damage: Similar to that caused by other bruchids. Emergence windows may be seen in still-infested seeds. To date, eggs have not been seen on infested seeds; however, the mere presence of bruchids is off-putting for customers who fear re-infestation in their storages.
Pest status: A pest of increasing significance in soy beans. Probably not a true stored-product pest but often found in soy beans at or after harvest. Identification: Adults are 2.5 mm long and are brown with speckled markings and sometimes paler patches on the elytra (Fig. 7.76). The antennae of the males are strongly pectinate. B. mackenziei has a large dark broad-based spine on each inner ventral femur ridge. The rear end of the spine is at 90° to the femur. May be confused with: Adults are similar to other bruchids of summer pulses, but have only one spine per femur as opposed to three to four for A. obtectus, and two spines for Callosobruchus spp. Host range: Reported in Australia only from soy beans. Life cycle: B. mackenziei is probably not a true stored-product pest. Adults lay eggs on the pods
Monitoring and action level: As for C. maculatus. Adults may be detectable in the field. Chemical control: Phosphine fumigation should kill any larvae still present in seed post harvest and reduce subsequent adult emergence; however, there is not yet any phosphine efficacy data specific to B. mackenziei.
Cowpea bruchid Callosobruchus maculatus (Fabricius) Coleoptera: F. Chrysomelidae, sub-family Bruchinae (formerly Family Bruchidae) Related to the less-common Callosobruchus species in Australia includes C. chinensis (adzuki bruchid) and C. phaseoli. Cowpea bruchids are
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sometimes called cowpea weevils but they are not true weevils. Distribution: Callosobruchus spp. bruchids are cosmopolitan pests throughout the tropics and subtropics. In Australia, C. maculatus is established south to at least central NSW. Pest status: Callosobruchus spp. bruchids, particularly C. maculatus, are major, widely distributed and regularly occurring pests of stored summer pulses. While Callosobruchus damage occurs in stored pulses, outbreaks can be initiated either before or after harvest but are usually not detected until well after harvest. Identification: Bruchids are small brown beetles with a rounded oval-outlined body that tapers towards the head, shortened elytra, and long antennae and legs (Fig. 7.77). The hind femurs are frequently enlarged. Bruchids do not have the elongated snout of true weevils. Cowpea bruchids (C. maculatus) are 3 mm long and typically orange–brown with dark markings. They have a large dark spot midway down the outer edge of each elytra and a stubby crossshaped orange patch on their back. The eggs are small and white and, despite being only 0.6 mm long, are readily visible on the surface of seeds (to which they are glued). The pale cream larvae are scarabaeiform with reduced legs and are rarely seen as they feed within seeds. Seeds with pupae inside have translucent windows through which emerging adults will emerge. May be confused with: Callosobruchus maculatus may be confused with other Callosobruchus spp.
Fig. 7.77. Cowpea bruchid (3 mm long) on mung bean seed. The circular exit hole is made by an emerging adult. White eggs are conspicuous on the seed surfaces. (DPI&F Qld: J. Wessels)
bruchids and with bruchids in other genera. Bruchid colouration may be variable. The most reliable genus and species indicators are spines and teeth on the underside of the femurs of the hind legs. Callosobruchus spp. bruchids have a single spine or tooth at the apical end of both the ventral femur ridges. In C. maculatus, the spines are triangular, and the outer spines may be obscured by hairs. For C. chinensis and C. phaseoli, the inner apical spines are slender and are longer than the broad-based outer spines. C. chinensis has a colour pattern similar to that of C. maculatus but is stouter with shorter elytra. C. phaseoli is more speckled with two dark patches on the lower elytra. Host range: Cowpea bruchids attack cowpeas, field pea, mung beans and soy beans. Chickpea and navy beans are less-favoured hosts. While chickpea acid usually protects chickpea pods in the field from attack, cowpea bruchids can attack chickpea seed in storage. Adzuki bruchids, C. chinensis, also attack chickpea, as well as cowpeas, mung beans and siratro. C. phaseoli mainly attacks Dolichos but sometimes also cowpeas, mung beans and navy beans. Life cycle: Callosobruchus adults can infest seeds in the field or in storage. Eggs are laid on maturing pods in the field, but in storage are laid directly on seeds. Eggs are firmly attached (glued) to pods and seeds. Hatching larvae burrow into the seed and complete their development (including pupation) within a seed before emerging as adults. Prior to pupation, larvae excavate an emergence tunnel to the surface of the seed, leaving a translucent but intact window in the seed coat. Adults do not feed on seeds, but may feed on pollen and nectar in flowers. Cowpea bruchids breed rapidly in stored seed and can complete a life cycle in as little as 28 days (at 30°C). Adults live only 10–12 days and females lay about 100 eggs. Adults are strong fliers and can travel 2–3 km. Infestations are often not detected until well after harvest. Risk period: Crops are at greatest risk during late podding. Seed in storage is at greatest risk if infested prior to harvest, if stored in bruchidinfested premises, if stored at too high moisture, if stored in bags, or if stored for an extended period of time (> 4 months). Damage: Larvae feed and pupate within the seed, eating large cavities inside the seed. 215
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Emerging adults make a neat circular exit hole in seeds. Seeds infested in storage are often covered with conspicuous white eggs. Heavy bruchid infestations can lead to the overheating of seed in storage, resulting in reduced quality (in addition to physical damage) and moulds. Damaged seed has poor germination, and hence sprouting. Monitoring: Inspect all seed carefully at intake, taking spear samples if necessary. Also look for adults on the surface of stored seed. Seed should be inspected for insects once a fortnight in summer and once a month in winter. Numbers present at intake may be very low and practically undetectable. Action level: Even an extremely low bruchid population at intake (e.g. undetectable levels of two bruchids per tonne or 0.00001% of seeds infested) could result in major damage (q 50% of seeds damaged) within 4–6 months of intake. Ideally, all seed should be treated at intake, particularly for pulse types and in regions with regular bruchid occurrence. Chemical control: Controlling cowpeas and other similar bruchids in the field is not effective and may not protect from storage damage. Anything less than 100% in-field control of cowpea bruchids could still result in significant bruchid damage in storage.
temperature and moisture should be monitored regularly during aeration. Cowpea bruchid development will cease at temperatures below 20°C. Grain in a good insulator and non-aerated pulse seed retains its temperature in storage during winter.
Redshouldered leaf beetle or Monolepta Monolepta australis (Jacoby) Coleoptera: Chyrsomelidae Distribution: Throughout northern Australia. Particularly abundant in cane-growing coastal regions such as Bundaberg in south-east Qld. Pest status: Moderate, widespread, regular. Identification: The beetles are 6 mm long and yellow, with a dark red (purple) band across the shoulders and two purple spots on the ends of the wing covers (Fig. 7.78). The flaccid yellowish eggs are small (< 1 mm across) and oval. The larvae are white, slightly flattened with sclerotised (hard brown) plates at both ends, and reach 5 mm in length. May be confused with: The adults are easily distinguished from other beetles (including ladybirds) by their bright but distinctive colouration.
Treating seed at intake is effective in minimising bruchid damage in storage. Phosphine fumigation is approved for bruchid control in pulses and all ‘at risk’ loads should be treated at intake, whether bruchids are detected or not. Correct fumigation is vital for successful bruchid control, with best results achieved in sealed (pressure-tested, gas-tight) silos (< 200 tonnes) or in sealed bag stacks. Seed should be fumigated for at least 10 days and tablets applied at recommended doses (two tablets per tonne capacity of wheat or three tablets per 2 m3). Remember that the dose relates to ‘whole of silo capacity’; that is, air space plus grain space. Best practice avoids mixing tablets with grain, as the residual tablet dust is toxic. Spread tablets in trays hung in silo headspace, or placed in bag stacks. Phosphine ‘blankets’ or ‘belts’ are an alternative to tablets.
Host range: Adults are polyphagous. Of the summer legumes, soy beans, navy beans and mung beans are particularly attractive hosts. Other hosts include avocado, cotton, lychee, macadamia, mango, strawberry, and numerous ornamentals. Larvae feed underground on the roots of sugarcane and pasture grasses.
Cultural control: Bruchid development in storage can be slowed by cooling aeration. Seed
Fig. 7.78. Redshouldered leaf beetle (5 mm), Bundaberg, Qld. (DPI&F Qld: J. Wessels)
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Chemical control: Monolepta are readily controlled with many (but not all) caterpillar pesticides already registered in soy beans. Consult your local agronomist or entomologists for advice about the latest pesticide options. Cultural control: Plant legume crops away from susceptible larval hosts if possible. Conservation of natural enemies: There are no known predators that are effective against high redshouldered leaf beetle. As infestations are often patchy, consider spot-spraying or perimeter spraying where numbers are highest, leaving most of the crop unsprayed.
Fig. 7.79. Redshouldered leaf beetles shredding soybean leaves, Bundaberg, Qld. (H. Brier)
Life cycle: Eggs are laid in the soil surface, mainly in pastures and in sugarcane. The larvae feed on grass roots and pupate in the soil. The life cycle takes about 2 months during summer and there are three to four generations annually. Adults usually emerge from the soil after heavy rains following a dry spell. If larval populations in the soil are high, the multitude of emerging beetles will form an aggregation and swarms may migrate into nearby soybean crops. Risk period: Soy beans are at greatest risk during flowering. Infestations are likely after heavy rainfall events. Damage: Monolepta attack leaves and flowers. High populations (e.g. > 50 per square metre) will shred leaves and denude crops of flowers (Fig. 7.79). Monitoring: Monolepta are readily assessed visually or with a beat sheet; however, the adults are extremely flighty and numbers are very difficult to accurately count on a beet sheet, as the beetles can literally ‘explode’ off the sheet in seconds. Estimate the number of groups of five or 10 beetles on the sheet to get a ‘ball park’ population estimate. Check crops after heavy rainfall events. Action level: Populations greater than 20 per square metre most likely cause significant damage in flowering crops. Defoliation thresholds are as for defoliating caterpillars.
B R E N T I DA E
Pod weevils Apion spp. including Apion congestum Lea Coleoptera: Brentidae Distribution: Pod weevils occur throughout Australia but Apion species attacking summer pulses are probably restricted to the tropics and subtropics. Apion sp. adults have been recorded on summer pulses (particularly mung beans) in the South Burnett region of south-east Qld. Apion congestum has been recorded in mung beans at Kununurra, WA, and from Tolga and Biloela in Qld. Pest status: Probably minor, widespread, regular. Apion spp. are regarded as major pigeon pea pests in India and Africa. Identification: Adults are small shiny black weevils (2–3 mm) with bulbous abdomens and typical weevil-type elongated heads (Fig. 7.80). The larvae are creamy white and of typical weevil larva outline. May be confused with: The distinctive adults are unlikely to be confused with other pests. Host range: Commonly encountered in mung beans, adzuki beans and cowpeas (all Vigna spp.), and also reported in pigeon peas and n avy beans. Life cycle: Larvae and adults of Apion sp. are reported feeding on mung bean flowers but other details on biology are lacking. 2 17
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preceding crop) results in plant deaths. Root damage in more advanced crops may reduce plant vigour. Monitoring: As for whitefringed weevils. Cultural control: Avoid successive plantings of amnemus weevil-susceptible crops such as soy beans in ground previously sown to pasture legumes.
Whitefringed weevil Fig. 7.80. Pod weevil, Apion sp., (2.5 mm) on mung bean buds. (J. Wessels)
Risk period: Crops are at greatest risk during flowering and early podding. Damage: Larvae and adults may attack flowers and most likely young seeds. Monitoring: Open flowers and look for the distinctive adults. Adults may also be seen on the outside of flowers. Cultural control: Vigorously growing crops should be better able to compensate for any damage caused by this very minor pest.
W E E V I L S ( C U RC U L I O N I DA E )
Amnemus weevil Amnemus quadrituberculatus (Boheman) Coleoptera: Cuculionidae N (Main entry in Chapter 13 Pastures—summer
rainfall.)
Pest status on summer pulses: Moderate, restricted, irregular. Identification and life history: See Chapter 13. Host range: Legumes; in summer pulses, reported mainly in soy beans (Glycine max). Risk conditions: Crops are at greatest risk if they follow another amnemus-susceptible host. Damage: Adults may feed on leaves but this is usually of little consequence. The symptoms are as for whitefringed weevil. Feeding on seedling tap roots by medium to large larvae (from a
Naupactus leucoloma (Boheman) Coleoptera: Cuculionidae Previously classified as Graphognathus leucoloma. N (Main entry Chapter 15 Pastures—winter rainfall.)
Pest status on summer pulses: Moderate, widespread and regular. Identification: See Chapter 15. May be confused with: Adult whitefringed weevils may be confused with sitona and vegetable weevils but are larger and more prominently striped. Whitefringed weevil larvae are difficult to distinguish from those of other soil-dwelling weevils. Leaf damage is similar to that caused by peanut scarabs. Host range: Pulses attacked include peanuts and chickpea. Other crops attacked include lucerne, lupins, potatoes and pumpkins. Life cycle: In northern Australia, this species has a 1-year life cycle. Eggs are laid in sticky clumps in plant debris at the base of plants. Hatching larvae burrow downwards in the soil attacking plant roots and pods. Risk conditions: Crops are at greatest risk if they follow another whitefringed weevilsusceptible crop. Damage: Adults may feed on leaves making ‘scalloped’ chewings in from the leaf margins. Leaf-feeding is usually of little consequence. Feeding on seedling tap roots by medium to large larvae (from a preceding crop) can sever roots resulting in plant death. In severely infested crops, plant numbers may be reduced by over 50%. Root damage in more advanced crops may reduce plant vigour. Damage to peanut pods is similar to peanut scarab damage but the holes
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made by weevil larvae are smaller than those made by scarabs. Pod damage reduces yields and increases the risk of aflatoxin. Monitoring: Look for leaf damage in young crops. If significant leaf damage is observed, look for adults at the base of plants. If seedling deaths occur, look for larvae under plants in the soil. Action level: Take action if significant adult activity is observed in seedling crops. Chemical control: Chemical control of whitefringed weevil adults in peanuts may be cost-effective. Foliar sprays are ineffective against larvae. Cultural control: Avoid successive plantings of crops susceptible to whitefringed weevils, such as peanuts following potatoes (as occurs in north Qld), or chickpeas following peanuts (as occurs in southern Qld), or peanuts following lucerne. By far the worst whitefringed weevil damage occurs where large larvae from preceding susceptible crops attack seedlings in following susceptible crops. Crops close to lucerne are also at risk from weevils walking from that crop.
Fig. 7.81. Sciarid larvae (2 mm) and scarification damage to a peanut pod. Note the larvae’s dark sclerotised head. (J. Wessels)
species such as Bradysia sp. also attack plant roots and peanut pods. Sciarid larvae can be pests in irrigated and glasshouse grown peanuts (because of the humid moist environment). Risk period: Podding. Damage: Larvae scarify the surface of pods but the wet environment favouring sciarids is unfavourable for the development of aflatoxin. Monitoring: Dig up pods carefully and look for damage and associated larvae. Action level: None specified.
F U N G U S G N AT S ( S C I A R I DA E )
Sciarids or fungus gnats Bradysia sp. Diptera: Sciaridae
Chemical control: No registered products, and probably none needed.
L E A FM I N E R F L I E S ( AG R O M Y Z I DA E )
Distribution: Most likely widespread but only active in irrigated crops.
Beanfly
Pest status: Minor.
Ophiomyia phaseoli (Tryon) Diptera: Agromyzidae
Identification: The larvae are small (3–5 mm), slender and pale with a dark sclerotised head (Fig. 7.81). The gut contents are often visible through their skin. The adults are small dark narrow-bodied flies and are only 3 mm long. May be confused with: Sciarid larvae many be confused with small Etiella larvae which are also pale with a dark head but which have prolegs. Damage (scarification) is similar to that caused by other soil pests such as peanut scarabs. Host range: Recorded from irrigated peanuts. Life cycle: The larvae are gregarious and live in moist soil and feed on organic matter. Some
Distribution: A worldwide pest of legumes. In Australia, beanfly is most common in tropical and subtropical coastal and sub-coastal regions. Pest status: Major, widespread and regular in its preferred range. Identification: Adults are small (3 mm long) and shiny black with clear wings (Fig. 7.82). The larvae (or maggots) are cream with dark mouthparts and reach 3 mm in length. Pupae are small, brown and cylindrical with rounded ends. May be confused with: Soybean podfly. 219
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Fig. 7.82. Adult beanfly (3 mm). (DPI&F Qld: J. Wessels)
Host range: Preferred crops are navy beans, adzuki beans and mung beans. They are rarely a pest of soy beans. Favoured non-crop hosts include phasey beans and pasture legumes. Life cycle: Female flies lay their eggs in young leaves. Upon hatching, larvae tunnel their way to the leaf mid-vein, make their way down the petiole and stem and pupate in the lower stem. Risk period: Crops are at greatest risk for 3–4 weeks from emergence, but later crops are sometimes attacked. Damage: Larval-tunnelling damages the plant’s vascular tissue, causing seedling death, reduced plant vigour and petiole droop in older crops (Fig. 7.83).
Fig. 7.84. Beanfly oviposition stings on navy bean leaf. (DPI&F Qld: J. Wessels)
Monitoring: Monitor seedling crops twice weekly. Look for the distinctive pale oviposition pinprick windows in the leaves (Fig. 7.84) and for larval tunnelling at the base of petioles and in the stems. Look for pupae and damaged stem tissue in the lower stems. Look also for adult flies. Action level: More than one larval tunnel per plant. Chemical control: Control is readily achieved in young crops with systemic pesticides, but repeat sprays (within 7 days) may be required for heavy infestations. Sprays against beanfly in mature crops may not be cost-effective. Cultural control: Ensure cropping areas are free of weed hosts such as phasey bean and volunteer crop legumes.
Soybean podfly Melanagromyza sojae (Zehntner) Diptera: Agromyzidae Distribution: A tropical species found in Africa, Asia, Australia and the western Pacific. In Australia, soybean podfly is most common in the tropics and subtropics. Pest status: Minor, widespread and irregular. Identification: Adults are small (3.5 mm long) and shiny black with clear wings. The larvae (or maggots) are cream with dark mouthparts. Fig. 7.83. Navy bean plant severely damaged by late beanfly infestation. (DPI&F Qld: J. Wessels)
May be confused with: Beanfly. Host range: Soy beans.
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Life cycle: Eggs are laid on pods. Larvae bore into pods and feed on developing seeds. Risk period: During podfill. Damage: Larvae infrequently damage seeds. Monitoring: Open pods to assess larval damage. Action level: No specified thresholds. Chemical control: No registered pesticides. It is doubtful whether pesticides could give effective control without compromising residue levels in the harvested seed. Cultural control: No recommended control. Conservation of natural enemies: Not applicable.
L E A F B LOTC H M I N E R S ( G R AC I L L A R I DA E )
Soybean leafminer Porphyrosela aglaozona (Meyrick) Lepidoptera: Gracillariidae
Moths are 2 mm long and are orangey-brown with dark and white markings. Their wings are folded above the body when at rest. May be confused with: Larval damage (leaf mines) may be confused with that made by young soybean moth larvae. Larvae of the two species can be distinguished by differences in body shape. Host range: Soy beans. Life cycle: Larvae spend their entire life feeding within the leaf (in leaf mines). Risk period: Crops are theoretically at greatest risk during podfill when they are least tolerant of defoliation. Damage: Larvae feed within the leaf-forming transparent mines or blisters (Fig. 7.85). Damage is cosmetic. Monitoring: Look for leaf mines. Open leaf mines to determine if they are made by soybean leafminer or by soybean moth larvae. Action level: No specific thresholds. Thresholds are therefore based on tolerable defoliation; that is, 33–40% pre flowering and 15–20% during early podfill.
Distribution: Native to Australia and reported from all soybean growing states (Qld, NSW, Victoria).
Chemical control: No products specifically registered or needed for this pest.
Pest status: Minor, widespread, irregular.
Cultural control and conservation of natural enemies: As for other Lepidoptera.
Identification: A very small species. Larvae reach 3 mm in length and are cream with a very small pale head. Their body shape is distinctive, being widest just behind the head and tapering towards the rear with deep constrictions between the body segments (Fig. 7.85).
G E L E C H I I DA E
Soybean moth Aproaerema simplexella (Walker) Lepidoptera: Gelechiidae Previously known as Stomopteryx simplexella. Distribution: Native to Australia and reported from most states. Very similar Aproaerema species occur in Asia and Europe. Pest status: Usually minor, widespread, regular. This pest occasionally occurs (once every 20 years) in very high numbers (1000s of larvae per square metre), which defoliate infested crops.
Fig. 7.85. Soybean leafminer larvae (2 mm). (DPI&F Qld: J. Wessels)
Identification: The small, elongated eggs are often laid on leaf veins. Larvae are pale green 221
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Fig. 7.88. Severe soybean moth damage, Kingaroy, Qld, 1988. (H. Brier) Fig. 7.86. Large soybean moth larva (7 mm long) showing typical feeding damage. (DPI&F Qld: J. Wessels)
with a dark head and only reach 7 mm in length (Fig. 7.86). Infested leaves are often crinkled and pulled together in the middle. The small narrow winged moths (only 6 mm long) are dark with a transverse white band (Fig. 7.87). May be confused with: Larvae may be confused with young soybean leafminer, legume webspinner and lucerne leafroller larvae. Host range: Soy beans and emu-foot (Psoralia tenax). Life cycle: Young larvae mine inside the leaves (Fig. 7.86) before emerging after 3–4 days to feed inside folded leaves. Larvae pupate within the folded leaves. Risk period: Crops are at greatest risk during podfill when they are least tolerant of defoliation. Drought-stressed plants or the most stressed sections of a crop are at greatest risk.
plague infestations (thousands per square metre) can totally defoliate a crop (Fig. 7.88). If this happens, larvae may graze on the surface of pods, but pods are not their preferred food. Monitoring: Look for the early warning signs of rare plague events, including very large numbers of soybean moths around lights at night, and numerous leaf blisters on leaves in the upper canopy. Soybean moth larvae normally only fold and distort a few leaves per square metre of crop. Action level: Indicative threshold is based on tolerable defoliation; that is, 33–40% pre flowering and 15–20% during early podfill. Chemical control: No specific registrations. Helicoverpa pesticides with some degree of persistence are most likely effective against soybean moth, but control is very rarely warranted. Cultural control: Irrigated crops are at lesser risk.
Damage: Young larvae form transparent mines or blisters, before emerging to web and distort leaves. Damage is usually cosmetic due to the small amount of leaf consumed; however, rare
Conservation of natural enemies: As for Helicoverpa and loopers. Small Ichneumonid wasps, Temclucha sp. (Fig. 7.89), have been recorded as parasitising soybean moth larvae.
Fig. 7.87. Adult soybean moth (6 mm long). (DPI&F Qld:
Fig. 7.89. Temclucha Gauld sp. 6, a small Ichneumonid parasitoid (8 mm) of soybean moth. (J. Wessels)
J. Wessels)
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Distribution: Widespread throughout Australia including Tas. Also reported from New Zealand. Pest status: Minor, widespread, irregular.
Fig. 7.90. A Lecithocera sp. larva on a peanut pod (10 mm). This species is very common at the base of peanut bushes. While they attack pods in the laboratory, they are thought to feed on dead leaves in the field. (DPI&F Qld: J. Wessels)
L E C I T H O C E R I DA E
Lecithocera sp. Lepidoptera: Lecithoceridae Distribution: Probably a native species. Very common in the south Burnett region, Qld. Pest status: Minor. Identification: The slender larvae reach 10 mm in length and are grey with hair tufts (rather than single hairs) (Fig. 7.90). The adults are tiny brassy grey moths (5 mm long) with long pale antennae. May be confused with: The larvae may be confused with those of Endotricha puncticostalis (Pyralidae) but have hair tufts instead of single hairs. They are considerably smaller and more slender than those of Hydrilloides lentalis. Host range: Peanuts, and possibly other summer pulses. Damage: Larvae make webs and feed on dead leaves (trash) at the base of peanut bushes. In the laboratory, larvae will feed on peanut pods but this type of feeding has not been observed in the field. Monitoring: Look for the larvae in trash at the base of peanut bushes. Action level and chemical control: Not required.
LE AFROLLER MOTH S ( TORTRICIDAE )
Identification: The moths are small, being only 8–10 mm long, and rest with the wings folded over their body. The leading edge (costa) of the forewing is strongly convex near the base. Female moths have uniformly cream wings, while males moths are cream with dark bands (Fig. 15.66) Young larvae are a pale with a dark head. Older larvae are yellow green with pale green stripes, a pale brown head and sparse short pale hairs. May be confused with: Larvae may be confused with beet webworm, legume webspinner and soybean moth larvae. Host range: While lucerne is this pest’s primary host, it has also been reported from soy beans, clover and sweet peas. Non-legume hosts include sunflower, geranium, garden mint, lettuce, carrots and dandelions. Life cycle: The larvae roll and web leaves together and feed within the resultant sheltered site. Larvae pupate within the folded leaves. Risk period: Crops are usually at greatest risk during early podding. Damage: Larvae web and roll leaves. Damage is usually only cosmetic with little leaf being eaten (because the larvae are so small). Monitoring: As for the soybean moth. Action level: None available. Chemical control: No registered products. Cultural control and conservation of natural enemies: As for other caterpillars. Small Ichneumonid wasps, Temclucha sp., parasitise the lucerne leafroller.
PLU M E M OT H S ( P T E RO PH O R I DA E )
Lucerne leafroller
Geranium plume moths
Merophyas divulsana (Walker) Lepidoptera: Tortricoidae
S. anisodactylus (Walker) and S. zanclistes (Meyrick) Lepidoptera: Pterophoridae 223
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Fig. 7.91. Plume moth larva, Sphenarches sp. (8 mm).
Fig. 7.92. Adult plume moth Sphenarches sp. (12 mm).
(J. Wessels)
(J. Wessels)
Previously referred to as Oxyptilus spp.
Risk period: Flowering and podding.
Distribution: Sphenarches anisodactylus is a cosmopolitan species reported in West Africa, Madagascar, India, Thailand, Japan, New Hebrides, tropical Americas, and Australia. In Australia, plume moths occur in the tropics and subtropics.
Damage: Larvae damage buds, flowers and pods. The damage does not significantly affect production.
Pest status: Minor, widespread and irregular. In India, the plume moth (Exelastis atomosa) is a major pest of pigeon peas. Identification: The larvae, pupae and moths are distinctive. Caterpillars reach 10 mm in length. They are variable in colour from green to orange and are covered in distinctively arranged long white spines or bristles (Fig. 7. 91). Some species may have a pattern down their back. The pupae are also covered in spines and can be mistaken for larvae. The moths are small (10 mm wingspan) with very narrow wings with long plume-like scales. The wings are buff with brown marks on some species. The moth’s legs are very long with large spines or spurs. Moths often rest with their abdomens curved up into the air, and their wings held at right angles to the body with the plumes folded (Fig. 7.92). May be confused with: Plume moths are unlikely to be confused with other Lepidoptera. Host range: Summer legumes attacked include cowpeas, dolichos, mung beans, navy beans, pigeon peas and soy beans. The geranium plume moth also attacks geraniums. Life cycle: Larvae feed by tunnelling into the flower buds and pods then pupate on the plant, usually at the base of flower or pod stalks.
Cultural control: Aim for uniform and timely plant emergence to assist in the synchrony of flowering, thus reducing the time a crop is at risk from flower-attacking pests.
P Y R A L I D M OT H S ( P Y R A L I DA E )
Bean podborer Maruca vitrata (Fabricius) Lepidoptera: Pyralidae Previously known as Maruca testulalis. Also known in some countries as the mung moth. Distribution: A cosmopolitan pest found in the Americas, Africa, Southern Europe, India, Asia and Australia. It is found in all parts of Australia but is far more abundant in tropical and subtropical coastal regions. Pest status: Major, widespread and regular in regions favouring this pest. A major pest of Vigna legumes (adzuki beans, mung beans and cowpeas), pigeon peas, and navy beans. Identification: The eggs are pale cream and flattened. Larvae are pale cream with two rows of distinctive paired black markings on their back (Fig. 7.93). In the final instar, these markings are often very pale. Larvae can reach 18 mm in length. Bean podborer moths have a 20–25 mm wingspan and a slender body characteristic of pyralids. They have brown forewings with a
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Host range: Favoured hosts include adzuki beans, mung beans, cowpeas, pigeon peas and navy beans. All of the above are semideterminate to indeterminate, with flowers and pods both present during at least during the earlier stages of reproductive development. Bean podborer is also reported from soy beans and peanuts, but is rarely damaging in these crops as flowering does not overlap with podding in soy beans, and peanuts have underground pods. Favoured weed hosts include Sesbania.
Fig. 7.93. Medium bean podborer larva (7 mm), showing characteristic paired black markings, in webbed adzuki flowers. (J. Wessels)
white band extending two-thirds down the wing from the leading edge (Fig. 7.94). Inside this band near the leading edge is a white spot. The hindwings are predominantly a translucent white with an irregular brown border. When at rest, they adopt a characteristic pose with outspread wings and the front of the body raised (Fig. 7.94). There are two colour forms in Australia, the second form having a slightly less extensive brown band on the hindwings and may represent a second species. May be confused with: Moths may be confused with those of the beet webworm (Spoladea recurvalis), but are slightly larger, and have predominantly pale hindwings, as opposed to the predominantly brown hindwings of S. recurvalis.
Fig. 7.94. Bean podborer moth, Maruca vitrata (Fabricius), in typical pose with body raised at front and wings outstretched (25 mm). (DPI&F Qld: J. Wessels)
Life cycle: Crops may be infested from early budding onwards. The eggs are laid on or in the flowers (inserted between the petals). Young larvae feed inside flowers for 5–7 days before moving to the pods when mid-sized. Favoured entry points are where flowers and pods are touching. After completing their development (10–15 days from egg hatch), larvae exit pods and pupate in the soil. Occasionally, M. vitrata larvae indulge in vegetative feeding and have been observed tunnelling in the stems and feeding inside the rolled and webbed leaves of luxuriantly growing (moisture-stress-free) peanuts. Risk period: From early budding onwards until the end of flowering. Indeterminate hosts are very susceptible, with both flowers and pods often present for at least a month. Damage: Seeds within damaged pods are totally or partially eaten out by bean podborer larvae. Entry holes also let in water, which stains the remaining non-eaten seeds. In peanuts, bean podborer larvae may web young leaves together, and may tunnel in plant stems. Such damage is not typical of its behaviour on other legumes and has only been witnessed by the author on rare occasions. Monitoring: The early sign of activity is webbing of flowers on reproductive racemes. Infested pods have a well-defined entry hole (usually one per larva) frequently ringed with moist larval frass. Look for these signs and for the distinctive moths, which may be flushed by a long-handled sweep net. Open all flowers from as many racemes as possible to look for larvae (at least 10 racemes randomly sampled across a crop). Divide the total number of bean podborer detected by the number of racemes sampled, and multiply by the estimated number of racemes per square metre. 225
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Action level: The indicative threshold is three larvae per square metre, but accurate assessments are difficult where larvae are in flowers. Chemical control: Pesticides are most effective if applied before larvae enter pods. Less than perfect control is often achieved as larvae can feed entirely inside flowers for 4–5 days before attacking pods. They consequently encounter only degraded spray deposits. Cultural control: Eradicate the weed host Sesbania spp. from cropping areas. Conservation of natural enemies: The avoidance of broad-spectrum pesticides prior to flowering may help conserve natural enemies; however, beneficial insects are unlikely to control populations above about 10 per square metre.
Beet webworm Spoladea recurvalis (Fabricius) Lepidoptera: Pyralidae Previously known as Hymenia recurvalis. One synonym is Hymenia exodias. Distribution: The Beet webworm is a cosmopolitan pest of tropical and subtropical regions, but in more recent times has been reported from more temperate countries including Belgium and Denmark. It is found throughout Australia but is most common in the tropics and subtropics. Pest status: Usually a minor leaf-feeding pest in Australian pulses but a significant pest in beet plantations. Identification: Beet webworm moths are dark brown with a white band across each forewing and hindwing, and an incomplete band across each forewing (Fig. 7.95). They have a wingspan of about 20 mm. Beet webworm larvae are pale green with a distinctive dark green dorsal stripe flanked by pale white stripes, sparse white hairs and a pale brown head. They have four pairs of ventral prolegs. Larvae can reach 20 mm in length and wriggle violently when disturbed. May be confused with: Beet webworm larvae may be confused with small loopers and Anticarsia irrorata; however, loopers have only
Fig. 7.95. Beet webworm moth (20 mm wingspan). (DPI&F Qld: J. Wessels)
two pairs of ventral prolegs, and A. irrorata larvae have pale transverse bands as opposed to the pale longitudinal stripes of the beet webworm. Beet webworm moths are sometimes confused with bean podborer moths (Maruca vitrata); however, the latter is easily differentiated because its hindwings are mostly pale with only a brown outer band, whereas beet webworm moths have mostly brown hindwings with a transverse white band. Host range: Pulse crops attacked include adzuki beans, mung beans, navy beans and soy beans. Other crops attacked include silverbeet and beetroot (Beta vulgaris), Trianthema postulacastrum, cockscomb (Celosia spp.), goosefoot (Chenopodium spp.), Portulaca spp., and Amaranthus spp. Life cycle: The life cycle is relatively short, with larvae developing in 2–3 weeks and pupation lasting 1 week. Where populations are high, larvae live and pupate communally in a silken web of leaves and frass on their food plants. Risk period: Podfill when crops are least tolerant of defoliation. Damage: Larvae are leaf-feeders, webbing leaves together. Silken webs and frass are indicative of beet webworm attack, but other pyralids can cause similar symptoms. Monitoring: Larvae can be detected when beatsheet sampling. Also inspect webbed leaves. Action level:. Indicative thresholds are based on tolerable defoliation; that is, 33–40% pre flowering and 15–20% during early podfill.
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May be confused with: Their darker colour differentiates larvae from other webspinner and leafroller species. Host range: Includes soy beans, cotton and lucerne.
Fig. 7.96. Cotton webspinner larva (15 mm long). (DPI&F Qld: J. Wessels)
Chemical control: No specific registrations. Any pesticides effective against Helicoverpa (except Helicoverpa virus) will most likely control beet webworm. Cultural control and conservation of natural enemies: As for Helicoverpa and loopers.
Cotton webspinner Achyra affinitalis (Lederer) Lepidoptera: Pyralidae Previously known as Loxostege affinitalis. Distribution: Throughout Australia. Pest status: Minor, widespread, irregular. Identification: Larvae are slender and are grey– green with a dark head (Fig. 7.96). They are sparsely haired and have a dark lateral stripe above each pale lateral band. They also have a dark stripe down the middle of their back flanked on each side by three rows of small dark spots. Larvae can reach 15 mm in length and wriggle violently when disturbed. Moths are pale yellow to cream with dark brown or reddish flecks with a 20 mm wingspan. At rest their wings are swept backwards forming a narrow triangle (Fig. 7.97).
Fig. 7.97. Adult cotton webspinner (20 mm wingspan). (DPI&F Qld: J. Wessels)
Life cycle: Moths may undertake long-distance migrations and the pest can suddenly appear in very large numbers. Risk period: Seedling stage, when sudden and devastatingly high populations can shred leaves. Damage: Larvae skeletonise leaves and web them together. Sudden and devastatingly high populations can ‘shred’ the leaves of seedling crops leading to plant death. Monitoring: Look for early infestation signs such as terminal damage, webbing and the windowing of leaves. Action level: The indicative threshold is based on tolerable defoliation; that is, 33–40% and greater than 25% terminal loss. Chemical control, cultural control and conservation of natural enemies: As for the legume webspinner. If spraying seedling crops, restrict the spray to a band over the plants, so as to minimise the impact on beneficials such as ants and spiders in the inter-row.
Endotricha puncticostalis Walker Lepidoptera: Pyralidae Distribution: Throughout tropical and subtropical Australia. Larvae are commonly found at the base of peanut bushes in southern and central Qld. Pest status: Minor, widespread, regular. Identification: The distinctive moths are small (wingspan 12–15 mm) and are brownish-purple with a wide pale cream band across each wing, and a narrower white band at this band’s inner edge. The moths often rest with outspread wings and arched body, looking like miniature deltawinged aircraft. The slender larvae are pinkishgrey to black in colour and reach 12 mm in length. Larval webbing is frequently found at the base of peanut bushes. The small pale cream eggs are similar to etiella eggs. May be confused with: Larvae may be confused with those of Lecithocera sp. (Lecithoceridae), 227
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pod damage depends on seasonal soil conditions.
Fig. 7.98. Pale Endotricha puncticostalis Walker larva (12 mm) and webbing on dead plant material at the base of a peanut bush. (J. Wessels)
which are very common under peanut bushes. Lecithocera sp. larvae are similar in size and colour but are grey with a pink suffusion. In addition, Lecithocera sp. larvae have hairs in multiple tufts, whereas E. puncticostalis larvae have single hairs. Webbing at the base of plants made by E. puncticostalis larvae is indistinguishable with that made by Etiella behrii. Endotricha puncticostalis moths are similar to those of E. mesenterialis, but are smaller and not as brightly coloured, having less purple on their wings. Host range: Peanuts and probably other summer pulses. Life cycle and damage: Larvae bore into young pods but are unable to penetrate maturing pods, merely scarifying the pod surface. Larval webbing is frequently seen under peanut bushes. Note that the similar looking Lecithocera sp. larvae most likely feed on dead peanut leaves at the base of plants. The species probably does not cause economic damage.
Identification: Eggs are small (0.6 mm diameter) and cream and are flattened (Fig. 7.99). Small larvae may be cream or pale green and lacking in stripes, and with a dark head. Mid-sized larvae may be pale green or cream, with pale brown or reddish stripes. Larger larvae are characteristically green with pink or reddish stripes and a brown head (Fig. 8.11). Larvae in the pre-pupal stage can be aqua blue or dark pink with no stripes. The moths are small (12 mm long at rest, 20–22 mm wingspan) and distinctively coloured. They are grey–brown in colour with a distinctive costal stripe along the leading edge of each forewing, and a transverse orange band on each forewing (about onequarter of the distance along each forewing from its base). Hindwings are pale grey in colour. The wings are folded back along the body when resting. Moths have a prominent ‘snout’ (formed by the labial palps), that is typical of pyralids (Fig. 8.11). May be confused with: Etiella behrii moths may be confused with those of E. zinckinella, which is very similar in colour (but which does not attack peanuts). Moths may also be confused with those of other non-pest Etiella spp., including the slightly larger E. scitivittalis, which has pinker forewings, and E. chrysoporella, which is a more golden brown. Because of their variable colouration, larvae are easily confused with
Risk period: During peanut podfill. Monitoring: As for E. behrii. Look for webbing and larvae at the base of plants.
Etiella (or lucerne seed web moth) Etiella behrii (Zeller) Lepidoptera: Pyralidae N (Main entry in Chapter 15 Pastures—winter rainfall.)
Distribution: Etiella behrii is found over much of South-East Asia: China, Indonesia, the Pacific Islands, and Australia including Tas. Pest status: Major, widespread, irregular. While frequently present in peanut crops, the level of
Fig. 7.99. Etiella behrii eggs (0.6 mm) on rattlepod. (DPI&F Qld: J. Wessels)
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those of other Etiella spp. and other small pyralid larvae including those of an undescribed genus similar to Faveria oculiferella (Meyrick), which has been recorded from the South Burnett region in south-east Qld. Larvae grow to 15 mm, are a dull green with dark stripes and a dark head capsule. The larvae are similar in appearance to Etiella behrii, but have black pigmentation behind the head, as opposed to red for etiella. The small moths are dark grey–brown with a pale orange head. Larvae feed secretively within webbed leaves of peanut plants. Webbing made by etiella larvae at the base of peanuts may be confused with that made by the (darker) larvae of Endotricha puncticostalis (Pyralidae) and Lecithocera sp. (Lecithoceridae). Host range: Summer pulses include peanuts, mung beans, adzuki beans, lima beans and soy beans. Other hosts include lentils, lupins, faba beans, field peas, seed lucerne, as well as native legumes such as Acacia spp.(wattles), native peas, Rhynchosia, and rattle pods (Crotalaria spp.). Rattle pods are also favoured hosts of other Etiella species. Life cycle: Eggs are laid on pods or under bracts and are very hard to detect. In legumes with aboveground pods, newly hatched etiella larvae bore straight into pods leaving a near-invisible entry hole. In summer pulses, larvae complete their development in a single pod before escaping through a characteristic pinhole-sized exit (2–3 mm diameter). In peanuts, many newly hatched larvae move down the plant and seek out the pods, which are below ground. The majority of pod-seeking larvae reach the pods within 24 hours of egg hatch. E. behrii is the only species in the genus with this behaviour. Pod damage in peanuts is most likely in drought years as soil moisture inhibits larval movement. Larval movement is assisted by cracks in dry soils. Larvae may also feed on flowers and in peanuts, and sometimes also feed on vegetative terminals, under bracts, and inside stems; however, laboratory and glasshouse trials suggest etiella larvae are unable to complete their development in peanut terminals. In peanut pods, the life cycle can be completed in 4 weeks at 30°C. Risk period: Crops may be infested from flowering onwards, but are at greatest risk during late podding. Peanuts are at particular
Fig. 7.100. Etiella damage to mung bean seeds showing typical pale frass, which is often mistaken for bruchid eggs. (J. Wessels)
risk during end-of-season droughts when the dry soil allows larval access to the pods. Such conditions also favour the development of aflatoxin in pods with etiella exit holes. Damage: Seeds are only partially eaten, often with characteristic pin-hole damage. This damage is very difficult to grade out and its unattractive appearance reduces seed quality. Larval frass adhering to damaged mung bean seeds is frequently mistaken for bruchid eggs (Fig. 7.100); however, unlike bruchids, etiella are unable to infest seed in storage. In peanuts, etiella damage is a major aflatoxin risk factor. This highly carcinogenic toxin is produced by an Aspergillus fungus, which gains entry to pods through holes (2–3 mm diameter) made by exiting etiella larvae (Fig. 7.101), but not through the very much smaller entry holes (0.2–0.3 mm diameter). Etiella-damaged pods can have aflatoxin levels 100 times greater than undamaged pods. Etiella infestations initiated close to harvest can result in the post-harvest emergence of larvae as peanut pods dry down in storage. Even a low-field infestation (< 1% of pods infested) can give rise to 10 000 or more
Fig. 7.101. Etiella damaged pod with gross fungal contamination. (DPI&F Qld: J. Wessels)
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Fig. 7.104. Agathis sp. (3.5 mm), a common braconid parasitoid of Etiella behrii. (H. Brier)
Fig. 7.102. Etiella cocoons in dust under a peanut-drying bin. Tens of thousands of larvae can emerge from a single tonne of harvested peanuts; however, etiella cannot re-infest peanuts in storage. (DPI&F Qld: J. Wessels)
larvae per tonne of harvested peanuts (Fig. 7.102). Monitoring: Etiella moths are readily monitored with light traps. Trap catches are a reliable indicator of likely pod damage. Inspect crops for both larvae and moths. In peanuts, inspect damaged terminals for etiella larvae. Such damage is most common at flowering (i.e. mid season) but may not be a reliable indicator of end-of-season etiella larval activity in pods. Etiella larvae in mature pods can only be detected by shelling pods. Webbing at the base of peanut plants and around small pods may be caused by etiella, but can also be caused by other less important and non-pest species.
Fig. 7.103. Larvae of the braconid parasite, Agathis sp., emerging from their etiella host. Note the parasite’s aqua blue colour matching the ventral colour of the host. (DPI&F Qld: J. Wessels)
Action level: There are no current thresholds for etiella in summer pulses. Future thresholds will be based on moth activity to minimise egg lay and subsequent larval damage. Chemical control: No products are registered against etiella in summer pulses. In practice, once larvae are in pods, pesticide control is not possible. Cultural control: Cultural control for etiella is most applicable to peanuts. Where water is available, late irrigations will keep soil moist and will lessen the risk of both etiella and associated aflatoxin damage. If larvae are detected in pods, an early harvest may reduce the aflatoxin risk, by removing peanuts from the field before larvae exit pods. Post-harvest management: Because etiella damage tolerance levels are extremely low in some summer pulses (e.g. a 1%), infrared or equivalent sorters are necessary to remove damaged seed post harvest. Conservation of natural enemies: Etiella larvae in pulses and weed hosts with aboveground pods (e.g. rattlepods) are often parasitised by Agathis sp. wasps (Braconidae) (Figs 7.103 and 7.104). However, no parasitism has ever been observed in etiella larvae collected from peanut pods, which is further evidence that most larvae infest peanut pods soon after egg hatch.
Legume webspinner or bean leafroller Omiodes diemenalis (Guenée) Lepidoptera: Pyralidae
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Fig. 7.105. Webspinner larval-grazing produces angular feeding areas, webbing and frass. (J. Hughes)
Previously referred to in Australia as Lamprosema abstitalis (Walker). Note that species in the genus Omiodes are sometimes referred to in the literature as Nacoleia sp. Distribution: A tropical species reported from India, South-East Asia, Micronesia, Pacific Islands, and northern Australia. Pest status: Minor, widespread in coastal regions, irregular. Identification: The larvae roll and web leaves together, but are considerably larger than larvae of the soybean moth or lucerne leafroller. Young larvae are pale green with dark heads. Older larvae are shiny green with pale brown heads and reach 14 mm in length (Fig. 7.105). The distinctive moths are brown with bright yellow patches and have an 18 mm wingspan (Fig. 7.106). May be confused with: Larvae may be confused with beet webworm larvae and (when small) soybean moth larvae. Host range: Soy beans, mung beans and navy beans Life cycle: The larvae roll and web leaves together and feed within the resultant sheltered site. Risk period: Crops are usually at greatest risk during early podding. Damage: Larvae are leaf-feeders, webbing leaves together. Silken webs and frass are indicative of webspinner presence (Fig. 7.105), but other pyralids produce similar signs.
Fig. 7.106. Adult legume webspinner moth (Omiodes diemenalis (Guenee)) (18 mm wingspan). (DPI&F Qld: J. Wessels)
Monitoring: Larvae will be sometimes detected when beat-sheet sampling. Also inspect webbed leaves. Action level: Indicative threshold is based on tolerable defoliation; that is, i.e. 33–40% pre flowering and 15–20% during early podfill. Chemical control: No specific registrations. Pesticides effective against Helicoverpa (except Helicoverpa virus) will most likely give control of legume webspinner. Cultural control and conservation of natural enemies: As for Helicoverpa and loopers.
Sorghum head caterpillar Cryptoblabes adoceta Turner Lepidoptera: Pyralidae Also called false blossom moth. Distribution: An exotic pest reported from NSW, NT and Qld. Also occurs in New Zealand. Pest status: Minor, widespread, irregular. Identification: Young larvae are yellow but darken with age to yellow–brown to grey, usually with dark and orange stripes (Fig. 7.107). Larvae reach 13 mm in length. Moths have grey forewings with a transverse two-tone pale/dark band. Moths rest with their wings folded back along the body and are only 8 mm long. 231
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Fig. 7.107. Sorghum head caterpillar Cryptoblabes adoceta Turner (10 mm) in adzuki pod. (J. Wessels)
Fig. 7.108. Brown colour form of Ectropis despicata looper (30 mm) on soy beans, Kingaroy, Qld. Note the single pair of ventral prolegs, characteristic of geometrid loopers. (J. Wessels)
Distribution: Probably an endemic species. May be confused with: Distinctive colouration differentiates larvae from other podborers. The striped larvae resemble small armyworms, but unlike armyworms they taper noticeably towards the head and rear. Host range: Sorghum but also recorded on adzuki beans and pigeon peas in south-east Qld. Life cycle: Eggs are probably laid on or near pods. Larvae bore into and feed inside pods. On pigeon peas, webbed leaves provide sheltered feeding sites. Risk period: Podding. Damage: Holes made in adzuki pods may let in water and increase seed weathering. Leaf webbing is unlikely to affect yield. Monitoring: Look for webbing and larvae in pods. Moths are difficult to detect.
Pest status: Minor, irregular. Identification: The larvae are large but slender geometrid loopers with one pair of ventral prolegs and reaching over 30 mm in length. Larvae have faint longitudinal strips, and vary in colour from pale green to pale brown (Fig. 7.108). About one-third of the way down of their body they have a pair of dark spots. The moths are pale grey with cryptic darker markings and a 30 mm wingspan (Fig. 7.109). May be confused with: Larvae may be confused with noctuid loopers but the single pair of ventral prolegs and their slender body are distinctive features. Host range: Larvae have been collected in soy beans in the South Burnett. This species is quite likely a polyphagous feeder as are other members of this genus.
Action level: No specified thresholds. Chemical control: No registered products. Pesticides targeting Helicoverpa are likely to be effective. Cultural control and conservation of natural enemies: As for bean podborer. GEOME TRID LOOPER S ( GEOME TRIDAE )
Large twig looper Ectropis despicata Walker Lepidoptera: Geometridae
Fig. 7.109. Ectropis despicata moth (30 mm wingspan) on soy beans, Kingaroy, Qld. (J. Wessels)
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Damage: Larvae feed on leaves. Life cycle, risk period, monitoring, control and natural enemies: As for other minor leaf feeding caterpillar pests.
Tiger looper Gymnoscelis lophopus Turner Lepidoptera: Geometridae Distribution: Coastal northern Australia from Cairns to northern NSW. Pest status: Minor, widespread, regular. Identification: The moths are small, nondescript geometrids (15–18 mm wingspan) with light brown wings with darker markings. Larvae are small (10 mm long) and very variable in colour (pale green to yellow to orange). Larvae sometimes have dark marks (stripes) on each segment (Fig. 7.110), hence the informal ‘tiger looper’ epithet. May be confused with: Other Gymnoscelis species. Host range: Mung beans and soy beans (and most likely other summer legumes). Life cycle: Larvae mostly found on flowering racemes. Risk period: Flowering and early podding. Damage: Larvae attack buds, flowers and small pods, and are reported as causing significant flower damage to experimental mung bean plots in the South Burnett region of south-east Qld.
Fig. 7.110. Gymnoscelis lophopus larva (10 mm) with distinct striping. (J. Wessels)
One synonym is Acidalia recessata. Distribution: Most summer pulse growing regions in eastern Australia. Pest status: Minor, widespread, irregular. Identification: Larvae are long and thin with only one pair of ventral prolegs. When resting, larvae often ‘stand’ erect mimicking a twig (Fig. 7.111). Larvae are fawn with small lateral dots each side near the front of the body. Early instars are striped, but the stripes are absent in later instars which reach 12–15 mm in length. Moths are almost white to cream, with a small black spot on each wing, and have fine brown to grey bands across their wings. The cup-shaped eggs are laid on stalks. Moths have a wingspan of 16 mm and rest with their wings outspread. May be confused with: Twig caterpillars may be confused with young bean loopers (Mocis sp.), which are also very thin; however, the latter species has two pairs of ventral prolegs.
Monitoring: Check for larvae in or on developing buds when checking for pests such as podborer and small Helicoverpa. Cultural control: Aim for uniform and timely plant emergence to assist in the synchrony of flowering, thus reducing the time a crop is at risk from flower-attacking pests. Conservation of natural enemies: As for other caterpillars.
Twig caterpillar or looper Scopula perlata (Walker) Lepidoptera: Geometridae
Fig. 7.111. Twig caterpillars looking like twigs. Larvae are up to 12 mm long, with a single pair of ventral prolegs. (J. Wessels)
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Host range: Soy beans and peanuts. Nonlegume hosts include the weed forget-me-not (Myosotis arvensis). Life cycle: Larvae feed on leaves and pupate in leaf litter at the base of plants. Risk period: Crops are not threatened at any stage due this species’ virtual non-pest status. Damage: This species has never been reported as causing damage of any significance. Monitoring: Can be sampled with a beat sheet. Action level: Not applicable. Control is not costeffective. Biocide: Bt is registered for twig looper control in all the main summer pulses. Conservation of natural enemies: As for other lepidopteran species.
B LU E S ( LYC A E N I DA E )
Bean flower caterpillar or dark cerulean Jamides phaseli (Mathew) Lepidoptera: Lycaenidae One synonym is Lampides oranigra. Distribution: Northern Australia, from Derby, WA, to Port Macquarie, NSW. Pest status: Minor, restricted, irregular. Identification: The butterflies have a 25–28 mm wingspan. Male wings are deep blue on top with a black margin. Hindwings have a series of spots along the rear margin, including a prominent black one at the tornus. Females are similar but are paler with a wider black margin, and are slightly larger than the males. Both sexes have a small tail on each hindwing with a single dark eye ringed with orange at their underside base. The undersides of the wings are pale brown with transverse pale dashes. Small larvae are green but larger larvae are brown with dark stripes. Larvae reach a length of 10 mm. Eggs are flattened and white. May be confused with: The butterflies are very similar to the pea blue but lack the distinctive
white band across the outer sector of the underside of the wings. The larvae are similarly shaped but are different in colour to those of the pea blue. Small larvae may be confused with those of the grass blue. Host range: Most members of the Fabaceae (legumes), including navy beans and mung beans. Life cycle: Probably similar to the pea blue. Larvae are often attended by ants. Risk period, damage, monitoring, action level, chemical control, cultural control and conservation of natural enemies: As for the pea blue.
Common grass blue (butterfly) Zizina labradus (Godart) Lepidoptera: Lycaenidae Also known as grass blue and lucerne blue. A very similar South-East Asian species, Z. otis, shares the same common name. Distribution: Native to and spread throughout Australia including Tas. Pest status: Minor, widespread; in eastern Australia, Zizina labradus is most damaging to soy beans in inland areas, irregular. Identification: The adult’s wings are pale dull blue on top with dark grey edgings, and lack tails and eye spots. The edging is broader in the females. The undersides of the wings are brown with soft markings. The eggs are relatively large (1 mm diameter) and are laid singly, and are bluish and flattened with a central depression. The small green slug-like larvae reach only 10 mm in length. Larvae are pale green with pale stripes on their back. The head is difficult to see and is usually tucked out of sight (Fig. 7.112). May be confused with: The larvae are sometimes confused with hoverfly (Diptera: Syrphidae) larvae, both of which may be found on leaf terminals. Hoverfly larvae are more tapered towards the head and often wave the front of their body from side to side. Other lycaenid larvae are also similar in outline and colour, particularly those of the bean flower caterpillar Jamides phaseli. The adults can be distinguisghed from other lycaenids such as the
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Islands. The race ‘platissa’ occurs in northern Australia. Pest status: Minor, widespread, irregular.
Fig. 7.112. Large larva of grass blue butterfly (10 mm long) and windowing of its feeding area. Note the larva’s pale stripes and patterning and that its head is hidden. (J. Wessels)
pea blue, Lampides boeticus, by their lack of eyespots and tails. Host range: May feed on most pulse legumes, but they are most common in soy beans. Life cycle: Eggs are laid singly on leaves. Larvae mostly feed on leaves. Larvae pupate in loose webbing under leaves. Risk period: Zizina labradus can attack at any stage. Less vigorous drought-stressed plants are at greatest risk as terminals are more likely to be attacked. Damage: They mostly feed on leaves and terminals, but occasionally feed on flowers. Damaged leaves may be windowed. Excessive terminal loss results in pods being set too close to the ground, which reduces crop harvestability. Monitoring: Check crops with a beat sheet, and also look for larvae inside terminals.
Identification: Pale pea blues have a 25 mm wingspan. Males are pale blue on top with a black spot and tail at the tornus (outer angle) of each hindwing. Females’ wings are blue on top with wide brown edging, and have an orangeringed black spot and tail on each hindwing. Underneath, they are fawn with irregular dark arcs edged in white. The hindwings each have one large and one small black spot edged in orange by the tornus. Underneath, both sexes are fawn with white markings. There is a small black eye spot, ringed with orange, at the base of each tail, on the underside of the wings. The larvae reach 10 mm in length, are yellowish green, with a reddish line edged in white along the back. Pupae are mottled brown and 8 mm long. May be confused with: The butterflies may be confused with those of the closely related lycaenids Jamides phaseli and Lampides boeticus, but can be distinguished by the twin spots underneath the wings, or the lack of a pale band on the underside of their wings, respectively. All these species are readily distinguished from the common grass blue by the presence of tails on their hindwings. Pale pea blue larvae are more prominently marked than those of the pea blue. Host range: Many members of the Fabaceae (legumes), including pigeon peas and rattle pods. Life cycle, risk period, damage, monitoring, action level, chemical control, cultural control
Action level: Control if terminal loss exceeds 25%. Chemical control: Most products targeting Helicoverpa (except Helicoverpa virus) will also control this pest.
Pale pea blue (butterfly) or forget-me-not Catochrysops panormus platissa (HerrichSchafer) Lepidoptera: Lycaenidae Distribution: This species occurs as various races from India across Asia to the Solomon
Fig. 7.113. Adult pea blue butterfly (30 mm wingspan). Note the prominent transverse white bands, eyespots and short tails. (DPI&F Qld: J. Wessels)
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and conservation of natural enemies: As for Lampides boeticus.
Pea blue (butterfly) Lampides boeticus (Linnaeus) Lepidoptera: Lycaenidae Distribution: Asia, Africa, India, southern Europe, Hawaii and Australia. It occurs throughout mainland Australia. Pest status: Minor, widespread, irregular. Identification: Pea blue butterflies have a wingspan of 30 mm. The males’ wings are pale blue on top, whereas females are blue with wide dark brown edges. Underneath, both sexes are pale brown with white markings and have a distinctive broad white band across the outer sector of both wings. Both sexes have a small thin tail on each hindwing. There are two small black eye spots, ringed with orange, at the base of each tail, on the underside of the wings (Fig. 7.113). Larvae reach 10 mm in length. Larvae are variable in colour ranging from a uniform pale cream to pale pink with darker pink markings. The pale brown head is usually retracted out of site when larvae are disturbed (Fig. 7.114). Younger larvae may be pale pink with darker pink markings. The eggs are flattened, white and only 0.2 mm across. Pupae are mottled brown with a length of 8 mm. May be confused with: The butterflies may be confused with those of the closely related lycaenids Jamides phaseli and Catochrysops panormus, but can be distinguished by the prominent pale band on the underside of their wings. All these species are readily
distinguished from the common grass blue by the presence of tails on their hindwings. Pea blue larvae are paler and less prominently marked than those of the other lycaenids mentioned in this chapter. Host range: Most members of the Fabaceae (legumes), including soy beans, mung beans, adzuki beans, navy beans, Dolichos and pigeon peas. Other recorded hosts include siratro, lupins, broad bean, garden pea and sweet pea and Sturt’s desert pea. Life cycle: Eggs are laid singly on buds and flowers. Larvae feed and pupate within the buds and flowers and possibly small pods. Pupae drop to the ground when flowers die and shrivel and drop off the plant. Risk period: Budding and flowering. Damage: The larvae eat out buds and flowers. Small pods may also be damaged. Monitoring: Look for larvae inside buds and flowers and for damage. The pale larvae may be difficult to see in pale coloured buds. Look also for the distinctive butterflies. Cultural control: Irrigation of crops at flowering will improve plant vigour and allow the crop to replace damaged flowers. Conservation of natural enemies: As for other Lepidopteran pests.
T U S S O C K M OT H S ( LYM A N T R I DA E )
Tussock moths Lepidoptera: Lymantridae Species attacking pulses include: Eubrostis sp. Painted pine moth, Orgyia australis Walker Distribution: Most summer pulse-growing regions. Pest status: Minor, widespread, irregular.
Fig. 7.114. Larva of pea blue (10 mm long), with head extended. (H. Brier)
Identification: Moths are stoutly bodied. Eubrostis sp. moths are uniformly pale cream to yellow in colour with a 30 mm wingspan. In contrast, Orgyia australis moths are brown with cryptic darker markings.
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Fig. 7.116. Crotalaria moth caterpillar (20 mm). (DPI&F Qld: Fig. 7.115. Tussock moth larva on soy beans, Kingaroy, Qld (20 mm long). (H. Brier)
Larvae are reasonably large (up to 30 mm long) and are covered in long hairs, some of which are arranged in clumps or ‘tussocks’ (Fig. 7.115). Larvae are often brightly coloured, a warning to potential predators that the hairs are extremely irritating. May be confused with: The distinctive arrangement of the hairs distinguishes tussock moth larvae from other caterpillars. Host range: Reported in soy beans and peanuts but may attack other summer pulses. Life cycle on pulses: As for other leaf-feeding Lepidoptera.
J. Wessels)
Distribution: A cosmopolitan tropical species, found throughout northern Australia. Pest status: Minor, widespread, regular. Identification: Larvae reach 25 mm in length and are grey with pale yellow stripes and orange spots, and are covered with stiff sparse hairs (Fig. 7.116). The moth is white with black and red patches on its wings (Fig. 7.117) and appears similar to Utetheisa pulchelloides (Fig. 7.118). Moths have a 30–35 mm wingspan. May be confused with: The heliotrope moth has a red spot on the tornus of each forewing, absent on the wings of the crotalaria moth.
Risk period: Podding, when plants are least tolerant of defoliation.
Host range: Reported in Australia from soy beans, pigeon peas and rattle pods (Crotalaria spp.).
Damage: Larvae feed on leaves.
Risk period: Flowering and podding.
Monitoring: Use a beat sheet.
Damage: Larvae attack leaves but cause no significant damage.
Action level: Use the same percentage defoliation thresholds as used for loopers and other defoliators. Chemical control: No specific products are registered but they should be controlled with products targeting Helicoverpa (except for nuclear polyhedrosis virus (NPV)-based products).
T I GER M OT H S ( A RC T I I DA E )
Crotalaria moth Lepidoptera: Arctiidae Utetheisa lotrix (Cramer) and heliotrope moth Utetheisa pulchelloides Hampson
Fig. 7.117. Crotalaria moth (30–35 mm wingspan). (J. Wessels)
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Fig. 7.118. Heliotrope moth, Utethesia pulchelloides Hampson (30–35 mm wingspan). (DPI&F Qld: J. Wessels)
Monitoring: Utetheisa lotrix larvae can be sampled with a beat sheet.
H E R M I N I I DA E
Fig. 7.120. Nodaria externalis moth (25 mm wingspan). (J. Wessels)
Nodaria externalis Guenée Lepidoptera: Herminiidae
been collected from soy beans in the South Burnett region of south-east Qld.
Distribution: Asia and Australia.
Pest status: Minor, widespread, irregular.
Pest status: Minor, irregular.
Identification: The larvae are dark with a small head, and have faint orange longitudinal stripes and bands between the body segments (Fig. 7.121). The moths’ forewings are fawn with a transverse pale band and curved pale line along the margin of each hindwing (Fig. 7.112).
Identification: Larvae are up to 15 mm long and are cream with a reticulated pattern of brown and grey stripes and spots (Fig. 7.119). The moths have a 25–28 mm wingspan and dark grey forewings with faint transverse markings (Fig. 7.120). The hindwings are pale grey. Their antennae may have thickened middle segments. Host range: Soy beans and peanuts. Damage: Larvae feed on leaves. Monitoring and action level: As for other leaffeeders.
Simplicia caeneusalis (Walker) Lepidoptera: Herminiidae
May be confused with: The moths’ colouration resembles that of the sugarcane looper, (Mocis frugalaris); however, the latter is larger (35 versus 28 mm wingspans) and has a straight rather than a curved band across each hindwing. Host range: Soy beans in the south Burnett region, Qld.
Distribution: This species ranges from India to Australia as far south as Sydney. Larvae have
Damage: Larvae in this genus are reported as feeding on dead leaves; however, Simplicia caeneusalis larvae will feed readily on green soybean leaves.
Fig. 7.119. Nodaria externalis larva (13 mm long) on soy beans, Kingaroy, Qld. (H. Brier)
Fig. 7.121. Simplicia caeneusalis, final-instar larva (25 mm). (H. Brier)
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Fig. 7.124. Adult Anticarsia irrorata (40 mm wingspan). (J. Wessels)
Fig. 7.122. Simplicia caeneusalis moth (28 mm wingspan). (J. Wessels)
A R M Y WO R M S , C U T WO R M S , N O C T U I D S , S E M I - LO O PE R S ( N O C T U I DA E )
Anticarsia irrorata (Fabricius) Lepidoptera: Noctuidae Previously known as Saroba irrorata. Distribution: India, South-East Asia, and Australia. In Australia, Anticarsia irrorata has been recorded from the Ord in WA, the NT and Qld. Pest status: Minor, widespread, irregular. The closely related velvetbean caterpillar, Anticarsia gemmatalis (Hübner), is a major legume pest in the Americas, particularly of soy beans. Identification: Larvae have four pairs of ventral prolegs. Young larvae are bright green with pale cream bands between the segments. When disturbed, they wriggle violently. Large larvae can reach 40 mm in length and are green with a pale pink stripe along each side (Fig. 7.123). Moths have a 35–40 mm wingspan and slightly
Fig. 7.123. Large Anticarsia irrorata larva (30 mm long). The lateral line on large larvae is frequently much pinker than on this specimen. (J. Wessels)
‘hooked’ forewings. Both forewings and hindwings are brown. The forewings have a distinctive B-shaped spot and an oblique line running from their apex down to and across the hindwings (Fig. 7.124). Eggs are spherical, white and ribbed. May be confused with: Young larvae may be confused with soybean and Chrysodeixis spp. loopers, but have four ventral prolegs, as opposed to two for loopers, and have transverse banding rather than the longitudinal stripes of loopers and webworms. The pink lateral stripes distinguish large Anticarsia larvae from Helicoverpa and loopers. Host range: In Australia, Anticarsia irrorata has been recorded from mung beans, soy beans and Dolichos. Other reported hosts (not in Australia) include kodo millet (Paspalum scrobiculatum) and rice (Oryza sativa). Life cycle: Eggs are laid singly on leaves and take 3–6 days to hatch. Larvae pass through six stages and take 3–4 weeks to complete their development. Fully developed larvae leave plants to pupate in the soil. Risk period: Crops are at greatest risk during flowering and podding but can be attacked at any stage. In southern regions, the greatest risk is during mid to late summer. In the tropics, activity may continue during winter (dry season) months. Damage: While being mainly foliage feeders, larvae may also attack flowers and pods (based on the feeding behaviour of A. gemmatalis). For this reason, adzuki beans, navy beans and mung beans are most likely at greater risk than soy beans, as they have pods that are more easily 239
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damaged by species that are not specialised podfeeders.
and sugarcane are the armyworms most commonly encountered in summer pulses.
Monitoring and action level: Refer to Helicoverpa and Mocis alterna, respectively. Tolerable defoliations in vegetative and flowering crops are 33% and 20%, respectively. In adzuki beans, navy beans and mung beans (which are more susceptible to flower and pod attack), the threshold during flowering and podding is provisionally three larvae per square metre.
Identification: See Chapter 2. Sugarcane armyworm moths have pale forewings with a dark line running the length of the forewing.
Chemical control: Anticarsia irrorata is most likely susceptible to most chemical insecticides used against Helicoverpa. The closely related A. gemmatalis is reported as being susceptible to Bacillus thuringiensis (Bt) but not Helicoverpa virus. Cultural control: Refer to soybean loopers. Trap crops of early plantings of soy beans are reported to be effective against moths of the closely related velvetbean caterpillar (A. gemmatalis) in the Americas. Conservation of natural enemies: The same principles apply as for Helicoverpa and loopers. While there is no Australian data specific to A. irrorata, tachinid flies are reported to be effective parasites of the closely related velvetbean caterpillar, A. gemmatalis. Several fungal pathogens have been associated with velvetbean caterpillar, the most important being the fungi Nomurea rileyi and Entomophthora sp., as well as a naturally occurring nuclear polyhedrosis virus. (This is not the virus that infects Helicoverpa.)
Armyworms
Risk conditions: Coastal soy beans are at greatest risk during the seedling stage. In the subtropics, spring-planted crops are at greater risk from common armyworm. Armyworm outbreaks sometimes occur after the end of a drought. Pulses are at risk after armyworms have eaten out their more favoured food. Soybean seedlings in coastal regions are sometimes damaged by sugarcane armyworms where crops are planted into cane trash blankets. Damage: Seedlings may be lopped and killed. Armyworm plagues have the potential to shred larger crops. Monitoring: Look for larvae in trash at the base of the plant during the day, particularly if there are suspicious seedling deaths. Action level: Take action if more than 20% seedling death is found. Chemical control: Band sprays are effective against armyworms in seedling crops. Armyworms are susceptible to most Helicoverpa pesticides. Cultural control: Avoid planting into trash blankets in paddocks with a history of armyworms. Natural enemies: Armyworms are often paraitised by Apantales and Coetesia sp. wasps. Fungal and viral (NPV) epizootics often decimate high-density armyworm populations.
Lepidoptera: Noctuidae Common armyworm, Mythimna convecta (Walker)
Cluster caterpillar, Spodoptera litura (Fabricius) Lepidoptera: Noctuidae
Northern armyworm, M. seperata (Walker)
Distribution: Asia, India, New Guinea, New Zealand, some Pacific islands and Australia. In Australia, Spodoptera litura is more common in tropical and coastal regions, but extends as far south as Perth and central NSW.
Sugarcane armyworm, Leucania stenographa Lower N (Main entry in Chapter 2 Cereals.)
Distribution: All summer pulse-growing regions. Pest status in summer pulses: Minor, widespread, irregular. The common, northern
Pest status: Moderate, widespread, irregular in present growing areas but this may change with the expansion of pulses into tropical Australia. Spodoptera litura has been recorded damaging peanuts in the NT and coastal soy beans in Qld.
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a
Fig. 7.126. Newly hatched cluster caterpillars (2 mm long) characteristically remain near the egg mass from which they hatch. (H. Brier) b
streaks (Fig. 7.125b) and translucent white hindwings edged with brown. May be confused with: Small to medium larvae (10 mm long) may be confused with Helicoverpa larvae, but may be distinguished by the ‘hump’ behind the head, and the row of dark spots along each side (Fig. 7.127).
Fig. 7.125. (a) Spodoptera litura egg mass with the furry appearance typical for this species, and (b) moth. (DPI&F Qld: J. Wessels)
Identification: Eggs are laid in a furry cream mass on the underside of leaves (Fig. 7.125a). Young larvae ‘cluster’ together and are translucent green with a darker thorax (Fig. 7.126). Middle-sized larvae are variable in colour and are smooth-skinned with a pattern of red, yellow, and green lines, a dark patch on the hump behind the head (mesothorax), and dark spots along each side (Fig. 7.127). Large larvae are initially brown with three thin pale yellow lines down the back: one in the middle and one on each side. They have a row of black dots along each side, and a row of conspicuous dark half-moons along the back (Fig. 7.128). Final instar larvae are dark and can exceed 50 mm in length. All larvae have four pairs of ventral prolegs, and are more solidly built than Helicoverpa larvae (Fig. 7.128). Moths are larger than Helicoverpa with a 40 mm wingspan. Moths have brown forewings with criss-cross cream
Fig. 7.127. Medium Spodoptera litura (left) showing the small ‘hump’ behind the head compared with Helicoverpa armigera (right) larvae (both 10 mm). Arrowed is the ‘saddle’ on the second and third abdominal segments of H. armigera, often but not invariably present on this species but not H. punctigera. (DPI&F Qld: J. Wessels)
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Distribution: Athetis is a widely distributed genus ranging from Africa to New Zealand. Athetis tenuis has been reported from Australia and New Zealand.
Fig. 7.128. Large Spodoptera litura larva (30 mm) showing the pattern of dark ‘half-moons’ along its back. (DPI&F Qld: D. Ironside)
Host range: Polyphagous; pulse hosts include adzuki beans, mung beans, navy beans, peanuts, pigeon peas and soy beans. Life cycle: Egg masses are laid on leaves. Young larvae feed on leaves but older larvae may feed on flowers and pods. Larvae pass through six larval stages and take 2–3 weeks to develop depending on temperature. Larvae pupate in the soil. Risk period: Flowering and podding. Damage: Small larvae window leaves, but older larvae chew holes in leaves. Older larvae also attack flowers and pods but are not as damaging as Helicoverpa. Larvae attack the pegs of peanuts and in light soils they may attack pods. Monitoring: As for Helicoverpa. Look also for the distinctive egg masses and clusters of young larvae. Action level: In pre-flowering crops, control is warranted if defoliation exceeds (or is likely to) exceed 33%. However, tolerable defoliation drops to 15–20% once flowering and podding commences. For adzuki beans, mung beans, navy beans and peanuts, an indicative threshold is three larvae per square metre. Chemical control and cultural control: Cluster caterpillars are controlled by most pesticides targeting Helicoverpa; however, cluster caterpillars are not controlled by NPV and are difficult to control with Bt unless very small. Thorough coverage is required in peanuts to reach cluster caterpillars attacking pegs and exposed pods at the base of the plants. Conservation of natural enemies: As for Helicoverpa and loopers.
Athetis tenuis (Butler) Lepidoptera: Noctuidae Synonyms include Proxenus tenuis and Radinogoes tenuis.
Pest status: Minor, widespread, irregular. Significant populations have occasionally been observed in peanut and soybean crops in the South and Central Burnett regions in Qld. Identification: The moths have brassy brown forewings with faint markings and a 25 mm wingspan. Larvae are stout-bodied and are translucent pale cream with a subtle darker criss-cross pattern. Host range: Recorded in soy beans and peanuts. Other hosts include barley. Damage: Larvae in the genus are reported as feeding on dead leaves. Cultured Athetis tenuis larvae feed readily on the green leaves of host plants and have been successfully reared through to adulthood (moths).
Cutworms, brown and black Lepidoptera: Noctuidae Agrotis munda (Walker) and A. ipsilon (Hufnagel) N (Main entry in Chapter 2 Cereals.)
Distribution: All summer pulse-growing regions. Pest status in pulses: Minor, widespread, irregular. May be confused with: Larvae may be confused with armyworms but are not as prominently marked. They may also be confused with larvae of other smaller noctuids commonly found at the base of plants, e.g. Hydrilloides lentalis. Risk period: Seedling stage, particularly if planted into weedy cultivation already hosting cutworms; however, Agrotis munda has been occasionally reported in large numbers in flowering and podding peanuts. Damage: Larvae can sever seedling stems and reduce plant stands. Later infestations in peanuts may also damage the pegs. Monitoring: Look for larvae in trash at the base of the plant during the day, particularly if there are suspicious seedling deaths or peg damage in peanuts.
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Action level: More than 20% seedling death. Chemical control: Band sprays are effective against cutworms in seedling crops. Cutworms are susceptible to most Helicoverpa pesticides. Cultural control: Avoid planting into weedy cultivation harbouring cutworms. Conservation of natural enemies: Cutworms are parasitised by a number of Ichneumonid wasps and Tachanid flies.
Ericeia inangulata Guenée Lepidoptera: Noctuidae Distribution: Most of Qld as far west as Cunnamulla and into northern NSW. Pest status: Minor, widespread, irregular. Identification: The moths have a 40–45 mm wingspan. All wings are fawn grey with a darker outer transverse band and numerous faint darker markings. The large thick-bodied larvae are grey and have a pale lateral line. Larvae reach 40 mm in length and have four pairs of ventral prolegs. May be confused with: The larvae are similar in size and colouration to cutworms. Host range: Larvae have been collected in soy beans in the south Burnett region, Qld. Life cycle: As for other noctuids. Risk period, damage, monitoring, action level, chemical control and conservation of natural enemies: As for other leaf-feeders.
Fig. 7.129. Eublemma dimidialis larva attacking mung beans, Darwin, NT. (M. Shepard)
fawn body (Eublemma) versus pink stripes on a green body (Etiella). Host range: Recorded from mung beans in the NT. Risk period: Flowering and podding. Damage: Larvae bore into pods and feed on the seeds (Fig. 7.129). Monitoring: Use a beat sheet. Action level: Provisionally three larvae per square metre.
H E L I C OV E R PA S PP.
Corn earworm and native budworm Lepidoptera: Noctuidae Helicoverpa armigera Hübner and Helicoverpa. punctigera Wallengren
Eublemma dimidialis (Fabricius) Lepidoptera: Noctuidae Distribution: South-East Asia, and the northern half of Australia. Pest status: Minor, restricted, irregular. Identification: The larvae are fawn with brown stripes and pale sparse hairs (Fig. 7.129). The moth is distinctively coloured with a 20 mm wingspan (Fig. 7.130). The forewings are pale cream proximally with a broad bright pink outer band. The hindwings are fawn. May be confused with: The pattern of larval striping is similar to that of Etiella behrii, but the colours are different, being brown stripes on a
Fig. 7.130. Eublemma dimidialis (Fabricius), Darwin, NT (20 mm wingspan). (M. Shepard)
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N (Main entry in Chapter 3 Cotton.)
Pest status: Both species are major, widespread and regular pests of summer pulses. Identification: See Chapter 3, Figs 3.18–3.23. May be confused with: Helicoverpa armigera larvae can be distinguished from soybean loopers by having four pairs of ventral prolegs instead of two pairs, and do not taper noticeably towards the head. Armyworm larvae can be distinguished by having fewer hairs and by bodies that taper slightly at both ends. Medium-sized Helicoverpa armigera larvae may also be confused with the cluster caterpillar (Spodoptera litura), but are hairier and lack the latter’s distinctive spots and hump behind the head (Fig. 7.127). Helicoverpa eggs are paler than looper eggs, which have a greenish tinge and which are squatter. Helicoverpa armigera pupae are readily separated from H. punctigera pupae, which have spines that are close together (Fig. 3.22). Damage to vegetative terminals and flowers of peanuts may be confused with that caused by Etiella larvae. Host range: Of the summer pulses, soy beans are the crop most attractive to Helicoverpa during the vegetative stage. Pigeon peas are a particularly attractive host from flowering onwards, as they have a protracted flowering period. Pigeon peas may be used as a Helicoverpa trap crop. Risk period: H. armigera and H. punctigera are the most damaging of summer pulse pests as they can severely damage all crop stages and all plant parts (except peanut pods which are underground). Crops are most attractive to moths during budding, flowering and early podding. Vegetative soy beans are more attractive than the vegetative stages of other summer pulses, and can even be damaged during the seedling stage. In southern Qld and further south, summer pulses are at greatest risk from H. armigera in the latter part of summer (January onwards); however, spring outbreaks are not uncommon. In the tropics, H. armigera is often active throughout the winter (or dry season) and ‘winter planted’ summer legumes are thus at risk. Spring outbreaks of H. punctigera are more likely after winter rains in inland Australia. Damage: Helicoverpa armigera larvae can attack all aboveground plant parts and can attack summer pulses at any stage from seedlings to podfill. In general, summer legumes can
Fig. 7.131. Foliage-feeding by Helicoverpa on flowering soy beans is characterised by rounded edges to the holes. (H. Brier)
tolerate considerable defoliation: up to 33% with no yield loss. Helicoverpa spp. defoliation is characterised by rounded chew marks and holes, as opposed to angular holes made by loopers (Fig. 7.131). High Helicoverpa populations in seedling or drought-stressed crops can cause considerable damage if vegetative terminals and stems are eaten, resulting in pods being set closer to the ground and making them difficult to harvest. On drought-stressed crops, the last soft green tissue is usually the vegetative terminals, which are thus more likely to be totally consumed than in normally growing crops. High Helicoverpa populations at seedling stage may totally consume leaves and then move to terminals and stems. Larvae select buds, flowers and pods when these become available except in soy beans, where young larvae prefer vegetative terminals, young leaves and flowers before attacking pods. Larvae target the seeds in large pods but destroy pods in the process (Fig. 7.132). Crops may compensate for early pod damage by setting additional pods, but where water is limiting significant early damage may delay or stagger podding, with subsequent yield and quality losses. Damage to welldeveloped pods frequently results in weather damage of uneaten seeds. Helicoverpa do not damage peanut pods that are belowground, but can attack the pegs that develop from fertilised flowers aboveground, and from which the pods develop after the pegs penetrate the soil. Monitoring: Inspect crops weekly during the vegetative stage and twice weekly from very early budding onwards until late podding.
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northern NSW soybean cultivars range from 1.2–2.0 larvae per square metre, depending on crop value and pesticide cost, and a suggested threshold of one larva per square metre for mung beans and navy beans. For peanuts, estimates for flowering threshold range between three to six per square metre.
Fig. 7.132. Pod-feeding by fifth-instar Helicoverpa armigera (20 mm) on adzuki beans. (DPI&F Qld: J. Wessels)
Increased scouting is recommended as crops approach budding, as there have been instances of young buds being totally consumed by Helicoverpa before flowering commenced. Beatsheet sampling is the preferred sampling method for medium to large Helicoverpa larvae. Small larvae should be also scouted for by inspecting (opening) vegetative terminals and flowers. Damage to vegetative terminals is often the first visual clue that Helicoverpa larvae are present. Summer legumes should be scouted for Helicoverpa eggs (Fig. 7.133) and moths, to pinpoint the start of infestations and to increase the chance of success of biopesticides such as Helicoverpa virus. Sample six widely spaced locations per field, and take five one-metre long samples at each site with a ‘standard’ beat sheet. Convert larval counts per metre to larvae per square metre by dividing counts by the row spacing in metres. Action level: Before flowering, summer pulses can tolerate 33% defoliation without yield loss. Control is warranted if loss of vegetative terminals (tipping) exceeds 25%. From flowering onwards, Helicoverpa thresholds are cropspecific. Thresholds for determinate Qld and
Fig. 7.133. Helicoverpa armigera eggs on pigeon pea buds. Note also the grass blue butterfly egg on the left (blue arrow). (H. Brier)
Chemical control: Biopesticides (particularly NPV) are recommended in the vegetative stage of summer pulses. While these may give less than perfect control (e.g. only 70%) they conserve beneficials which buffer against Helicoverpa attack during later stages. In addition, perfect control of Helicoverpa is not necessary during a legume crop’s vegetative stage. To manage resistance of H. armigera, control post flowering should target small larvae (a 7 mm), and the number of spray applications per crop is usually restricted to one. For best results, all ingestion type products (including biopesticides) require thorough plant coverage by using multiple nozzles, droppers, high pressure and high spray volumes (at least 30 and 100 L ha–1 for aerial and ground applications, respectively). For biopesticides, the addition of a feeding supplement is recommended. Cultural control: Where possible, avoid successive plantings of summer legumes. Maintenance of large, vigorously growing plants reduces the risk of terminal damage as the terminals of water-stressed plants are attractive to larvae. Vigorously growing plants with adequate available moisture are also better able to replace damaged leaves and compensate for flower and pod damage. Survival of overwintering pupae in summer pulses is very low (often < 10%) due to predation and high levels of parasitism. In regions where diapause occurs, pupal survival may be further reduced by cultivating soil to a depth of 10 cm before moth emergence in August. Conservation of natural enemies in summer pulses: Predatory bugs attacking Helicoverpa eggs and larvae include: spined predatory bug (Oechalia schellenbergii (Guérin-Méneville)) (Fig. 8.20), glossy shield bug (Cermatulus nasalis (Westwood)) (Figs 3.38–3.39), Pacific damsel bug (Nabis kingbergii Reuter), bigeyed bugs (Geocoris lubra Kirkaldy (Fig. 3.35) and less frequently Germalus sp.), brown smudge bug (Deraeocoris signatus (Distant)) (Fig. 3.36), apple dimpling bug 245
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(Campylomma liebknechti (Girault)), and assassin bugs (Pristencanus spp.) (Fig. 3.34). Predatory beetles include red and blue beetle (Dicranolaius bellulus), predatory ladybirds and (less frequently) soldier beetles (Chauliognathus spp., Fig. 7.70). Other important predators include ants, spiders and lacewings. Helicoverpa parasitoids include the tiny wasps Trichogramma spp. (Fig. 3.50) and Telenomus sp. (egg parasitoid), the wasp Microplitis demolitor (Fig. 3.52) and Lissopimpla sp. (Fig. 8.19) (small to medium larvae), Heteropelma and Netelia (Fig. 8.18) (medium to large larvae) and Ichneumon sp. (Fig. 8.19) (pupae), as well as numerous species of tachinid flies (Fig. 8.17) (larvae). With the exception of the egg parasitoids and Microplitis, the parasitoids above do not kill until the pupal stage. Mice, predatory earwigs and wireworm larvae are significant predators of Helicoverpa pupae. In addition to beneficial arthropods, naturally occurring caterpillar diseases frequently have a marked impact on Helicoverpa in summer legumes. Epizootics of Helicoverpa NPV occur frequently in crops with high Helicoverpa populations. Peanuts, being close to the ground and soil splash, provide good transmission conditions.
Hydrilloides lentalis Guenée Lepidoptera: Noctuidae Distribution: India, Asia and Australia south to Sydney. Common in peanut crops in the tropics and subtropics. Pest status: Minor, widespread, with significant populations in many peanut crops.
Fig. 7.134. Hydrilloides lentalis larva (20 mm long) on peanuts, Kingaroy, Qld. (J. Wessels)
Endotricha puncticostalis (Pyralidae) and Lecithocera sp. (Lecithoceridae). Host range: Peanuts. Life cycle: Larvae feed at the base of plants and hence often go unnoticed. Damage: Hydrilloides lentalis has been observed feeding on green (living) peanut leaves in the South Burnett region, Qld. Monitoring: Look for larvae in leaf trash at the base of plants.
N O C T U I D LO O PE R S
Castor oil looper Achaea janata (Linnaeus) Lepidoptera: Noctuidae Also known as the croton looper. One synonym is Phalaena melicarta. Distribution: India, South-East Asia, Taiwan, Papua New Guinea, New Zealand, most Pacific
Identification: Larvae can reach 20 mm in length and are relatively stout and have a wrinkled ‘rough-skinned’ appearance. They are dark with sparse pale hairs, and have two pale brown–orange bands along their back, which may be faintly patterned with further pale and dark markings (Fig. 7.134). The moths have a 22– 25 mm wingspan and have dark grey inner and outer forewings with a wide mustard-coloured central transverse band. Each forewing has a small dark ‘quater moon’ shape in the middle of this band (Fig. 7.135). May be confused with: Small larvae may be confused with those of the much smaller
Fig. 7.135. Hydrilloides lentalis moth (25 mm wingspan). (J. Wessels)
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Monitoring, action level, chemical control, cultural control and conservation of natural enemies: Refer to soybean loopers.
Eight spots moth Amyna axis (Guenée) Lepidoptera: Noctuidae Also known as Amyna octo (Guenée). Fig. 7.136. Castor oil looper (45 mm long), on a soybean leaf, Kingaroy, Qld. (DPI&F Qld: J. Wessels)
islands including Hawaii, Chile and Australia. In Australia, it is common in tropical and subtropical regions. Pest status: Minor, restricted, irregular. While its large size means A. janata is potentially very damaging, populations in summer legumes are usually too low to be of concern. Identification: The eggs are blue and spherical and about half a millimetre in diameter. They are laid singly on the food plant. Larvae are initially brown with a black and white head, a red knob on the tail, and a transverse black mark on the back of the second abdominal segment (Fig. 7.136). The spiracles on each side of the abdominal segments are black. The underside and hind legs are pale. In the last instar, the brown turns to bluish-grey and the point on its tail turns black. The first pair of prolegs is degenerate and they consequently only have three pairs of functional ventral prolegs. Larvae move with a typical looper action. The moths are large with a wingspan of 50–55 mm. Their forewings have variable patterns of light and dark brown, while the hindwings are black with distinctive white markings. May be confused with: This species is very distinctive and should not be confused with other looper pests in Australian summer legumes.
Distribution: Africa, India, Indonesia, Cook Islands and Australia. In Australia, it most likely extends as far south as northern NSW. Larvae have been recorded (> three per square meter) in mung bean crops in the south Burnett region of south-east Qld and have also been reported from Biloela in central Qld. Pest status: Minor, widespread, irregular. Its incidence may be underestimated as larvae may be misidentified in the field as Chrysoieixis spp. loopers. Identification: The thin-bodied larvae are pale bright green with faint white stripes and have two pairs of ventral prolegs (Fig. 7.137). Larvae reach 23 mm in length and wriggle violently when disturbed. The moths’ forewings are chocolate-brown with numerous faint grey– white lines and a faint small figure eight on each forewing (Fig. 7.138). Males have a small semitransparent patch on each forewing. The hindwings are lighter with very faint markings. The moths have a 23 mm wingspan. May be confused with: The larvae look very much like small Chrysodeixis spp. loopers and Anticarsia irrorata larvae. Their violent wriggling and parallel body differentiates them from Chrysodeixis spp. loopers, which taper noticeably towards the head. Amyna axis larvae have fewer ventral prolegs than Anticarsia (two versus four pairs), which is also a ‘wriggler’.
Host range: Soy beans, Acacia spp., Bauhinia, castor oil plants, cotton, crotons, eucalypts, hoop pine, lychees, macadamias, roses and white mangrove. Life cycle and risk period: As for soybean looper. Damage: Most likely a defoliator of soy beans. Populations are usually too low to be damaging. The adult moth may feed on some fruits by piercing the skin and sucking the juices.
Fig. 7.137. Final-instar larva (22 mm long) of the eight spots moth on mung beans, Byee, Qld. (J. Wessels)
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Fig. 7.138. Eight spots moth (23 mm wingspan), Byee, Qld. (J. Wessels)
Fig. 7.139. Mocis alterna eggs, globular and slightly ribbed. (DPI&F Qld: J. Wessels)
Host range: Mung beans, cowpeas, soy beans and lucerne.
Larvae of this distinctive species can be cream, charcoal, bright orange or brown. Larvae may have dark stripes along their back and a cream or yellow band along each side (Fig. 7.140); however, pale cream larvae may only have very faint stripes. Larvae can reach 40 mm in length, have two pairs of ventral prolegs, move with a looping action, and are more slender than Helicoverpa and soybean loopers. A distinctive feature of this species is the forward-sloping and striped head. Larvae pupate in a cocoon inside curled leaves. Moths have a 30–32 mm wingspan and more rounded wings than Helicoverpa. Moths are grey or grey–brown with dark bands and markings on all wings (Fig. 7.141).
Life cycle pulses: As for other loopers. Risk period: Podfill. Damage: Probably feed mostly on leaves. Monitoring: Use a beat sheet. Action level: Use defoliation thresholds as for loopers. Chemical control: Most likely controlled with pesticides targeting other caterpillars. Cultural control and conservation of natural enemies: As for loopers.
M O C I S S PP. LO O PE R S
Bean looper or Mocis Mocis alterna (Walker) Lepidoptera: Noctuidae Distribution: Native to Australia. Mocis alterna is widespread through northern Australia from the north-west of WA, across the top end of NT, most cropping regions in Qld, and as far south as northern NSW. Within this range, the bean looper is more commonly encountered in coastal and sub-coastal regions.
May be confused with: Mocis alterna moths may be confused with those of the larger Mocis trifasciata (45 mm wingspan). However, they may be distinguished by the greater ‘scalloping’ of the dark markings on the forewings, and by the ‘angling in’ of these markings at their anterior end (when wings are extended in a setting board position). Mocis alterna larvae may be confused with those of the less common (in legumes) sugarcane looper, Mocis frugalaris. Larvae of this
Pest status: Moderate, widespread, regular. The most common Mocis species looper in Australian summer pulses. Identification: Eggs are globular, pale green and slightly larger than Helicoverpa eggs (Fig. 7.139).
Fig. 7.140. Larva of bean looper (26 mm long), typical colour form on mung beans, Kingaroy Qld. (J. Wessels)
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Fig. 7.142. Large Mocis trifassiata larva (45 mm) on soy beans, Bundaberg, Qld. (J. Wessels) Fig. 7.141. A bean looper female (32 mm wingspan). (DPI&F Qld: J. Wessels)
species are brown and have two transverse black bands across their thorax (these are most visible when larvae arch their backs). Host range: Mocis alterna has been recorded from adzuki beans, mung beans, navy beans and soy beans. Other host crops include lucerne. Life cycle and risk period: As for soybean loopers. Damage: Larvae are primarily foliage feeders in soy beans but will attack flowers and developing pods of adzuki beans, mung beans and navy beans. Monitoring, action level, chemical control, and cultural control: Refer to soybean loopers. Conservation of natural enemies: As for soybean loopers, except that there are fewer records of braconid wasp parasites.
Mocis trifasciata (Stephen) Lepidoptera: Noctuidae
darker bands across their thorax (most visible when larvae arch their backs). The larval head capsule is striped as in M. alterna. Larvae can reach 50 mm in length and have two pairs of ventral prolegs. Moths have a 45 mm wingspan and are larger than other Mocis species. Moths are brown with two dark bands across each forewing. Their hindwings also have dark bands (Fig. 7.143). May be confused with: Its larger size and the pattern of its dark markings on the forewings distinguish M. trifasciata moths from M. alterna. Larvae may be confused with those of M. alterna but are distinguished by the transverse dark bands on the thorax. Medium-sized larvae are very similar to medium Pantydia metaspila larvae, but unlike the latter, have prominently striped head capsules. Host range: Reported in cowpeas (Vigna spp.) in north Qld and from soy beans at Bundaberg. Other hosts include rice and Brachiaria. Life cycle and risk period: As for soybean loopers.
Distribution: Indonesia, Papua New Guinea, the south-west Pacific and Australia. In Australia, it is found in the NT and Qld, south to Brisbane. It is more common in coastal regions. Pest status: Minor to moderate, most common and frequent in coastal regions. Identification: Medium larvae are brown and prominently striped, often with a darker area down the middle of the back. Large larvae are pale brown (fawn) or orange with faint stripes and two broad diffuse grey bands along their back (Fig. 7.142). They have two transverse
Fig. 7.143. Female Mocis trifassiata (45 mm wingspan) on soy beans, Bundaberg, Qld. (J. Wessels)
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Damage: Primarily a foliage-feeder on cowpeas and soy beans. There are no records or observations from other summer legumes. Monitoring, action level, chemical control and cultural control: Refer to soybean loopers. Conservation of natural enemies: As for Mocis alterna.
Sugarcane looper Mocis frugalaris (Fabricius) Lepidoptera: Noctuidae Distribution: The species is found across Africa, Asia (including India), Papua New Guinea, many Pacific islands and Australia. In Australia, its range is similar to that of M. alterna. Pest status: Minor, most likely restricted, irregular. Identification: Younger larvae are brown with pale stripes. Large larvae are usually dark (Fig. 7.144) with a transverse black band across their thorax (only visible when larvae arch their backs). Larvae can reach 40 mm in length and have two pairs of ventral prolegs. Moths have a 35 mm wingspan and have slightly more angular wings than Mocis alterna. Moths are brown with a diagonal dark line with a pale inner edge across each forewing. Hindwings are slightly darker at their outer edge (Fig. 7.145). May be confused with: The single dark line across the forewings distinguishes M. frugalaris moths from other Mocis species. Larvae may be confused with those of M. alterna but are distinguished by the transverse black bands on the thorax.
Fig. 7.145. Moth of sugarcane looper (30 mm) dark form on soy beans, Kingaroy, Qld. (H. Brier)
Host range: Occasionally found in soy beans. Preferred hosts are grasses including sugarcane and oats. Life cycle and risk period: As for soybean loopers. Damage: Primarily a foliage-feeder on soy beans. There are no records or observations from other summer legumes. Monitoring, action level, chemical control, and cultural control: Refer to soybean loopers. Conservation of natural enemies: As for Mocis alterna.
Lepidoptera: Noctuidae Pantydia spp. including Pantydia sparsa Guenée, P. metaspila Walker and P. capistrata Lucas One synonym for P. sparsa is Toxocampa orthosiodes. Distribution: Pantydia sparsa is found in the southern half of Australia as far north at least as the Wide Bay Burnett in Qld, as well as islands in the South Pacific. Pantydia metaspila and P. capistrata have been collected from soybean crops at Bundaberg and Kingaroy in Qld, respectively, but are most likely more widely distributed in Australia. Pantydia metaspila has also been recorded from Hong Kong. Pest status: Minor, widespread, irregular.
Fig. 7.144. Larva of sugarcane looper (30 mm) dark form on soy beans, Kingaroy, Qld. (H. Brier)
Identification: Larvae can reach 35–40 mm in length. Medium Pantydia metaspila larvae are brown with darker stripes, but large P. metaspila larvae develop a mottled appearance and have a distinctive pale ‘blaze’ on their head capsule.
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Fig. 7.146. Final-instar Pantydia capistrata Lucas larva (30 mm) on soy beans, Kingaroy, Qld. (H. Brier)
Pantydia capistrata larvae are brown with a darker strip down the middle of the back (Fig. 7.146) and two transverse black bands only visble when the caterpillars arch their backs. Their head capsule is dark with four prominent white stripes (Fig. 7.146). Small to medium Pantydia spp. larvae have two pairs of ventral prolegs, but large larvae may develop three pairs of ventral prolegs. The moths have a wingspan of 33–40 mm, Pantydia metaspila being the largest of these species. Pantydia sparsa and P. capistrata are greyish-fawn with variable darker markings, those on P. sparsa being more extensive. Pantydia metaspila have dark brown forewings with a thin pale band across their outer sector (Fig. 7.147). Hindwings of all species have a broad dark outer band. May be confused with: The moths may be confused with those of Helicoverpa moths but are slightly larger and stouter and greyer. Helicoverpa moths also lack Pantydia’s black ‘face’. Young larvae may be confused with Mocis larvae.
Fig. 7.147. From top to bottom: Pantydia sparsa Guenée, P. capistrata Lucas and P. metaspila Walker. Note the dark flattened C-shapes at the bottom of forewings for P. sparsa and P. metaspila. Respective wingspans of these specimens are 33, 36 and 40 mm. (DPI&F Qld: J. Wessels)
Host range: All three Pantydia species have been recorded from soy beans. Other reported hosts include egg and bacon pea (Dillwynia ericifolia,) and lucerne.
Distribution: Africa, Asia, Papua New Guinea and Australia. First reported in Australia in 1976 in Qld. Now also recorded from NSW and Tas but most likely more widespread in Australia. This pest is more common in coastal regions.
Life cycle: As for Helicoverpa. Risk period: During podfill. Damage: Pantydia has been observed attacking leaves. It is not known whether larvae attack pods. Monitoring: As for Helicoverpa and loopers.
Soybean looper Thysanoplusia orichalcea (Fabricius) Lepidoptera: Noctuidae One synonym is Diachrysia aurifera.
Pest status: Moderate, widespread, regular. Identification: Eggs are pale yellow–green and ribbed and are flatter than Helicoverpa eggs. Larvae move with a distinctive looping action and have only two pairs of ventral prolegs. Their body tapers noticeably towards the head. Larval colour can vary considerably. Very small larvae are green all over but medium larvae usually have prominent dark and white striping. Large larvae are usually green with white stripes (Fig. 7.148). Larvae have pale hairs, which are most prominent on darker and newly moulted specimens. Larvae can reach 45 mm in length. 251
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Host range: Soy beans, mung beans, navy beans and most likely adzuki beans. Rarely seen on peanuts. Also recorded from lucerne, sunflowers, potatoes and parsley. Life cycle: Looper eggs hatch in 3–6 days. There are six larval stages. Larvae take 2–3 weeks to develop before pupating under the leaves in a loose silken cocoon.
Fig. 7.148. Large soybean looper (Fabricius) (30 mm) with green colouration, numerous narrow white bands and a thicker white lateral band. (DPI&F Qld: J. Wessels)
Unlike Helicoverpa, which pupate in the soil, soybean looper larvae usually pupate under the leaves in a thin silken cocoon. Pupae are dark above and pale underneath. The moth’s forewings are very distinctive, being brown with a large bright golden patch (Fig. 7.149). The hindwings are fawn-coloured, darkening towards the outer margins. It has a wingspan of about 4 cm. May be confused with: Larvae are unlikely to be confused with Helicoverpa larvae, having only two pairs of ventral prolegs instead of four, a tapering body, and a looping action when moving; however, medium-sized darker specimens look very similar to Helicoverpa larvae. Soybean looper larvae may be confused with vegetable and tobacco loopers (Chrysodeixis spp.), but are more prominently striped. Mistaking soybean loopers for these closely related species is of no consequence as they cause similar damage and all are easily controlled.
Fig. 7.149. Adult soybean looper at rest. (DPI&F Qld: J. Wessels)
Risk period: Crops can be attacked at any stage but are at greatest risk during flowering and podding. Summer legumes are least tolerant of defoliation at these stages, and in some crops (adzuki beans, mung beans and navy beans) flowers and young pods are at direct risk of looper attack. In subtropical regions, crops are at greatest risk from mid to late summer; however, in the tropics, winter-planted ‘summer legumes’ may also be at risk. Damage: Damage depends on host crop. Thysanoplusia orichalcea is a more serious pest in mung beans and navy beans than in soy beans. Small larvae feed on only one side of the leaf, leaving translucent ‘feeding windows’. As larvae develop, they chew holes in the leaf, and then feed from the leaf margin. Larvae are primarily foliage feeders in soy beans but will attack the flowers and developing pods in adzuki beans, mung beans and navy beans. Looper leaf damage is different to Helicoverpa damage, the feeding holes being angular rather than rounded (Fig. 7.150). Monitoring: Inspect crops weekly during the vegetative stage and twice weekly from very early budding onwards until crops are no longer
Fig. 7.150. Soybean crop shredded by a soybean looper population exceeding 50 larvae per square metre. The inter-vein feeding and angular holes differentiates looper damage from that caused by Helicoverpa larvae, which chew through the leaf veins. (DPI&F Qld: J. Wessels)
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susceptible to attack (late podding). Increased scouting is recommended as crops approach flowering, as loopers may attack flowers in some crops, and all crops become less tolerant of defoliation at this stage. Beat-sheet sampling is the preferred sampling method for looper larvae. Small larvae should be also monitored for early signs of damage. Crops should also be scouted for looper eggs and moths, so as to pinpoint the start of infestations and to increase the chance of success of biopesticides such as Bacillus thuringiensis (Bt). Sample six widely spaced locations per field, and take five 1-m long samples at each site with a ‘standard’ beat sheet. Convert larval counts to larvae per square metre by dividing counts per metre by the row spacing in metres. Action level: In pre-flowering crops, looper control is warranted if defoliation exceeds (or is likely to exceed) 33%; however, tolerable defoliation drops to 15–20% once flowering and podding commences. In adzuki beans, mung beans and navy beans, where flowers and pods are attacked, the threshold is set at three looper larvae per square metre. Chemical control: Soybean loopers can be controlled with most pesticides targeting Helicoverpa, but are not controlled by products containing NPV. Small loopers (< 12 mm long) can also be controlled with Bt. For best results, ensure thorough coverage and add a feeding stimulant such as amino feed. In pre-flowering crops, and especially in areas at risk from whitefly attack, use the softest option possible (preferably a biopesticide). Where there is sustained and heavy egg-laying pressure, more than one biopesticide application will be necessary or else a product with greater residual activity, and/or activity against looper moths, may be required, especially where flowers and pods are being attacked.
Fig. 7.151. Litomastix wasp pupae visible through the skin of a parasitised and mummified soybean looper. (DPI&F Qld: J. Wessels)
numerous predators and parasites. Many of these also attack Helicoverpa (e.g. predatory bugs, tachinid flies, braconid wasps and ichneumonid wasps). Loopers are frequently parasitised by Litomastix sp. (Encyrtidae) (Fig. 7.151), with scores of parasite larva developing per looper host (Fig. 7.152). The use of Bt for looper control will help preserve beneficial insects and also reduce the risk of subsequent whitefly and mite attack. Epizootics of looper NPV are frequently observed in crops with high populations; however, larvae are usually not killed by virus until they are medium to large (instars 4–5). Looper virus is not the same as Helicoverpa virus.
Cultural control: Any agronomic practices promoting plant growth increase a crop’s tolerance to looper attack. On larger plants, there will be less defoliation in percentage terms, for a given looper population. Vigorously growing plants will be better able to compensate for flower and pod damage, and damaged leaves will be replaced more quickly. Conservation of natural enemies: Soybean loopers in summer pulses are attacked by
Fig. 7.152. Litomastix sp. wasps from single soybean looper (Trichoplusia orichalcea). (DPI&F Qld: J. Wessels)
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C H RYS O D E I X I S S PP. LO O PE R S , L E PI D O P T E R A : N O C T U I DA E
Tobacco looper Chrysodeixis argentifera (Guenée)
Vegetable looper Chrysodeixis eriosoma (Doubleday) Distribution: Chrysodeixis argentifera (synonym Plusia secundaria) is endemic to Australia and is occasionally reported in New Zealand and Norfolk Island. C. argentifera is common throughout mainland Australia and Tasmania. Chrysodeixis eriosoma occurs throughout Asia and the Pacific to Hawaii and Easter Island and in Australia. Chrysodeixis eriosoma occurs throughout northern and eastern Australia as far south as central NSW. Pest status: Moderate, widespread, regular. The status of these pests depends on their host crop as this influences the type of damage. Chrysodeixis loopers damage flowers and pods of adzuki beans, mung beans and navy beans and the leaves of soy beans. Identification: Eggs are ribbed and pale yellow– green, and are flatter than Helicoverpa eggs. Larvae move with a distinctive looping action and have only two pairs of ventral prolegs. Their body tapers noticeably towards the head. Small larvae are pale green but larger larvae are green with white striping (Fig. 7.153). Tobacco looper larvae have a darker green line along the middle of their back, flanked by two white stripes. Their sides have parallel white lines, and sometimes a row of black spots. Vegetable looper larvae have faint white stripes. Larvae can reach 40 mm in length. Unlike Helicoverpa, which pupate in the soil, larvae of these loopers usually pupate under the leaves in a thin silken cocoon (Fig. 7.154). Pupae are dark above and pale underneath. The moths of both species are very similar in size (30 mm wingspan) and have dark brown forewings with small silver ‘figure eight’ markings. On tobacco looper moths these markings are fused (Fig. 7.155), whereas on vegetable looper they are separated (Fig. 7.156). Tobacco looper also has a small silver S-shaped mark above the figure eight on each forewing.
Fig. 7.153. Larva of the tobacco looper (30 mm). Note the 2 pairs of ventral prolegs, as opposed to 4 pairs for Helicoverpa. (DPI&F Qld: J. Wessels)
This mark is absent on vegetable looper. Male vegetable loopers also have long orange hair-like scales on either side of the abdomen, which are characteristic for this species. May be confused with: Larvae are unlikely to be confused with Helicoverpa larvae, having only two pairs of ventral prolegs instead of four, a tapering body, and a looping action when moving (Fig. 7.153). Chrysodeixis loopers may be confused soybean loopers but are not as prominently striped. They may also be confused with the similar but much smaller green looper of the eight spots moth Amyna axis Guenée (Noctuidae), sometimes present in summer pulses. This species reaches only 25 mm in length and wriggles violently when disturbed. The moth of this species is very different to Chrysodeixis spp. moths. Host range: Both Chrysodeixis species have been recorded from soy beans, mung beans and navy beans. They most likely attack adzuki beans but are rarely seen on peanuts. C. argentifera has also been recorded from clover, canola, sunflowers, silverbeet, tomatoes and tobacco. C. eriosoma also attacks tomatoes, as well as citrus, dahlias, egg plants, geraniums, lettuce, parsley, potatoes, and cobblers pegs.
Fig. 7.154. Vegetable looper pupa in webbing in a curled soybean leaf. (J. Wessels)
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Fig. 7.157. Triginoides hyppasia larva (33mm) on soy beans, Kingaroy, Qld. This species is easily confused with Mocis spp. loopers. (J. Wessels)
Fig. 7.155. Tobacco looper moth (40 mm wingspan). Note the small silver ‘S’ below the large silver marking. (DPI&F Qld: J. Wessels)
and are very distinctive with fawn forewings, each with a large black triangle (Fig. 7.158). Each triangle is bisected by a thick pale line, and bounded on two sides by thinner white lines.
Life cycle, risk period, damage, monitoring, action level, chemical control, cultural control and conservation of natural enemies: As for soybean loopers.
May be confused with: The larvae are very similar in colouration, size and shape to Mocis alterna larvae.
Trigonoides hyppasia Cramer Lepidoptera: Noctuidae
Life cycle: As for other leaf-feeding noctuids.
Formerly known as Chalciope hyppasia. Distribution: Ethiopia, Asia, Pacific Islands and Australia. Widely distributed in northern and eastern Australia as far south as south coastal NSW. Pest status: Minor, widespread, irregular. Identification: Larvae are cream with numerous faint longitudinal lateral stripes, and a darker series of stripes along their back (Fig. 7.157). Their head capsule is also striped. Larvae reach 35 mm in length and have two pairs of ventral prolegs. The moths have a 35–38 mm wingspan
Fig. 7.156. Vegetable looper Chrysodeixis eriosoma (Doubleday). Note the separated larger silver marks and the absence of a small silver ‘S’. (DPI&F Qld: J. Wessels)
Host range: Soybean crops and the native Glycine clandestina in the laboratory.
Risk period: Crops are at greatest risk during flowering and early podfill. Damage: Larvae feed on leaves. Monitoring: Use a beat sheet. Action level: Use the standard defoliation thresholds for summer pulses. Chemical control: Most likely controlled by pesticides targeting Helicoverpa and other more common caterpillars. Conservation of natural enemies: As for other Lepidoptera.
Fig. 7.158. Triginoides hyppasia Cramer moth (37 mm wingspan). (J. Wessels)
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Sources of information Australian Faunal Directory. (requires authorisation to view) Australian Plant Pest Database. (requires authorisation to view) Brier, H., Creagh, R., Knight, K. and Wessels, J. (2004). Common salt as a mirid management tool. In: Proceedings of the XXII International Congress of Entomology, August 2004, Brisbane, Qld. Brier, H., Collins, P., Burrill, P. and Lucy, M. (2005). Grain storage – Bruchids in mungbeans and other pulse crops: a major threat to the pulse grains industry. DPI&F Note No. ISSN 01553054. 4pp. Brier, H., Mills, G., McLennan, A., Lucy, M. and Wessels, J. (2004). What soybean insect is that? A QDPI&F Insects extension publication. 16pp. Brier, H., Knight, K. and McLennan, A. (2004). Insect management. In: Module 4 of the ‘Accredited Mungbean Agronomist Manual’ (2004 edition). 60pp plus attachments. Brier, H. (2001). Soybean tolerance to attack by Nezara viridula (L.): formulating realistic economic thresholds. Masters thesis, University of Queensland. 149pp. Brier, H., Knight, K., Wessels, J., Lucy, M. and Ferguson, J. (2001). Controlling mirids in mungbeans: how to identify mirids, assess their damage potential and control them. 2nd edition. CROPLINK Pest Update, DPI Farming Systems Institute. Q101051. 8pp. Brier, H., Dougall, A., Leighton, M., Skilton, J., Russo, P., Tuite, M. and Wessels, J. (2005). Multipest IPM in coastal soybeans. In: Proceedings of the 13th Australian Soybean Conference, Beruga, NSW. Brisbane Insects and Spiders. Caterpillars. Clarke, A.R. (1991). The current distribution and pest status of Nezara viridula (L.) (Hemiptera: Pentatomidae) in Australia. Journal of the Australian Entomological Society 31(4): 289–297. Clarke, A.R. (1990). The control of Nezara viridula L. with introduced egg parasitoids in Australia. A review of a ‘landmark’ example of classical biological control. Australian Journal of Agricultural Research 41(6): 1127–1146. Clarke, R.G. and Wilde, G.E. (1971). Association of the green stink bug and the yeast-spot disease organism of soybeans. III. Effect on soybean quality. Journal of Economic Entomology 64(1): 222. Common, Ian F.B. (1990). Moths of Australia. Melbourne University Press, Melbourne. 452pp. Coombs, M. (2003). Post-release evaluation of Trichopoda giacomellii (Diptera: Tachinidae) for efficacy and non-target effects. In: Proceedings of the 1st International Symposium on Biological Control of Arthropods, Honolulu, Hawaii, January 2002. (Ed: Van Driesche, R.G). pp. 399–406. CSIRO Entomology, Australian Insect Common Names. Department of Primary Industries and Fisheries, Queensland. Faleiro, J.R., Singh, K.M. and Singh, R.N. (1990). Influence of abiotic factors on the population build up of important insect pests of cowpea Vigna unguiculata (L) Walp. and their biotic agents recorded at Delhi. Indian Journal of Entomology 52(4): 675–680. Fehr, W.H. and Caviness, C.E. (1977). Stages of soybean development. Iowa State University Cooperative Extension Service, Special Report 80: 1–12.
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Feiber, D. (2002). Pink hibiscus mealybug biological control program established. DOACS Press Releases. Grundy, P. and Maelzer, D. (2000). Assessment of Pristhesancus plagipennis (Walker) (Hemiptera: Reduviidae) as an augmented biological control in cotton and soybean crops. Australian Journal of Entomology 39(4): 305–309. Gyawali, B.K. (1987). Influence of rice on lygaeiid and mirid bugs of soyabean. Quarterly Newsletter – Asia and Pacific Plant Protection Commission 30(3–4): 27–30. Kiritani, K. and Hokyo, N. (1962). Studies on the life table of the southern green stink bug, Nezara viridula. Japanese Journal of Applied Entomology and Zoology 6: 124–139. McDonald, F.J.D. and Edwards, P.B. (1978). Revision of the genus Oncocoris Mayr (Hemiptera: Pentatomidae). Australian Journal of Zoology Supplementary Series 26(62): 1–53. Murray, D.A.H. (1982). Life history of Monolepta australis (Jacoby) (Coleoptera: Chrysomelidae). Journal of the Australian Entomological Society 21(2): 119–122. Panizzi, A,R. and Slanski, F. (1985). Review of phytophagous pentatomids (Hemiptera: Pentatomidae) associated with soybean in the Americas. Florida Entomologist 68(1): 184–214. Prasada Rao, R.D.V.J. (2003). The host range of Tobacco streak virus in India and transmission by thrips. Annals of Applied Biology 142(3): 365–368. Pyke, B.A. and Brown, E.H. (1996). The Cotton Pest and Beneficial Guide. GOPRINT, Woolloongabba, Qld. Rees, D. (2004). Insects of Stored Products. CSIRO Publishing, Collingwod. 192pp. Rogers, D.J., Brier, H.B. and Houston, K.J. (1992). Scarabaeidae (Coleoptera) associated with peanuts in southern Queensland. Journal of the Australian Entomological Society 31(2): 177–181. Russin, J.S. (1988). Incidence of microorganisms in soybean seeds damaged by stink bug feeding. Phytopathology 78(3): 306–310. Shepard, M., Lawn, R.J. and Schneider, M.A. (1983). Insects on Grain Legumes in Northern Australia: A Survey of Potential Pests and Their Enemies. University of Queensland Press, Brisbane. Staddon, B.W. (1997). A new shield-bug species Piezodorus grossi (Het. Pentatomidae) from the Australasian region previously confused with P. hybneri (Gmelin). Journal of Natural History 31: 1859–1863. Strickland, G.R. (1981). Integrating insect control for Ord soybean production. Journal of Agriculture, Western Australia 22(2): 81–82. Turner, J.W. and Brier, H. (1979). Effect of leafhopper control on yield of peanuts and navy beans. Tropical Grain Legume Bulletin No. 16, pp. 23–25. USDA-APHIS (July 1996). Factsheet: The Hibiscus or Pink Mealybug. Velasco, L.R.I. and Walter, G.H. (1992). Availability of different host plant species and changing abundance of the polyphagous bug Nezara viridula (Hemiptera: Pentatomidae). Environmental Entomology 21(4): 751–759. Waite, G.K. (1976). Effect of various temperatures on development of the lucerne jassid Austroasca alfalfae (Evans) (Homoptera: Cicadellidae) with reference to population levels in lucerne. Queensland Journal of Agricultural and Animal Sciences 33(1): 39–42. Ward, A.L. and Rogers, D.J. (2006). Population ecology of Heteronyx piceus (Coleoptera: Scarabaeidae) in a peanut/maize cropping system. Bulletin of Entomological Research 96(2): 129–136.
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8 PULSES—WINTER M.M. Miles, G.J. Baker and W. Hawthorne
(b) Field peas at late flowering, early podding (Pulse Australia:
(a) Lupins (Pulse Australia: W. Hawthorne)
W. Hawthorne)
seeding germination growth flowering podding maturation drying harvest
Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec Jan
The winter pulse season in Australia: generalised phenology of winter pulse crops grown in temperate southern Australia. In warmer growing areas of northern WA (including the Ord River Irrigation area), Eyre Peninsula of SA and central Qld, flowering (July–September 50% flowering) and harvest may be earlier than indicated. In the Ord Irrigation Area the crop is harvested during September–October.
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PEST (major pests in bold) Germination snails redlegged earth mite blue oat mites lucerne flea false wireworms seedling maggot Growth green peach aphid cowpea aphid bluegreen aphid pea aphid brown pasture looper and green looper caterpillars Flowering and podding thrips pea weevil etiella moth native budworm corn earworm lesser budworm
Areas of crop in Australia, 5-year average to 2005 (Australian Crop Report no. 111, ABARE):
Chickpeas, Cicer arietinum. Origin: Asia. Australian area: 237 000 ha, of which 37% is grown in NSW, 27% in Qld, 17% in Vic. and 17% in WA. Faba beans, Vicia faba. Origin: Europe. Australian area: 165 000 ha, 41% of which is grown in SA, 31% in Vic., 17% in NSW and WA. Field peas, Pisum sativum. Origin: Europe. Australian area: 361 000 ha, 38% of which is grown in SA, 38% in Vic. 18% in WA and 5% in NSW. Lentils, Lens culinaris, Origin: Asia. Australian area: 116 000 ha, 64% of which is grown in Vic., 32% in SA and the remainder in NSW and WA.
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Pulse crops are grown as a cash crop, as a cereal disease control break, for diversity of weed control and because they fix their own nitrogen and aid soil fertility. Pulses often follow a cereal or canola crop, and recent practice is to sow pulses into the stubble of the previous crop rather than a cultivated paddock. Pulses usually precede a cereal or canola crop. Pulses, and lupins in particular, can also be sown into a grassy pasture or be part of re-establishing a sown pasture. In some cases the pulse is sown as a multi-purpose crop, being either grazed, cut for hay, green manured, desiccated as ‘green manure’, grazed as a standing crop or harvested for grain.
Lupins, yellow-leafed, Lupinus albus, and narrow-leafed lupin, Lupinus angustifolius. Origin: Mediterranean basin, southern Europe, Ethiopia. Australian area: 1 220 000 ha, 82% of which is grown in WA, 9% in NSW, 6% in SA and 3% in Vic.
Lupins are grown predominantly on acidic to neutral soils that are non-calcareous, and often sandier and lower in fertility. The other pulses are better suited to the less acidic, neutral to alkaline soils with heavier texture. The pulses can be grown in most rainfall districts, but lupins, field peas and vetch handle the drier rainfall condition better than the others.
Vetch, Vicia sativa. Origin: west Asia. Mainly used as a fodder crop but some grown for grain to feed stock.
The previous crop and management can influence the insect pests in the pulses (e.g. snails, slugs, false wireworm, bronzed field
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beetle, seedling maggot). The neighbouring crop, pasture or fence line can also affect the insect pest incidence (e.g. pea weevil, aphids, lucerne seed web moth) (Hamblin et al. 2000).
Natural enemies: A number of bird species eat snails, but usually do not give significant control.
Redlegged earth mite G E R M I N AT I O N
Common white snail and white Italian snail N (Main entry in Chapter 2 Cereals.)
Pest status on pulses: Major, restricted, regular. Life cycle on pulses: Refer to cereals. Risk period on pulses: Adult snails may cause some damage to germinating crops in autumn but the main risk is harvest contamination. Damage: These snails clog machinery at harvest and, as a contaminant of the harvest grain, invoke quarantine restrictions by importing countries. Field peas, lentils and vetch are particularly vulnerable. Monitoring: Monitor for snails in emerging crops by inspecting all parts of all fields. In late winter or early spring, monitor crop edges and, if snail densities warrant, apply a border bait treatment. Action level: Bait if there are five or more snails per square metre. Chemical control: Treat with formulated bran pellet baits containing a molluscicide when snails are active during cool, moist conditions: 5–80 snails per square metre, apply bait at 5 kg ha–1; over 80 snails per square metre, apply bait at 10 kg ha–1. Baiting must be completed at least 8 weeks before harvest due to the risk of bait/ residue contamination in grain. Cultural control: Refer to Chapter 2. Harvest the pulse as early as possible. A number of harvest and cleaning operations can assist in minimising snail contamination in pulse grain, including: windrowing just ahead of the harvester, ‘pusher bars’ on the harvester comb, specialised concaves and sieves, snail crushers and specialised grain cleaners. Chickpeas are less susceptible to snails and can be used as break or buffer crops.
N (Main entry in Chapter 15 Pastures—winter rainfall.)
Pest status on faba beans, field peas, lentils lupins and vetch: Major, widespread, regular. Not a recorded pest of chick peas. Risk period: Autumn–winter, particularly during seedling stage. Damage: Silvering on the upper surface of cotyledons and leaves. Lupins (epigeal emergence) are particularly susceptible at emergence as uncontrolled damage results in seedling death. Other pulses (hypogeal emergence) can recover from mite damage at emergence by re-shooting from the base. Faba beans (Fig. 8.2), field peas (Fig. 8.1) and yellow lupins are particularly susceptible to foliar damage. Vetch (Fig. 8.2), narrow-leafed and white lupins are less so, and on chick peas redlegged earth mite is not reported as a significant pest. Mites are reported as damaging lentils but they do not appear to reproduce on this host. Monitoring: Regular inspection of susceptible crops from time of first emergence of seedlings. Estimate numbers in a 10 cm × 10 cm (100 cm 2) ground area. Repeat at five to 10 sites in the crop. Avoid monitoring in bright light. Monitor on cloudy days, early morning or late afternoon. Action level: On susceptible crops, treatment is warranted if any mites are detected. Insurance spraying, even if no mites are detected, is often used as a cost-effective method of managing risk of crop failure. Chemical control: Seeds may be treated with a systemic insecticide before planting for 4–6 weeks’ protection, but seedlings should be monitored to determine the need for foliar spray protection. Alternatively, apply contact insecticides to the bare earth as a post-sowing, pre-emergent treatment to kill mites on the soil surface, or apply foliar spray treatments to the emerging seedlings to kill mites as they commence feeding. During the growing season, 261
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Cultural control: Prior growing of a non-host crop (such as cereals, chickpeas) helps reduce numbers. Control of weeds along fence lines reduces mite refuges. Natural enemies: There is no evidence of natural enemies preventing redlegged earth mite damage to pulse crops.
Blue oat mites Acarina: Penthaleidae
Fig. 8.1. A bare patch in pea crop caused by redlegged earth mite-feeding on seedlings compared with treated peas. (SARDI: P. Taverner)
application is timed 2 weeks after the first rains of autumn, when most eggs have hatched. One application is sufficient to ensure establishment and prevent yield losses. The use of the TIMERITE® management strategy in a preceding pasture phase may reduce the carryover of eggs to the cropping phase. a
b
Penthaleus major (Dugès), P. falcatus Qin & Halliday and P. tectus Halliday Distribution: In Australia, the ranges of P. major and P. falcatus overlap from southern Qld to much of the south-eastern mainland and Tas. P. major is recorded from south-western WA. P. tectus has restricted distributions in NSW and Vic. Pest status on faba beans, field peas and lupins: Major, widespread, regular. Identification: Adult mites are about 1 mm long with a purple–blue or greenish body and red legs. Blue oat mite species can be distinguished by their body hairs. P. major has comparatively long hairs running in four or five longitudinal rows down the mite’s back. P. falcatus has short hairs covering the back of its body. P. tectus has short hairs on the anterior of its body and longer hairs running in longitudinal rows on the posterior of its body. May be confused with: The redlegged earth mite which, however, lacks the orange–red oval patch found on the back of blue oat mites (Fig. 15.35). Life cycle: There are up to four generations per year. Blue oat mites reproduce asexually, developing from unfertilised eggs to produce all-female clones. Risk period: Autumn–winter, particularly during the seedling stage.
Fig. 8.2. (a) Redlegged earth mites on mature vetch leaves (SARDI: R. Matic) , and (b) faba beans are susceptible to leaf damage (Pulse Australia: W. Hawthorne) .
Damage: Silvering on the upper surface of cotyledons and leaves. Yellow lupins, field peas and faba beans are particularly susceptible, narrow-leafed and white lupins less so, and on chick peas blue oat mites are not reported as significant pests. Blue oat mites will damage lentils, but do not appear to reproduce on this host.
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Monitoring: Regular inspection of susceptible crops from the time of first emergence of seedlings. Action level: Of the susceptible crops, any observable mite activity on newly germinated plants warrants treatment. Chemical control: Preventative application of insecticide following autumn rain is less effective for blue oat mites than for redlegged earth mites. Blue oat mites may emerge earlier from summer diapause, and within the species complex there is variation in the timing of emergence. Management using insecticides is similar to redlegged earth mites but, while chemical control of P. major may be costeffective, P. falcatus appears tolerant to many insecticides. Natural enemies: There is no evidence of natural enemies preventing blue oat mite damage to pulse crops.
Lucerne flea N (Main entry in Chapter 15 Pastures—winter rainfall.)
Pest status: A major pest of lentils and vetch and a minor pest of faba beans, widespread in southern growing areas, regular occurrence. Not a recorded pest of chickpeas. May be confused with: Damage occurs at similar times to redlegged earth mite. However, the lucerne flea eats small areas (‘windows’) in leaves (Fig. 15.42), whereas the earth mite and blue oat mite produce small white feeding areas (Figs 15.37 and 15.38). Host range: Most pulses and legume pastures and weeds such as shepherd’s purse, chickweeds, sow thistles and wild radish. Risk period: Autumn–winter, particularly during the seedling stage of pulses. Damage: The young nymphs eat through the upper surface of a leaf, leaving the opposite epidermis as a ‘window’ (Fig. 15.42) characteristic of lucerne flea-feeding. Monitoring techniques: Regular inspection of the crop from time of first emergence of seedlings. Action level: At first evidence of feeding damage.
Chemical control: Pre-sowing seed treatment with insecticide. Apply contact insecticides to the bare earth as a post-sowing, pre-emergent treatment, or apply foliar spray treatments to the emerging seedlings to kill fleas as they commence feeding. Care with chemical selection is important, as some products that control redlegged earth mite are ineffective against lucerne flea. Cultural control: Clean-fallowing and elimination of weeds around the perimeter at least 1 month before sowing.
False wireworms, grey false wireworms and bronzed field beetles Coleoptera: Tenebrionidae Gonocephalum spp. and Pterohelaeus sp., Isopteron spp. and Adelium brevicorne Blessig N (Main entry for false wireworms in Chapter 5
Oilseeds.)
Pest status: Minor pests of winter pulses, restricted, irregular. Identification: See Chapter 5. Life cycle on pulses: These species are irregular soil-inhabiting beetles with life cycles of usually one generation per year. Adults of these may be found in fields from late spring to autumn. They are often associated with the use of minimum tillage, where trash helps support the developing larvae. Risk period: During germination and crop establishment. Damage is most severe in crops sown into dry seed beds and when germination is slowed by continued dry weather. Occurrence of damage is unpredictable. Damage: Larvae eat cotyledons and roots of germinating seedlings, and a symptom of their activity is small bare patches and sections of missing rows in the crop. They generally prefer light, dry soils. Monitoring: Monitor paddocks prior to sowing. See monitoring for wireworms in Chapter 5. Chemical control: Gonocephalum may be controlled using a registered seed treatment. 263
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Pest status: Minor, widespread, irregular. Identification: Medium-sized, wingless forms may be coloured in one of various shades of green. The winged form has a black patch on the abdomen. May be confused with: Other greenish coloured aphids. The green peach aphid is not shiny, does not have the bluish colouration of the bluegreen aphid, and is smaller than pea aphid (Table 15.1, Fig. 15.46). Fig. 8.3. Seedling maggot, 4 mm long. (SARDI: G.J. Baker)
Isopteron is controlled using soil-incorporated insecticide prior to sowing. Adelium may be controlled by spraying the seedling crop soon after germination. Cultural control: Post-sowing rolling, and increased seedling density, can reduce the impact of these pests on crop establishment.
Seedling maggot Delia platura (Meigen) Diptera: Anthomyiidae Pest status: Minor, restricted, irregular. Not a recorded pest of chick peas. Identification: Larvae are legless, tapered, white to creamy maggots (Fig. 8.3), which grow to 6–7 mm length and may pass through a number of generations per year. Damage: They occasionally cause damage to pulse crops, particularly lupins. They favour moist soils with rotting organic matter. The larvae (maggots) attack the stems just below the soil surface. Damage may appear similar to root rot until the stem is split open to reveal the larva. In some instances onion seedling maggot may be a secondary pest, infesting plant tissues that have become diseased with a fungal or bacterial pathogen.
G ROW T H
Green peach aphid Myzus persicae (Sulzer) Hemiptera: Aphididae Distribution: Probably a native of Asia, now distributed worldwide. In Australia, its range includes all cropping areas.
Host range: Hosts of the green peach aphid include many families of plants, including tree and field crops. The green peach aphid may be found on most winter pulse crops in southern Australia but it is often not the most common aphid species. Life cycle on pulses: The green peach aphid has a number of generations each year. Winged adults fly into a crop from early winter onwards and form colonies of wingless aphids by asexual reproduction. Aphid build-up is favoured by fine, warm weather. By early summer, some winged aphids are found in these colonies. Development of aphid numbers is retarded by hot days in early summer and when its host plant senesces. Risk period: The main risk period for disease transmission occurs from early winter onwards, when winged aphids fly from infected hosts. Damage: The green peach aphid causes damage mainly as a vector of virus diseases. Monitoring: Inspect crop and sample leaves for winged aphids from early winter onwards and for aphid colonies in spring and early summer. Action level: Virus transmission may occur on crops at very low aphid densities and transmission may have occurred by the time aphids are detected. There is no recommended action threshold to prevent virus infection. Chemical control: Except in seed crops, chemical control is usually not cost-effective. Cultural control: See box. Natural enemies: Field crop aphids are often parasitised, but the parasitoids usually occur too late to prevent disease transmission.
Cowpea aphid Aphis craccivora Koch Hemiptera: Aphididae
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Fig. 8.4. Colony of cowpea aphid on a faba bean shoot. One shiny black adult (2 mm long) appears on the lower left; the remainder are immature with a waxy coating. (SARDI: G.J. Baker)
Distribution: A native of Europe, now distributed worldwide. Pest status on pulses: Major, widespread, irregular. Identification: Adult aphids are shiny black. May be confused with: Unlikely to be confused with other aphids of winter pulses, as it is the only black aphid numerous on these crops. Host range: Recorded on all winter pulses in Australia, and is also found on many other legume and non-legume plants. Life cycle on winter pulses: Similar to the green peach aphid. Risk period: Dry, warm winter–spring weather favours aphid build-up. High aphid densities combined with moisture or other stresses weaken plants and reduce yield. The main risk period for disease transmission is in early
Fig. 8.6. Lupins susceptible to aphid damage (right) and resistant variety in pod (left). (Pulse Australia: W. Hawthorne)
spring, when winged aphids fly from infected hosts. In early summer, high aphid densities combined with moisture stress weaken plants. Damage: The cowpea aphid causes damage ainly as a vector of virus diseases. Dense colonies may cause physical damage to chickpeas, faba beans, lentils, lupins and vetch, especially when plants are under water stress (Fig. 8.4). Monitoring techniques: Inspect crop and sample leaves for winged aphids from early winter onwards and for aphid colonies in spring and early summer. Action level: Virus transmission may occur on crops at very low aphid densities and transmission may have occurred by the time aphids are detected. There is no recommended action threshold to prevent virus infection. Chemical control: Except for seed crops, chemical control to prevent virus transmission is usually not cost-effective (Bwye et al. 1997). Spraying of dense colonies of cowpea aphid in spring may prevent yield losses from physical damage. Cultural control: See box. Choose a pulse variety with resistance to aphids or the important viruses transmitted by them. Natural enemies: Parasitoids are often found in aphids. They usually occur too late to prevent disease transmission in pulses. Heavy rainfall can reduce aphid populations significantly.
Bluegreen aphid Fig. 8.5. Cowpea aphid colonies on a faba bean plant. Combined with water stress, these aphids may retard crop growth. (SARDI: G.J. Baker)
Acyrthosiphon kondoi Shinji Hemiptera: Aphididae 265
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N (Main entry in Chapter 15 Pastures—winter rainfall.)
Distribution: Originally from Asia, now distributed worldwide. In Australia, distributed in southern pulse areas. Pest status: Major pest of lupins, restricted, irregular. Identification and life cycle: See Chapter 15 (Table 15.1, Fig. 15.47). Host range: Many legume species. In southern Australia, the greatest densities are found in lucerne crops. Risk period: Spring to early summer, when winged adults fly into the crop from perennial legume hosts. Damage: The bluegreen aphid causes damage mainly as a vector of virus diseases. Monitoring techniques: Inspect crop and sample leaves for winged aphids in spring and for aphid colonies in spring and early summer. Action level: Virus transmission may occur on crops at very low aphid densities, and transmission may have occurred by the time aphids are detected. There is no recommended action threshold to prevent virus infection Chemical control: Usually not cost-effective. Cultural control: See box. Choose a pulse variety with resistance to aphids or the important viruses transmitted by them.
Risk period: Spring to early summer, when winged adults fly into the crop from perennial legume hosts. Damage: The pea aphid is a vector of bean yellow mosaic disease and bean leafroll virus in chickpea. The aphid is rarely associated with physical damage to pulse crops in Australia. Monitoring: Inspect crop and sample leaves for winged aphids in spring and for aphid colonies in spring and early summer. Action level: Virus transmission may occur on crops at very low aphid densities and transmission may have occurred by the time aphids are detected. There is no recommended action threshold to prevent virus infection. Chemical control: Usually not cost-effective. Cultural control: See box. Choose a pulse variety with resistance to aphids or the important viruses transmitted by them. Natural enemies: Parasitic wasps and fungus disease.
Looper caterpillars Chrysodeixis sp. Lepidoptera: Noctuidae Brown pasture looper, Ciampa arietaria, Guenée Lepidoptera: Geometridae. Green looper caterpillars, Chrysodeixis spp. Lepidoptera: Noctuidae N (Main entry in Chapter 5 Oilseeds.)
Pea aphid
Pest status: Minor, restricted, irregular.
Acyrthosiphon pisum (Harris) Hemiptera: Aphididae
Identification: Looper caterpillars walk in a looping motion, which is enabled by abdominal legs (prolegs), in addition to three pairs of legs on the thorax.
N (Main entry in Chapter 15 Pastures—winter rainfall.)
Distribution: A native of Europe, now distributed worldwide. Pest status: A minor pest of faba bean and field pea crops, widespread, irregular. Identification and life cycle: See Chapter 15 (Table 15.1, Fig. 15.46). Host range: Many legume species. In southern Australia, the greatest densities are found in lucerne crops. Pea aphids are recorded in generally low densities on faba bean, chickpea and field pea crops.
The female moths of the brown pasture looper (Fig. 5.13) lay eggs in autumn. The larvae are very small when the crop emerges, and grow up to 20 mm in length, are grey or brown, and have a dark back stripe bordered by a thin yellow line on either side running along their backs. Damage by brown pasture loopers has mainly been recorded from lupin crops. High larval densities can reduce plant numbers, and defoliation may reduce vigour and yield. Damage is most severe along edges of crops near pastures containing broadleaf weeds.
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Aphid transmission of virus diseases in Australian pulse crops Worldwide, many species of aphids are vectors of plant disease viruses and many of these aphids are now in Australia. Symptoms of these diseases include yellowing or mottling of leaves, leaf distortion, stunted plant growth and ultimately loss of yield. These symptoms may be confused with those of nutrient deficiency, herbicide damage and waterlogging. Aphids contact these viruses by feeding on infected plants and transmitting the virus on their mouthparts to uninfected plants. Some of the viruses are seed-borne, and aphids transmit the virus from an infected plant to nearby uninfected plants, resulting in a patchy distribution of the disease. Other diseases are transmitted from alternative hosts by winged aphids that fly into growing pulse crops, producing disease symptoms around the perimeter of the crop. The main species of aphids that form colonies in pulse crops include cowpea aphids, green peach aphids and bluegreen aphids. Virus outbreaks are generally more common in areas of higher rainfall, or in wetter seasons when aphid and virus hosts/ reservoirs are abundant in and around pulse crops (Jones 1997). Bean yellow mosaic virus damages chick peas, faba beans, field peas lentils and lupins. The virus is transmitted by aphids from legume pastures to crops during winter and early spring. The virus does not persist in aphids, which lose the infection after one or two feeds on uninfected plants. Bean yellow
mosaic virus is found more frequently on plants at the edge of crops than in the centre, reflecting migration patterns of the aphids (Jones 1997). Cucumber mosaic virus damages chick peas, lentils and lupins (narrow-leafed and yellow) (Bwye et al. 1995). Pea seedborne mosaic virus damages chick peas and lentils and causes minor damage to faba beans and field peas. Alfalfa mosaic virus damages chick peas and lentils. These three viruses are introduced to crops by infected seed, and aphids spread the disease to nearby plants. These viruses do not persist in their aphid host. Subterranean clover red leaf virus and related viruses damage faba beans (Fig. 8.7), chick peas and lentils. These viruses are transmitted from clover hosts to crops by cowpea and other aphids during spring. These viruses persist in the aphid, which may be infective for many weeks. Additional persistent aphid transmitted virus diseases that have been recorded on chickpeas include lettuce necrotic yellows virus, bean leafroll virus, beet western yellows and subterranean clover stunt virus (Swinghamer et al. 2002). Chickpea chlorotic dwarf virus is a persistent virus transmitted by leafhoppers. (Swinghamer et al. 2002). In India, the leafhopper vector is reported as Orosius orientalis (Horn et al. 1993). A combination of methods can be used to manage virus diseases. Use of tolerant varieties
and seed free of virus are important when available. Avoidance of sowing near infected host plants reduces transmission risk and early sowing allows crop growth before the time when aphids invade. Promotion of a vigorous plant canopy allows healthy plants to suppress sick plants. Minimising the perimeter of a field and avoiding bare patches reduces edges from which aphids invade the crop. Planting of a cereal crop around the perimeter or within the crop may help trap aphids and disinfect their mouthparts of virus. Retention of stubble from the previous crop or straw ground cover may deter aphids from landing (Jones 2001). Insecticidal control is usually not cost-effective as small numbers of aphids may have transmitted disease some weeks before symptoms are apparent (Bwye et al. 1997).
Fig. 8.7. Faba beans with symptoms of subterranean clover red leaf virus, persistently transmitted by aphids. (UnivAd: J. Randles)
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Green looper caterpillars (Fig. 5.15) grow up to 40 mm in length and are coloured green with thin white stripes running the length of the body. In spring, they may be found in pulse crops eating leaves, but do not enter pods. They do not cause economic damage in pulse crops, and should not be confused with native budworm.
a
F LOW E R I N G A N D P O D D I N G b
Flower thrips Thysanoptera: Thripidae Including possibly:
Plague thrips, Thips imaginis Bagnall Onion thrips, T. tabaci Lindeman Tomato thrips, Franklinella schultzii (Trybom) Western flower thrips, F. occidentalis Pest status on pulses: Minor, restricted, irregular.
Fig. 8.8. (a) The adult pea weevil (length 5mm) which, in spite of its common name, is not a true weevil, and (b) Harvested peas with windows from which adult pea weevils will emerge in autumn, or as a response to jolting. (SARDI: G.J. Baker)
Pest status: Thrips damage to faba beans is reported from NSW and large numbers of thrips may cause bud and flower loss in some pulse crops.
Distribution: A native of west Asia, now found in most field pea-growing areas of the world, including Australia. Not in Tas. or some regions of NSW.
Damage: Thrips feeding on flowers of faba beans may cause discoloration of the seed coat and distortion of pods, making them difficult to harvest. Risk of damage is increased during dry weather.
Pest status: Major on field peas, restricted, regular.
N (Main entry in Chapter 7 Pulses—summer.)
Monitoring: Thrips numbers can be estimated by picking and opening a representative sample of flowers from across the field. Action level: An action threshold of two to four thrips per flower is suggested for NSW crops. Chemical control: Thrips may be controlled by registered insecticides but the cost-effectiveness is yet to be reported.
Identification: In spite of its name, the pea weevil is not a true weevil but belongs to another group of plant-feeding beetles. The adult is about 5mm long and has a brown body flecked with white, black and grey patches. The wing covers are shorter than the abdomen (Fig. 8.8). The egg is orange, cigar-shaped and 1.5mm long (Fig. 8.9). The larva is legless and white with a small brown head. May be confused with: The pea weevil is unlikely to be confused with any other field pest of peas. However, in storage it may be confused with other bruchid species (see Chapter 7).
Pea weevil
Host range: Field peas.
Bruchus pisorum (Linnaeus) Coleoptera: Chrysomelidae
Life cycle on field peas: The pea weevil has one generation each year. Adults become active in
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adults to emerge. Adult beetles fly to hibernation sites, which may include sheds, silos and grain bins, under the bark of trees, particularly pines, and in cracks and crevices in posts. They remain in hibernation during summer, autumn and winter, and resume activity when spring temperatures reach about 20°C and fly to the edge of the nearest pea crop. Risk period: From when pods first appear in spring to drying of the pods in early summer. Damage symptoms: A circular hole of 3 mm diameter in pea seeds. Heavily infested crops may suffer significant yield loss (grain weight) from feeding by pea weevil. Fig. 8.9. Eggs (each of 1.5 mm length) of the pea weevil are laid on pea pods; detail of an egg (inset). (SARDI: G.J. Baker)
early spring when temperatures reach around 20°C and move into pea crops from their hibernation sites (under tree bark, posts, along fence lines, around sheds, bins, etc.). Dispersing adults characteristically invade from the edge of the field, and often do not move more than 50 m into the crop. Females must feed on pea pollen to their mature eggs, which usually takes several weeks. When mature, females lay eggs on the outside of green pea pods. The orange eggs are easily seen by the naked eye glued to the pod tissue. When hatched about 2 weeks after laying, the larva channels through the tissue directly below the egg and does not come into contact with the exposed surface of the pod. More than one egg laid on the surface of a pod usually results in an equivalent number of damaged seeds. The minute larva enters a seed and feeds for 5–7 weeks, in which time it grows to occupy much of the seed. When it is nearly fully grown, the larva chews a circular exit hole, 3 mm diameter, leaving only a translucent skin above the hole (Fig. 8.8). By this time, the seed has dried and may be harvested during larval development. The larva pupates in the seed (pupation usually occurs after harvest). The pupa develops into the adult, usually during the time when the grain is put into storage. Adults emerge over several months starting in mid-December. Jolting of the seed during handling may induce a number of
Monitoring techniques: Use a sweep net to detect when and how many adult beetles occur in the crop. Start in early spring during flowering, but before first pods have formed. Sweep every 5–7 days around the crop edge, about 2 m into the crop. Take 10 sweeps at a time and count the number of pea weevils caught. Sweep at five to 10 sites along the crop edge, particularly where trees and other hibernation sites are located. Action level: Spray if an average of one or more adults are collected per 10 sweeps. Chemical control: A spray to kill beetles should be applied before they lay eggs on pods. The spray should be applied if the action level is exceeded and, if so, when the first pods appear in the crop. A spray applied to the border (40–50 m) of the crop, or as indicated by monitoring, is usually sufficient to prevent damage (Fig. 8.10). Cultural control: Early harvest of the pea crop when the insect is still in the larval stage removes it from the field before adults emerge. If the crop is left in the field until mid-December, adults start to emerge and will infest the following season’s crop. Sheep in the pea field following harvest will eat any fallen seed and further reduce the carry-over of pea weevil. Post-harvest fumigation: Infested seed may lose up to 70% of weight after harvest as larvae continue to feed. To prevent this, to meet quality standards for sale, and to reduce carry-over into the following season, fumigation of the seed prior to storage may be necessary. However, planting clean seed and pre-harvest control in 269
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a
T b
Adult density high 50m
medium low
T
tree c
Fig. 8.10. Typical distribution of adult pea weevil in a large pea field during spring. A single perimeter spray usually controls most damage. (SARDI)
the field are more cost-effective than postharvest control. Natural enemies: No effective natural enemies have been recorded in Australia. Maintenance of area-freedom (Tas. and parts of NSW): Ensure all purchased pea seed is phosphine-fumigated prior to delivery. The fumigation must occur in a sealed silo, or under gas-proof sheeting, for 10 days. If possible, avoid purchasing seed from a pea weevil-infested district.
Etiella moth (or lucerne seed web moth)
Fig. 8.11. (a) Etiella moth adult, 15 mm long, showing the prominent white streak along the forewing and the prominent ‘snout’; (b) egg, 0.6 mm-long laid near a developing lentil pod; and (c) fully grown larva. (SARDI: C. Krawec)
Etiella behrii (Zeller) Lepidoptera: Pyralidae N (Main entry in Chapter 13 Pastures—summer rainfall.)
Distribution: East and South-East Asia, some pacific Islands and Australia. In Australia, its range includes all pulse areas but it is not common in Tas. Pest status on pulses: Major on field peas, lentils and lupins, restricted, irregular. Identification: Eggs are oval-shaped, 0.6 mm long, colourless when first laid, soon turning yellow, and prior to hatching (Fig. 8.11) a black head can be seen in the egg. Eggs are laid singly. The mature larva has a pale golden-brown head, behind which is a dark shield, with the
remainder of the body green with a pinkish tinge (Fig. 8.11). The adult is a greyish moth, 10– 15 mm long, with a snout-like head. The leading edge of the forewing has a white stripe along its full length and at the base of the forewing is an orange band. At rest, the moth has an ‘alert’ appearance, with the front end of the body raised (Fig. 8.11). May be confused with: Larval damage to field pea seeds is similar to that of native budworm. Damage by this moth is sometimes wrongly attributed to native budworm. Host range: Legumes, including lucerne, lupins, lentils and some native legumes.
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a
chews into the pod and feeds on the seeds as its growth progresses. Risk period: When pods are green. Dry pods are not at risk.
b
Damage: A significant pest of field peas and lentils in eastern Australia. Pod set can be reduced by etiella damage to flowers, and pulse grain quality is downgraded by even pin-prick holes or small ‘drill’ holes in otherwise intact seed. Gross larval damage is indicated by the jagged remains of eaten seeds and frass in the pod. Monitoring techniques: Monitor for moth flights using light traps; detection in sweep nets is unreliable. Adults may be seen making short flights in the crop when disturbed and alight on the undersides of leaves. In lentil crops, adults may be monitored using a sweep net during the risk period. Make at least three lots of 20 sweeps at random locations in the crop.
c
Action level: Evidence of relatively few adults in a crop may be indicative of economic losses. An indicative threshold for lentils is two or more adults per 20 sweeps. Chemical control: Should aim to kill adults before egg laying. Larvae cannot be controlled once inside the pods. If the crop has been sprayed for native budworm, an etiella spray will not be necessary for a further 10–14 days.
Fig. 8.12. (a) Field pea pod opened to show a larva of etiella moth (length 20 mm) eating fresh peas; (b) webbing and frass are indicators of lucerne-seed web moth damage (SARDI: P.T. Bailey) ; and (c) lentil grain damaged by etiella larvae (SARDI: C. Krawec) .
Life cycle on pulses: Etiella moth has three or four generations per year, of which only the spring generation damages winter pulses. Larvae from the previous autumn overwinter in a pupal chamber in the soil, and adults emerge from September. Adult moths fly into nearby crops and lay eggs on developing pods. Female moths lay about 200 eggs, which may hatch in 1–14 days depending on temperature. The egg hatches into a small larva, which spins a protective web around itself. Both the web and entry hole soon disappear. The small larva then
Natural enemies: Six species of insects (five wasps and a fly) parasitise up to 10 per cent of large larvae when they emerge from the pods and build their webbed retreats. Disease has been recorded as killing up to 40 per cent of larvae (Austin et al. 1993).
Native budworm Helicoverpa punctigera (Wallengren) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Pest status on pulses: Major, widespread, regular. Identification: See Chapter 3. May be confused with: In harvested lentil and field pea seed, damage is similar to that of the lucerne seed web moth but lacks webbing. In 271
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field peas, the jagged edge of native budworm damaged seed contrasts with the smooth, cylindrical exit-holes made by pea weevil. Corn earworm, Helicoverpa armigera (Hübner), larvae occur in low frequency in winter pulse crops. Looper caterpillars also occur in pulse crops during spring. Hosts: All broadleafed crops and many native and introduced plants are larval hosts of native budworm. All pulses are hosts. Native budworm may lay eggs on a number of weeds commonly found in pulse crops.
a
b
Life cycle on pulses: Only the spring generation damages pulses. Adult moths fly into southern crops from inland winter breeding areas. Female moths lay eggs on leaves, flowers and fruit. Young larvae graze the surface of leaves, leaving skeletonised areas of leaf. Older larvae enter pods and eat seeds, leaving a jagged feeding area. Risk period: When green pods are present, usually late spring to pre-harvest. Damage: Young larvae feed on leaves. High numbers on vegetative chickpeas are recorded as defoliating and lopping branches in northern regions. Where moisture is adequate, most pulse crops compensate for damage prior to flowering (Fig. 8.13). Later, larvae damage buds, flowers, developing and filling pods by their feeding. Larger larvae burrow into filling pods, consuming some or all of the developing grain. One larva may destroy several seeds in up to four or five pods. Damaged seed remnants have jagged edges, and pods may contain faecal pellets. Monitoring: Light traps or pheromone traps can indicate the presence of immigrant adults in a district. At present, no good relationship between the number of budworm moths trapped and subsequent egg laying and damage to pulse crops has been demonstrated. Searching for eggs gives an indication of the activity of native budworm in the crop. Eggs are usually laid by female moths on the upper surface of leaves but may also be found on the lower surface of leaves and on flowers and pods. Start searching for eggs in mid-August. ‘Indicator plants’, such as unopened soursob flowers, may be inspected in addition to monitoring crop plants.
Fig. 8.13. (a) Native budworm (arrowed) feeding on faba bean leaves causes little damage but (b) is indicative of likely seed damage at harvest. (Pulse Australia: W. Hawthorne, SARDI: P.T. Bailey)
Larval monitoring is the most widely-used method of deciding when and whether to apply an insecticide. For field peas, chickpeas, lentils and grain vetch sown on narrow rows, or broadcast, use a sweep net to drag through the top one-third of the crop. Swing the net in an arc of 180° in front of the body. Stop after five or 10 sweeps and count and record the number and size of native budworm larvae collected. Repeat at five to 10 sites in the crop and average the count for decision making (Fig. 8.15). Chick pea and faba bean crops planted on 1 m row spacings are checked using a 1 m beat sheet to estimate larval density and stage of development. Lie the sheet on the ground across the space between the rows, and over the adjacent row. Vigorously beat the plants in 1 m of row onto the beat sheet and count and size the dislodged caterpillars. Repeat at 10 randomly selected sites. Monitor crops weekly from flowering to dessication, more frequently during podding or when populations are close to threshold.
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a
b
Fig. 8.15. A sweep net may be used to monitor for both native budworm larvae and pea weevil adults in field peas. The brown strip in the field pea crop was untreated and has been eaten out by native budworm. (SARDI: P.T. Bailey)
weeks’ residual life will kill larvae that hatch from eggs during this period. Fig. 8.14. (a) Native budworm eggs on the bud of weed (soursob) that matures earlier than grain legume crops, and (b) ‘brown ring’ stage, indicative that the eggs are mid-way through development. (SARDI: G.J. Baker)
Action level: The level of damage at harvest is directly related to the number of budworm per square metre. For field peas and lupins, an average of one to two larvae in 10 sweeps indicates treatment may be necessary. For chickpeas, the suggested thresholds are one to four larvae per square metre (assessed by beat sheet), and five larvae per 10 sweeps (assessed by sweep net). For faba beans, treat if there is an average of one or more larvae per sample. The action level is lowered as the crop sets more pods and starts to mature, or if the crop is moisture stressed, as it is less able to compensate for damage. Action thresholds will be higher or lower depending on crop value, cost of control and tolerance of feeding damage. For example, some crops have a lower action level when they are destined to be marketed for human consumption rather than stock feed or processing. Chemical control: Insecticides are generally more effective on younger larvae (first to third instar) than older larvae. Aim to control larvae before they enter pods where they are protected from insecticides. An insecticide with two
Biocidal control: Commercially available nuclear polyhedrosis virus (NPV) is effective against native budworm (Fig. 8.16), and used on chickpeas gives about 80% control. NPV may not reduce an extremely high population below threshold. NPV should only be used in chickpea with a recommended additive to boost efficacy. A bacterial product, the crystalline spores of Bacillus thuringiensis (Bt) is also effective against Helicoverpa spp. in chickpeas. Natural enemies: Different species of Trichogramma (Figs 3.50 and 3.51) have been recorded as naturally occurring parasites of native budworm eggs. However, only in western Victoria has Trichogramma sp. (near ivalae) in some years prevented damage to field pea crops, and obviated the need for spraying (Ridland et al.
Fig. 8.16. Helicoverpa larva killed by formulated nuclear polyhedrosis virus (NPV) sprayed onto a chickpea crop. (DPI&F Qld: M. Miles)
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a
a
b
b
Fig. 8.17. (a) Tachinid fly preparing to lay its egg on the larva of Helicoverpa armigera; (b) three tachinid eggs laid on a caterpillar. (DPI&F Qld: J. Wessels)
1993). Trichogramma are not active in chickpea crops. Tachinid flies (Fig. 8.17) (most commonly Carcelia illota Diptera: Tachinidae) and the hymenopteran parasitoid, Heteropelma scaposum Hymenoptera: Ichneumonidae, are the most a
b
Fig. 8.19. Ichneumonid wasps: (a) banded caterpillar parasite, Ichneumon prommisorius, is a natural enemy of noctuid larvae in northern winter pulse areas (K. Power) ; and (b) orchid dupe, Lissopimpla excelsa (Costa), pollinates native orchids and also parasitizes noctuid caterpillars, among others (K. Power) .
frequently observed larval parasitoids of native budworm larvae on chickpeas in southern Qld. The larval parasitoid Netelia producta (Hymenoptera: Ichneumonidae) (Fig. 8.18) and the pupal parasitoid Ichneumon prommisorius (Ichneumonidae) (Fig. 8.19) are irregular parasitoids of Helicoverpa species in chickpeas. Several species of predatory bugs, including Nabis tasmanicus (Nabidae) (Fig. 8.20) and the spined predatory shield bug, Oechalia schellenbergii (Pentatomidae) (Fig. 8.20), prey directly on native budworms and transmit NPV disease through their faeces after feeding on an infected larva. The polyhedra in the faeces remain infective in the crop for several days (Cooper 1981).
Corn earworm Fig. 8.18. (a) Adult orange caterpillar parasite, Netelia producta (K. Power) , and (b) egg laid by Netelia producta onto native budworm larva. On hatching, the parasitoid larva enters the body of its host (DPI&F Qld: J. Wessels) .
Helicoverpa armigera (Hübner) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
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a
Chickpeas as trap crops to reduce carry over of insecticide-resistant corn earworm into summer crops
b
Chickpea trap crops have been used in areawide management of corn earworm in southern Qld to capture the local overwintering population as it emerges from diapause. This local population are the survivors from the previous season and potentially carry the genes for resistance to insecticides. Chickpea trap crops are planted on 1% of a field or a minimum of 2 ha. The trap crops are planted as a winter crop but later than commercial chickpea crops so that flowering occurs as commercial crops senesce, and there are few other hosts in the cropping area. The aim is to capture a large proportion of the local H. armigera. In this strategy, the trap crops are used to remove a proportion of the local population before it can move into the summer crops. Corn earworm larvae in the trap crop are killed by slashing and cultivation.
Natural enemies: As for native budworm.
Lesser budworm Fig. 8.20. Predatory bugs: (a) Nabis tasmanicus (SARDI: G.L. , and (b) a spined predatory shield bug feeding on Helicoverpa larva (DPI&F Qld: C. Freebairn) . Predatory bugs can spread NPV in their faeces.
Baker)
Pest status: Corn earworm frequently occurs in pulse crops in northern NSW and southern Qld, where it is a pest of chickpeas, and it may sometimes occur in low numbers in pulse crops in more southerly areas. Monitoring: As for native budworm. Because of its resistance to a number of insecticides, application of chemical control to populations of corn earworm must target small larvae (< 7 mm). It is difficult to accurately distinguish corn earworm and native budworm in the field, so it is prudent to assume mixed populations in all northern regions (north of Dubbo, NSW). Action level: As for native budworm. Biocidal control: As for native budworm.
Heliothis punctifera Walker Lepidoptera: Noctuidae Distribution: Australia. Pest status: Minor, widespread, irregular. Identification: The striped larvae resemble larvae of Helicoverpa species. May be confused with: Helicoverpa punctigera, which however has dark body hairs (Fig. 3.21) compared with the light body hairs of Heliothis punctifera. Helicoverpa armigera also has light body hairs (Fig. 3.21) but the last spiracle is in the area of the light stripe, compared with Heliothis punctifera, where the spiracle lies in the dark area (arrowed in Fig. 8.21). Adults (Fig. 8.22) have patterned forewings. Hosts: Crop hosts include cereals, lentils, lupins, medics and vetch. Native hosts include Chenopodiaceae, Compositae and Labiatae (Matthews 1999). 275
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Fig. 8.23. Heliothis punctifera larvae grazing leaves of field peas. They also eat seeds of legume crops and cereal heads. (SARDI: G. Caon) Fig. 8.21. A larva of Heliothis punctifera may be distinguished from Helicoverpa punctigera by its white hairs and from H. armigera by the last spiracle (arrowed) in the dark band. (SARDI: G. Caon)
Life cycle: The species is known from inland breeding areas and has been caught in light traps over eastern Australia as far south as Tas.
It was not recorded as a common species in southern cropping areas until the spring of 2005, when large numbers of moths migrated and laid eggs on spring crops. It is possible that the life cycle is similar to that of native budworm. Risk period: Spring. Damage: Damage appears similar to that of native budworm. Larvae graze on foliage, causing little damage, but also eat seeds, and some grain losses have been recorded. Vetch seems particularly susceptible to damage. Monitoring in pulse crops: Similar to native budworm. Action level: Unknown, but those used for native budworm may be used as an indicative level.
Fig. 8.22. Heliothis punctifera moth. (SARDI: G. Caon)
Control: Provisional control options are similar to those of native budworm.
Sources of information Austin, A., White, T.C.R, Maelzer, D.A. and Taylor, D.G. (1993). Biology of Etiella behrii Zeller (Lepidoptera: Pyralidae): a pest of seed lucerne in South Australia. Transactions of the Royal Society of South Australia 117: 67–76. Bwye, A., Jones, R. and Proudlove, W. (1995). Cucumber mosaic virus in lupins. W. A. Journal of Agriculture 36: 124–130. Bwye, A.M., Proudlove ,W., Berlandier, F.A. and Jones R.A.C. (1997). Effects of applying insecticides to control aphid vectors and cucumber mosaic virus in narrow-leafed lupins (Lupinus angustifolius). Australian Journal of Experimental Agriculture 37: 93–102. Cooper, D.J. (1981). The role of predatory Hemiptera in disseminating a nuclear polyhedrosis virus of Heliothis punctiger. Journal of the Australian Entomological Society 20: 145–150. Hamblin, J., Hawthorne, W. and Perry, M. (2000). Regional reviews: The Australian scene. pp. 131–142. In: Linking Research and Marketing Opportunities for Pulses in the 21st Century. R. Knight (ed.). Kluwer Academic Publishers. 276
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Horn, N.M., Reddy, S.V., Roberts, I.M. and Reddy, D.V.R. (1993). Chickpea chlorotic dwarf virus, a new leafhopper-transmitted geminivirus of chickpea in India. Annals of Applied Biology 122: 467–479. Jones, R. (1997). Bean yellow mosaic virus in lupins. W.A. Journal of Agriculture 38: 42–48. Jones, R.A.C. (2001). Developing integrated disease management strategies against non-persistently aphid-borne viruses. A model program. Integrated Pest Management Reviews 6: 15–46. Matthews, M. (1999). Heliothine Moths of Australia: A Guide to Pest Budworms and Related Groups. CSIRO Publishing, Collingwood. Ridland, P.M., Smith, M.A., McDonald, G. and Cole, P. (1993). Developing efficient management procedures for control of budworm in field peas. Australia: Grains Research and Development Corporation, final project report, Project DAV 17G. Swinghamer, M., Schlig, M., Moore, K., Kumari, S., Srivastava, M., Wratten, K., Murray, G., Knights, T., Bambach, R. and Southwell, R. (2002). The virus situation in chickpea, faba bean and canola in New South Wales and Southern Queensland. Updates of research in progress at the Tamworth Agricultural Institute pp. 44–47.
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9 RICE M.M. Stevens N Oryza sativa, Poaceae. Origin: southern Asia
(a) Rice plant
(b) Rice paddock
The temperate rice season
sowing establishment vegetative flowering grain filling harvest Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun Jul Aug
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Darwin
2
1
3 4 5
NT QLD WA
Brisbane
SA NSW
Perth Adelaide
Former tropical rice areas 1 Ord River (WA) 2 Humpty Doo (NT) 3 Mareeba (Qld) 4 Ingham (Qld) 5 Ayr (Qld) Current temperate rice area 6 Murrumbidgee and Murray Valleys
6 VIC
ACT
Sydney
Canberra
Melbourne
TAS Hobart
Rice growing areas: current temperate rice area (Murrumbidgee and Murray Valleys, NSW); former tropical rice areas are shown by dots at Mareeba, Ayr, Ingham, NT (Humpty Doo) and Ord River.
PEST (major pests in bold) Temperate rice Establishment aquatic earthworms water snails tadpole shrimp yabby bloodworms rice leafminer Vegetative growth to harvest locusts and grasshoppers rice root aphid green vegetable bug rice stink bug
PAGE
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PEST (major pests in bold)
PAGE
common armyworm sugarcane and maize stemborer hairy rice caterpillar Tropical rice (not presently grown commercially; prior pest status noted) Establishment bloodworms rice leafminer Vegetative growth to harvest rice leafhopper brown planthopper paddy bug rice stink bug caseworm rice leaffolder white rice stemborer darkheaded riceborer striped riceborer armyworms: common, northern, dayfeeding and lawn rice skipper
Temperate rice is currently grown under irrigation in the Murrumbidgee and Murray Valleys of southern NSW, and is limited by climate to a single crop per year, sown in October–November and harvested in March– May. Tropical rice has previously been produced in the NT, the Ord region of WA, and in north Qld (see map). In these areas it was possible to grow two crops per year (wet season and dry season); however, tropical rice is no longer produced commercially for various reasons including lack of processing infrastructure, variable grain quality, pest damage, and the cost of irrigation infrastructure. Upland (rain-fed) rice has not been produced commercially in Australia.
Temperate rice Temperate rice production in NSW varies seasonally according to the availability of irrigation water. Production in 2003–2004 was approximately 530 000 t of paddy; however, production has been as high as 1.7 million t. Yields vary with seasonal conditions, but typically average around 8–9 t ha–1. Over 85% of rice crops are sown aerially (into flooded fields) or are dry-broadcast and then flushed once prior
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292 292 292 292 293 293 294 294 294 294 294 295 295
to permanent flooding. The remaining crops are drill- or sod-sown into prepared seed beds or clover pastures and flushed several times before application of permanent water.
E S TA B L I S H M E N T
Aquatic earthworms Eukerria saltensis (Beddard) Oligochaeta: Ocnerodrilidae Distribution: Introduced, believed to originate from South America. A widely distributed peregrine species thought to have been introduced into Australia during the 19th century. Within Australia, known from NSW, Qld, Vic. and WA. Pest status in temperate rice: Major, widespread, irregular. The worst damage occurs in southern rice areas on dispersive clay soils. Serious damage is generally confined to aerially sown and dry-broadcast crops. Identification: Red to red–brown segmented worms growing to 75 mm length with relatively
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Fig. 9.1. Aquatic earthworms (Eukerria saltensis). Length 60 mm. (M. Stevens)
thin bodies (about 2 mm in mature specimens) (Fig. 9.1). May be confused with: Bloodworms (Fig. 9.15); however, bloodworms do not exceed 18 mm in length and have a distinct head capsule. Host range: Earthworms ingest soil and organic matter in suitable habitats. Life cycle in temperate rice crops: Eggs are laid within protective cocoons (Fig. 9.2) in the soil and hatching is regulated by soil moisture. Young earthworms grow progressively until reproductive maturity. Risk period: 0–100 days after sowing. The main risk period is shortly after sowing before the shoots emerge above the water surface. Damage: Aquatic earthworms do not feed on healthy rice plants. When dense populations
Fig. 9.2. Cocoons of aquatic earthworms, Eukerria saltensis, 2 mm long. (G. Warren)
Fig. 9.3. Tunnelling activity and castings of aquatic earthworms (Eukerria saltensis) lead to a fine silty layer on the soil surface that prevents root anchoring and increases water turbidity. (M. Stevens)
are present, the tunnelling activity of aquatic earthworms reduces soil compaction, preventing the plants from anchoring effectively (Figs 9.3–9.5). On dispersive soils, turbidity increases, lowering light penetration and reducing photosynthesis. High turbidity also slows plant growth by lowering water temperature. Earthworm tunnelling increases nitrogen and phosphorus in the water column, which can lead to algal blooms. Dense earthworm infestations often attract large flocks of ibis, which cause secondary damage by trampling the young seedlings. Monitoring: Deep water areas generally support the highest densities of earthworms. The bases of irrigation banks also often support high densities because of the moisture retained in the banks during winter.
Fig. 9.4. Aquatic earthworm castings on the surface of a recently drained rice field. Under flooded conditions these disperse to increase water turbidity. (M. Stevens)
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Fig. 9.6. Shells of common water snails from Australian temperate rice crops. Left to right: Austropeplea lessoni, Isidorella newcombi, Glyptophysa sp. A. lessoni is not regarded as a pest species. Shell height for A. lessoni 12 mm. (M. Stevens) Fig. 9.5. Poor plant establishment in a mature rice crop caused by aquatic earthworm (Eukerria saltenesis) damage during seedling establishment. (M. Stevens)
Action level: None established, as variability in soil type has a major effect on damage levels associated with a given population density. Action should be based on seedling health and numbers. Chemical control: None available. Cultural control: Crop rotations are important in the management of aquatic earthworms. Cultivation of non-irrigated winter crops prior to rice helps to reduce earthworm populations, while growing irrigated clover pastures immediately prior to rice cultivation should be avoided in paddocks supporting earthworm infestations. Drill- or sod-sowing also eliminates most earthworm damage. Uniform shallow water across infested fields assists rapid crop establishment, while gas-operated scare-guns can be used to deter ibis.
Water snails Gastropoda: Planorbidae
commonly occur in temperate rice but are not regarded as crop pests. Identification: Small to medium-sized water snails (shell height up to 14 mm) lacking an operculum, predominantly mid to dark brown in colour, shell helicoid, sinistral (aperture on left when shell held upright with aperture facing observer) (Figs 9.6–9.8). May be confused with: Several lymnaeid snail species that are not recognised pests. Common rice field lymnaeids are generally paler in colour and have dextrally coiled shells. Host range: In addition to rice, Isidorella and Glyptophysa feed on a range of aquatic plants, including both macrophytes and algae. Life cycle in temperate rice crops: Egg masses are attached to plant stems or plant debris (Fig. 9.9), and hatch in 3–7 days. The young snails feed and grow continuously until they reach sexual maturity. When the fields are drained prior to harvest, mature snails burrow into the soil and enter dormancy until the fields are inundated again. Survival between consecutive rice crops can approach 50%.
Isidorella newcombi (Adams & Angas) sens. lat. and Glyptophysa sp. Distribution: Isidorella: all states and mainland territories except for the northern tropics, and possibly Tas. (Walker 1988). I. newcombi: NSW, SA (central), Vic. (northern) (Smith and Kershaw 1979). Glyptophysa: all states and mainland territories (Walker 1988). Pest status in temperate rice: I. newcombi: major, widespread, regular. Glyptophysa sp.: minor, widespread, irregular. Other snail species
Fig. 9.7. Isidorella newcombi. Shell height 14 mm. (M. Stevens)
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Fig. 9.8. Glyptophysa sp. Shell height 12 mm. (M. Stevens)
Risk period: 0–100 days after sowing. Damage: Snails feed on the root tissue at the base of the plant, sometimes detaching the plant from the soil. Plants surviving early damage often show reduced tillering and delayed maturity as a consequence of root pruning (Fig. 9.10). Monitoring: Snails are often distributed in patches across fields (Fig. 9.11), with higher densities adjacent to irrigation stops and in areas of deeper water. Monitoring should be conducted along transects through each bay to obtain an accurate estimate of snail densities. Action level: During the crop establishment period chemical control should be considered if there are obvious signs of plant damage, there are more than five large or 10 small snails per square metre, and the density of healthy seedlings is at risk of falling below a target level of 150 plants per square metre (Stevens et al. 2004).
Fig. 9.10. Rice plants damaged by water snails have smaller root systems and tillering is reduced or delayed. An undamaged plant of the same age is on the left. (M. Stevens)
against snail eggs necessitating a second application 7–14 days after the initial treatment. Cultural control: Most serious snail infestations arise from snails emerging from soil dormancy. Whilst I. newcombi can survive between consecutive rice crops, survival is limited over longer periods, and a crop rotation that involves
Chemical control: Control with molluscicides is effective; however, some materials are inactive
Fig. 9.9. Isidorella newcombi egg mass. Length 7 mm. (G. Warren)
Fig. 9.11. Isidorella newcombi often congregate near irrigation stops, particularly when water levels are lowered. (G. Warren)
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breaking the cycle of consecutive summer rice crops is the most effective way to avoid damaging snail infestations.
Tadpole shrimp Triops australiensis australiensis (Spencer & Hall) Notostraca: Triopsidae Tadpole shrimps are often found in temporary aquatic habitats in southern Australia, and are sometimes found in rice crops. Other tadpole shrimp species have been recognised as rice pests in other countries, particularly the USA (California). Distribution: Drier parts of all mainland states and territories. Absent from the northern tropics and wetter areas of southern WA, south-east Qld, eastern NSW and much of Vic. Not known from Tas. (Williams 1980). Pest status in temperate rice: Minor, restricted, irregular. Damage is usually confined to aerially sown and dry-broadcast crops. Identification: Tadpole shrimps commonly grow to 5 cm in length (including tail), but those found in rice crops during the establishment phase of crop growth generally do not exceed 25 mm in length. They are characterised by a brownish, oval body and a segmented tail with two terminal bristles (Fig. 9.12). May be confused with: Unlikely to be confused with other rice invertebrates. Host range: Although an occasional pest of rice crops, tadpole shrimps are generally regarded as detritivores (Gooderham and Tsyrlin 2002).
Fig. 9.13. Cast skin of a tadpole shrimp, Triops australiensis australiensis. Body length (excluding tail bristles) 25 mm. (M. Stevens)
Life cycle in temperate rice crops: Tadpole shrimp eggs are highly resistant to desiccation, and hatching is triggered by wetting. Microscopic larvae gradually take on the adult form after a series of moults (Williams 1980). Risk period: 0–65 days after sowing. Damage: Young tadpole shrimps feed on newly sown seed, but larger shrimps will attack both the roots and shoots of young seedlings. Burrowing at the soil surface often leads to discolouration of the water. Monitoring: Plant damage can be similar to that caused by bloodworms. Tadpole shrimp are often detected in rice fields from the accumulation of cast skins at the downwind end of bays (Fig. 9.13). Action level: None established. Overseas guidelines are generally based on critical seedling densities, rather than pest numbers. Chemical control: Although tadpole shrimps are known to be susceptible to some insecticides, there are no compounds currently registered for their control in Australia.
Fig. 9.12. Tadpole shrimp (Triops australiensis australiensis), dorsal view. Body length (excluding tail bristles) 25 mm. (G. Warren)
Cultural control: None available. The long persistence of viable eggs in dry soil prevents short- to medium-term crop rotations from significantly impacting on tadpole shrimp populations. 285
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Host-plant resistance: None known. Conservation of natural enemies: Rapid emergence of tadpole shrimps from dormant eggs after fields are flooded suggests invertebrate predator populations will probably increase too slowly to exert effective control.
Yabby Cherax destructor Clark Malacostraca: Parastacidae Distribution: The most widely distributed species of crayfish in Australia. Distributed from cool wet regions of Vic. to arid regions of central Australia (Merrick 1993). Pest status in temperate rice: Minor, widespread, regular. Identification: Large crayfish up to 280 mm, predominantly dark brown, dorsal area of abdomen lacking spines (Fig. 9.14). May be confused with: Unlikely to be confused with other freshwater crayfish in rice-producing areas. Host range: Yabbies are opportunistic omnivores but primarily feed on detritus and carrion. Unlike some overseas species, they do not feed directly on rice plants. Life cycle in temperate rice crops: Fertilised eggs are attached to the female pleopods (abdominal swimming appendages). After hatching the young go through three larval stages, with only the third larval stage actively feeding. The larvae remain attached to the pleopods until the final larval stage, and then moult to become independent juveniles.
Juveniles continue to moult at regular intervals as they mature. Yabbies construct burrows in the soil, and when water levels drop they can seal the burrow entrances, allowing them to survive long periods without inundation. Risk period: Complete cropping cycle, from flooding to draining. Damage: Yabbies dig burrows into field and channel banks. These burrows may breach the banks causing significant water loss, and cause banks to erode and collapse. Monitoring: Yabbies are always present in irrigation channels and in field banks more than 1 year old. Damage is most likely where banks are relatively thin and have not been re-formed for 3 or more years. Action level: Not established. Chemical control: None available. Cultural control: The most effective method for reducing yabby numbers is to cultivate and reform irrigation banks after every two rice crops. Thorough cultivation will effectively destroy yabbies within their burrows.
Bloodworms Diptera: Chironomidae Chironomus tepperi Skuse, and other chironomid species Rice fields support a wide range of chironomid species, and while C. tepperi is known to be the principal cause of crop damage during the first 2–3 weeks after sowing, other species have been implicated in damage occurring during the later part of the establishment period. Distribution: C. tepperi: ACT, NSW, Qld, SA, Vic. and WA (Freeman 1961). Pest status in temperate rice: Major, widespread, regular. Bloodworm damage is usually confined to aerially-sown and drybroadcast crops.
Fig. 9.14. A mature yabby, Cherax destructor. Body length 120 mm. (M. Stevens)
Identification: Bloodworms are the larval stage of chironomid midges. The larvae are aquatic, grow to approximately 18 mm length, and are often red in colour (Fig. 9.15). The adult midges are up to 5 mm in length and resemble mosquitoes (Fig. 9.16), but they do not bite.
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Fig. 9.15. Final-instar larva and pupa of the rice bloodworm (Chironomus tepperi). Larval length 16 mm. (G. Warren)
May be confused with: Bloodworms may look similar to aquatic oligochaetes (Fig. 9.1), but bloodworms have a distinct head capsule and, in the case of Chironomus, short, tentacle-like posterior appendages. They are also much shorter than oligochaetes such as Eukerria saltensis, which may be up to 75 mm in length. a
Fig. 9.17. Egg masses of different bloodworm species. A rice bloodworm, Chironomus tepperi egg mass is on the right, 16 mm long. (G. Warren)
Host range: None of the bloodworm species found in rice crops are obligate rice feeders. Most will also feed on decaying organic matter and detritus, while others are filter-feeders or facultative predators. Life cycle in temperate rice crops: C. tepperi generally has only one generation in rice, but other species have multiple asynchronous generations. Female midges lay eggs into the water after the fields are flooded. Gelatinous masses of up to 350 eggs are laid, often in spirals and attached to twigs or other organic matter (Fig. 9.17). Eggs hatch in 1–3 days and the larvae have four instars, of which the third and fourth are most commonly noticed. The short pupal stage does not feed. Mature pupae rise to the surface where the adult midges emerge. Risk period: 0–45 days after sowing.
b
Fig. 9.16. (a) Adult female of the rice bloodworm (Chironomus tepperi), body length 5 mm (G. Warren) . (b) Adult male of the rice bloodworm (Chironomus tepperi), body length 4 mm (G. Warren) .
Damage: Drill-sown rice is less vulnerable to bloodworm damage than aerially sown rice. Larval feeding causes damage to the seed and primary root systems of young plants (Fig. 9.18). Damage is rarely significant after the secondary root system starts to develop. Monitoring: Aerially sown crops should receive a prophylactic treatment on the day of sowing, while dry-broadcast crops should be treated within 3 days of permanent water being applied. Crops should be monitored regularly for 45 days after sowing or application of permanent water to determine if a second treatment is required. Action level: Due to the small size of earlyinstar larvae and difficulties associated with 287
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a
Conservation of natural enemies: C. tepperi colonises rice fields within 3 days of flooding, and there are few effective predators present in the fields at this time.
Rice leafminer Hydrellia michelae Bock Diptera: Ephydridae
b
Fig. 9.18. (a) Damage to seedlings caused by rice bloodworm (Chironomus tepperi). An undamaged rice plant of equivalent age is on the right (M. Stevens) . (b) Rice seed damaged by larvae of the rice bloodworm, Chironomus tepperi (G. Warren) .
identifying bloodworms to species level, quantitative action thresholds have not been developed. An initial prophylactic treatment should be applied routinely to aerially sown and dry-broadcast crops, while any subsequent treatment (or any initial treatment of drill or sod-sown crops) should only be applied in response to substantial bloodworm numbers and clear evidence of plant root damage.
Extensive surveys have not been conducted and other Hydrellia species may also be involved. H. griseola (Fallén) was reported damaging rice in Mareeba (Qld) by Halfpapp (1989), however this record requires confirmation. Bock (1990) described a new Hydrellia species, H. mareeba, which was bred from rice leaves collected in the area, and did not recognise H. griseola as being present in Australia. Distribution: H. michelae: NSW, NT, SA, Vic. and WA (Bock 1990; Stevens 1997). H. mareeba is only known from Qld. Other Hydrellia species (particularly H. griseola and H. philippina Ferino) are major pests of rice in many countries. Pest status in temperate rice: Major, restricted, irregular. Identification: Larvae, when removed from mines, are typically maggot-like, up to 4 mm long and with black mouthparts (Fig. 9.19). The adult leafminer is a metallic green fly approximately 2.5 mm in length (Fig. 9.20). a
Chemical control: Seed dressing with insecticides and controlling damaging populations with liquid insecticide formulations are the main methods of protecting crops against damaging populations of bloodworms. Cultural control: Drill- or sod-sowing will prevent most bloodworm damage, but may compromise weed control programs. It has been alleged that maintaining high organic matter levels in rice fields provides an alternative food source for bloodworms and may reduce crop damage, but this has not been scientifically investigated. Host-plant resistance: No Australian commercial rice varieties have any appreciable level of resistance to bloodworm attack.
b
Fig. 9.19. (a) A leafminer, Hydrellia michelae, larva within a mined rice leaf (G. Warren) , and (b) leafminer larva 4 mm long (G. Warren) .
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a
a
b
b
Fig. 9.20. (a) Leafminer pupae removed from rice leaves, 5 mm long (G. Warren) , and (b) adult leafminer, Hydrellia michelae, length 2.5 mm (G. Warren) .
May be confused with: Unlikely to be confused with other Australian rice insects. Host range: Hydrellia species feed on many aquatic and terrestrial plants, including other graminaceous crops. H. michelae and H. mareeba are only known from rice; however, other hosts presumably exist for H. michelae, which is distributed throughout areas where there is no commercial rice cultivation. Life cycle in temperate rice crops: The female fly lays eggs singly on the leaves of young rice plants. The larvae mine into the leaf blade, extending the mine as they feed. Although not strictly aquatic, larvae can tolerate periods of full submersion. Larvae pupate within the leaf. Risk period: Young plants after permanent water has been applied. Damage increases in response to cool weather and slow plant growth. Damage: Larval mines weaken and kill the leaf blades, and may reduce tillering and delay maturation. Thinning of the crop around the field edges is often an indication of leafminer infestation (Fig. 9.21). Monitoring: Search for early leaf damage and for dead leaves lying flat on the water surface (Fig. 9.21). Larvae and pupae can be detected as distinct ‘bumps’ within the leaf blades (Fig. 9.19). The incidence of plants with leafminers is counted along two diagonal transects across
Fig. 9.21. (a) Dead and dying leaves on the water surface beneath rice plants damaged by leafminers (M. Stevens) , and (b) rice crop showing poor plant establishment along the field margins caused by a heavy leafminer infestation (M. Stevens) .
each rice bay, with at least eight points being sampled along each transect. No more than two points per transect should be within 8 m of the crop edge, and none should be within 3 m. Action level: Action level is dependent on seedling density: < 80 plants per square metre, 5% of plants infested; 80–120 plants per square metre, 10% of plants infested; > 120 plants per square metre, 20% of plants infested (Stevens et al. 2004). Chemical control: Aerial applications of registered insecticides provide adequate control. Newer systemic insecticides have considerable potential for leafminer control, but have not yet been developed for this use pattern in Australia. Cultural control: The risk of severe damage can be reduced by avoiding deep water during crop establishment. Conservation of natural enemies: Larvae are known to be parasitised by several wasp species, however they are not thought to exert significant control. 289
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V E G E TAT I V E G R OW T H TO H A R V E S T
Locusts and grasshoppers Including:
Orthoptera: Acrididae Australian plague locust, Chortoicetes terminifera (Walker) Small plague grasshopper, Austroicetes cruciata (Saussure) N (Main entry in Chapter 18 Locusts and grasshoppers of
water levels. Colonisation of fields can be prevented by ensuring that plant roots are always fully covered with water. Fully grown aphids are 1.2–2.2 mm long and dark green to grey–brown in colour, while nymphs are lighter coloured with a reddish area on the tip of the abdomen (Fig. 9.22).
Green vegetable bug Nezara viridula (Linnaeus) Hemiptera: Pentatomidae
pastures and rangelands.)
N (Main entry in Chapter 7 Pulses—summer.)
Locusts and grasshoppers have previously been reported as rice pests; however, recent evidence suggests that crop losses are rarely significant. The majority of confirmed damage has occurred during establishment of drill-sown crops when fields have dried out between flushes.
The green vegetable bug has been reported as a significant rice pest in Japan, but while present in temperate Australian rice crops it has yet to cause significant damage. Feeding on maturing grain has the potential to reduce quality by increasing the proportion of ‘pecky’ (blotched) grain. The risk period is between early February and harvest. Green vegetable bugs are most prevalent at the edge of fields, especially when rice is grown adjacent to other host crops. Control is not presently recommended.
Rice root aphid Rhopalosiphum rufiabdominalis (Sasaki) Hemiptera: Aphididae Rice root aphids have been found feeding on rice plants in temperate crops; however, they have not caused significant crop damage. They suck fluids from the plant roots, but can only do so when the bases of the plants are exposed by low
Rice stink bug Eysarcoris trimaculatus (Distant) Hemiptera: Pentatomidae N (Main entry in Chapter 7 Pulses—summer.)
Although present in temperate Australian rice areas, rice stink bugs have yet to occur in sufficient densities to justify control. This is thought to be related to the low winter temperatures and low summer humidity levels in southern NSW. Control is not presently recommended.
Common armyworm Leucania convecta (Walker) Lepidoptera: Noctuidae N (Main entry in Chapter 2 Cereals.) Fig. 9.22. Wingless adult and nymph of the rice root aphid, Rhopalosiphum rufiabdominalis. Length of adult 1.5 mm. The nymph has a red area (arrowed) at the tip of the abdomen. (G. Warren)
Pest status on rice: Major, may be restricted and irregular, occasionally in plague numbers. Identification and biology: See Chapter 2.
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a
b
c Fig. 9.23. Damage to rice foliage caused by common armyworm, Leucania convecta. (K. Bechaz)
Risk period: Generally from flowering to harvest, but earlier damage is possible. Damage: Larval feeding defoliates plants and often results in stems being severed below the panicles (Fig. 9.23). Monitoring: Early detection is important, and the incidence of damage and insects should be sampled along two diagonal transects per bay. At least eight points should be sampled per transect, with no more than two sampling points per transect within 8 m of the crop edge and none within 3 m.
Fig. 9.24. (a) Sugarcane and maize stemborer, Bathytricha truncata, larva removed from rice stem, length 20 mm (M. Stevens) , (b) adult moth, 15 mm long (NSWDPI) , and (c) whitehead caused by sugarcane and maize stemborer attack after panicle formation (M. Stevens) .
Action level: When the panicles are not yet exposed, treat when 35% or more of plants have at least 25% defoliation. When panicles are exposed and there is more than 2 weeks to harvest, treat when there are one or more caterpillars per plant. If less than 2 weeks to harvest, treat only if there are two or more caterpillars per plant (Stevens et al. 2004). A less conservative protocol for drill-sown crops has been proposed by Elder et al. (1992), who suggest treatment is required when infestation levels exceed four larvae per metre of row.
As the common name suggests, this species also occurs on other graminaceous crops. Female moths lay eggs on the leaf sheath of rice plants and the newly hatched larvae graze the surface of the leaf before boring into the stem, which they eventually hollow out as they develop, producing a ‘dead heart’. If stemborers attack after the panicle has formed they produce ‘white heads,’ dead panicles containing empty or only partly filled grains (Fig. 9.24). Pupae overwinter within rice stubble.
Control: Insecticide sprays should be applied before larvae grow to full size (about 40 mm). Older larvae are harder to kill and may have already caused significant damage.
Hairy rice caterpillar
Sugarcane and maize stemborer Bathytricha truncata (Walker) Lepidoptera: Noctuidae
Celama taeniata (Snellen) Lepidoptera: Noctuidae Caterpillars of this native moth have been reported feeding on rice panicles and developing grain in the Murray Valley. Larvae grow to approximately 7 mm in length, pupate in the rice stem and may overwinter in rice
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a
b
Queensland. Damage is largely consistent with that in temperate rice, however bloodworms are considered less significant in Qld and WA than they are in temperate crops. This may reflect differences in the species present, or may reflect the strong reliance on aerial sowing in temperate rice.
Rice leafminer Hydrellia mareeba Bock Diptera: Ephydridae N (Main entry in this chapter, page 288.)
Fig. 9.25. (a) Hairy rice caterpillar, adult moth, (Celama taeniata) and pupal case, length of adult 6 mm (G. Warren) , and (b) hairy rice caterpillar alongside rice grain, length of caterpillar 5 mm (G. Warren) .
stubble. The moth that emerges is 6 mm long with a wingspan of 12 mm. The wings are brown with white markings (Fig. 9.25).
Tropical rice Rice is not currently being produced commercially in northern Australia, although it was previously grown in the NT (Humpty Doo area), WA (Ord River Irrigation area) and, until relatively recently, the Mareeba–Ayr–Ingham areas of north Qld. Many tropical pest species attack the crop in these areas, and the pest fauna shows strong similarities to that found in SouthEast Asia. The major pests of rice in northern Australia are discussed briefly below, with much of the information drawn from Elder et al. (1992) and Pathak and Khan (1994).
E S TA B L I S H M E N T
Bloodworms Chironomus sp. Diptera: Chironomidae N (Main entry in this chapter, page 286.)
Little is known about the composition of chironomid communities associated with rice crop injury in Western Australia and north
Considerable uncertainty exists in regard to records of H. griseola (Fallén) attacking rice in Qld. Damage is identical to that caused by H. michelae in temperate rice. Two species of parasitoids are recorded from rice leafminer larvae in Qld, Dolichogenidea sp. and an unidentified species of Alysiinae (Hymenoptera: Braconidae) (Halfpapp 1989).
V E G E TAT I V E G R OW T H TO H A R V E S T
Rice leafhopper Nephotettix malayanus Ishihara & Kawase Hemiptera: Cicadellidae Rice leafhoppers are minor pests in Qld. Nephottettix species are known vectors of rice diseases that occur in South-East Asia (yellow dwarf, tungro disease, and others), but there are no records of N. malayanus vectoring rice diseases in Australia.
Brown planthopper Nilaparavata lugens (Stål) Hemiptera: Delphacidae The brown planthopper is widely distributed throughout eastern, southern, and South-East Asia and is also found in northern Australia. Sap-feeding by nymphs and adults causes the lower leaves to yellow, producing characteristic ‘hopper burn’ (Elder et al. 1992). Brown planthoppers act as vectors for rice diseases (grassy stunt, ragged stunt) over much of their range (Pathak and Khan 1994). Cultural and biological control are important for brown planthopper management. Ratoon crops and
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Aquatic predators in rice fields Flooded rice fields provide a habitat for many species of aquatic predators. These include damselfly (Fig. 9.26) and dragonfly nymphs (Odonata) (Fig. 9.27), dytiscid (Fig. 9.28)
Fig. 9.26. Damselfly nymph, Odonata: Coenagrionidae. Body length 14 mm. (G. Warren)
Fig. 9.28. Dytiscid water beetle larva, 20 mm long. (G. Warren)
Many of the pesticides used for rice pest and weed control kill predatory aquatic insects, and care should be taken to minimise their use, particularly after the early crop establishment period.
and hydrophilid beetles, and a range of predatory Hemiptera, including water boatmen (Notonectidae) (Fig. 9.29), water scorpions and needle bugs (Nepidae) (Fig. 9.30), and giant water bugs (Belostomatidae). These predators colonise fields relatively slowly, and while they appear to have little impact on early establishment pests such as Chironomus tepperi, there is strong evidence to indicate they play an important role in rice field ecosystems, particularly in control of mosquito larvae.
Fig. 9.27. Dragonfly nymph, Hemianax papuensis. Length 24 mm. (G. Warren)
Fig. 9.29. A notonectid, or backswimmer, ventral view. Length 7 mm. (G. Warren)
Fig. 9.30. Water scorpion, Laccotrephes tristis Stål. Body length (excluding siphon protruding from back of body) 28 mm. (M. Stevens)
stubble regrowth should be avoided and stubble destroyed as early as possible. Excessive use of pesticides for the control of planthoppers and other pests will damage populations of beneficial species and result in the resurgence of planthopper populations. Resistant rice varieties are available, however their performance depends on the selection of varieties with resistance to particular planthopper biotypes.
Paddy bug Leptocorisa acuta (Thunberg) Hemiptera: Alydidae Several genera of Alydidae damage rice in Asia, Africa and South America; however, Leptocorisa is the most important genus worldwide. L. acuta
is found in Asia and northern Australia (Pathak and Khan 1994). Crop damage is similar to that caused by rice stink bugs, with both nymphs and adults feeding on the developing grains, reducing yield and quality. Damaged grains show brown spots at the points where the feeding punctures were made (Halfpapp 1982). Adult L. acuta are approximately 14 mm long, 3 mm wide, pale brown in colour (Fig. 7.34), and give off an oily smell when disturbed. They are a significant pest of rice in north Qld, particularly in coastal areas (Elder et al. 1992).
Rice stink bug Eysarcoris trimaculatus (Distant) Hemiptera: Pentatomidae 293
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a
Caseworms are recorded as pests of rice in east and South-East Asia, Africa and South America. In Australia they are minor pests of tropical rice. Larvae construct cases of cut leaf fragments and damage growing rice leaves by their feeding. The short-lived adult moths are white with pale brown to black wing markings. A more detailed description of biology and crop damage is given by Pathak and Khan (1994).
Rice leaffolder b
Cnaphalocrocis medinalis (Guenée) Lepidoptera: Pyralidae The rice leaffolder is a major pest throughout South-East Asia. From the second instar onwards the larvae fold the leaves over to form a shelter using secreted threads that contract as they dry. Larvae feed by scraping away the leaf mesophyll within the shelters.
White rice stemborer Scirpophaga innotata (Walker) (= Tryporyza innotata) Fig. 9.31. (a) Eggs and first-instar nymph of the rice stink bug, Eysarcoris trimaculatus, length of nymph 1 mm (G. Warren) , and (b) adult rice stink bug, 6 mm long (G. Warren) .
Although only rated as a potential pest of temperate rice, the rice stink bug is a major pest in Qld rice (Elder et al. 1992), and also significant in WA (Learmonth 1980). Eggs are laid in groups on the surface of rice leaves (Fig. 9.31). Small nymphs that emerge move into panicles and use their piercing mouthparts to feed on developing rice grains. This leads to grain loss or grains with dark blemishes, known as ‘pecky’ rice. Chemical control is effective, however parasitism by tachinid flies has been recorded (Kay 2002), and egg parasitism by Hymenoptera is also believed to play a significant role in regulating stink bug populations.
Caseworm Nymphula depunctalis (Guenée) Lepidoptera: Pyralidae
Darkheaded riceborer Chilo polychrysa (Meyrick)
Striped riceborer Chilo suppressalis (Walker) Lepidoptera: Pyralidae The white rice stemborer is found throughout eastern Asia, and is the most serious insect pest affecting tropical rice production in the NT and WA. It is also regarded as a major pest in rice in Qld. The darkheaded riceborer and striped riceborer are minor pests of rice in the NT (Li 1990). The larvae of all species cause similar damage to the native sugarcane and maize stemborer: developing larvae hollow out stems producing ‘dead hearts’, however if stemborers attack after the panicle has formed they produce ‘white heads,’ dead panicles containing unfilled grains. Egg and larval parasitoids play an important role in suppressing stemborer populations and integrated control programs
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need to be focussed on the conservation of natural enemies (Learmonth 1980, 1981b).
Armyworms Lepidoptera: Noctuidae Common armyworm, Leucania convecta (Walker) N (Main entry in this chapter, page 290.)
Northern armyworm, Leucania separata (Walker) Dayfeeding armyworm, Spodoptera exempta (Walker) Lawn armyworm, Spodoptera mauritia (Boisduval)
All four species of armyworm attack tropical rice and can occur as plagues. Common, dayfeeding and lawn armyworms are major pests of rice in Qld (Elder et al. 1992), while northern armyworms are serious pests in WA (Learmonth 1980, 1981a). General biology and damage is similar for all species.
Rice skipper Parnara sp. Lepidoptera: Hesperiidae Larval rice skippers are serious rice pests in north Qld. The larvae form tube-like shelters from rice leaves and feed on the leaf blade. Severe defoliation can occur. Chemical control is necessary when more than 15% of leaves are severely damaged (Elder et al. 1992).
Sources of information Bock, I.R. (1990). The Australian species of Hydrellia Robineau-Desvoidy (Diptera: Ephydridae). Invertebrate Taxonomy 3(7): 965–993. Elder, R.J., Brough, E.J. and Beavis, H.S. (1992). Managing Insects and Mites in Field Crops, Forage Crops and Pastures. Department of Primary Industries, Queensland, Brisbane. 143pp. Freeman, P. (1961). The Chironomidae (Diptera) of Australia. Australian Journal of Zoology 9: 611– 737. Gooderham, J. and Tsyrlin, E. (2002). The Waterbug Book: A Guide to the Freshwater Macroinvertebrates of Temperate Australia. CSIRO Publishing, Collingwood. 232 pp. Halfpapp, K.H. (1982). Insect pests of rice. Queensland Agricultural Journal 108(5): xxix–xxx. Halfpapp, K.H. (1989). Rice leaf miner Hydrellia griseola in Australia. International Rice Research Newsletter 14(6): 32. Kay, I.R. (2002). Parasitism of Eysarcoris trimaculatus (Distant) (Hemiptera: Pentatomidae) by two tachinid flies (Diptera: Tachinidae). Australian Entomologist 29(1): 21–24. Learmonth, S.E. (1980). Rice insect pests and their management in the Ord River irrigation area of Western Australia. International Rice Research Newsletter 5(1): 14. Learmonth, S.E. (1981a). Exotic parasite helps control Ord armyworms. Journal of Agriculture – Western Australia 22(2): 59. Learmonth, S.E. (1981b). Parasite ‘hitch hikers’ hit Ord rice pest. Journal of Agriculture – Western Australia 22(2): 68–69. Li, C.S. (1990). Status and control of Chilo spp, their distribution, host range and economic importance in Oceania. Insect Science and its Application 11: 535–539. Merrick, J.R. (1993). Freshwater Crayfishes of New South Wales. Linnean Society of New South Wales, Sydney. 127pp. Pathak, M.D. and Khan, Z.R. (1994). Insect Pests of Rice. IRRI/ICIPE, International Rice Research Institute, Manila. 89pp.
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Smith, B.J. and Kershaw, R.C. (1979). Field Guide to the Non-marine Molluscs of South Eastern Australia. Australian National University Press, Canberra. 285pp. Stevens, M.M. (1997). Common Invertebrates of New South Wales Rice Fields. Biology, Pest Status and Control. NSW Agriculture, Yanco. 55pp. Stevens, M.M., Brickhill, J. and Brown, P. (2004). Pests of rice crops – vertebrates and invertebrates. Ch. 10 (28 pp.) in: Kealey, L. (ed.) Production of Quality Rice in South Eastern Australia. Rural Industries Research and Development Corporation, Canberra. Walker, J.C. (1988). Classification of Australian buliniform planorbids (Mollusca: Pulmonata). Records of the Australian Museum 40(2): 61–89. Williams, W.D. (1980). Australian Freshwater Life. The Invertebrates of Australian Inland Waters. 2nd ed., Macmillan Australia, Melbourne. 321pp.
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10 SORGHUM N Sorghum bicolor (Poaceae). Origin: north-east Africa
B.A. Franzmann
(a) Sorghum crop, Hay, NSW (SARDI: P.T. Bailey)
(b) Red seed (DPI&F Qld: B. Franzmann)
sorghum seeding germination growth head emergence flowering green seed red seed & harvest
Jul
Aug Sep
Oct
Nov Dec
Jan Feb Mar
Apr May Jun
Phenological stages of a sorghum crop.
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PEST (major pests in bold)
PAGE
Germination black field earwig false wireworms cutworms Growth corn aphid northern armyworm common armyworm dayfeeding armyworm Flowering, head and seed development corn earworm Rutherglen bug sorghum midge sorghum head caterpillar yellow peach moth
298 298 299 299 300 300 300 300 301 301 302 303
Grain sorghum is the most important summer cereal in Australia with an average area of 691 000 ha. The grain is used mainly for feeding stock. Approximately 65% of the crop is grown in central and southern Qld, 34% in NSW and the remainder in NT and the north-east of WA. Most is grown as a dryland crop except in southern NSW, where the crop is mostly irrigated.
False wireworms
In southern Qld and northern NSW the most common planting time is October, and in central Qld in January. However, sorghum is planted whenever planting rains arrive between October–January in southern areas and December–March in central Qld.
N (Main entry in Chapter 5 Oilseeds.)
Coleoptera: Tenebrionidae Southern false wireworm, Gonocephalum macleayi (Blackburn) Large false wireworm, Pterohelaeus alternatus Pascoe Pest status on sorghum: Major, widespread, irregular. Identification. See Chapter 5. May be confused with: True wireworms (Elateridae). Host range: All field crops and many weeds.
G E R M I N AT I O N
Black field earwig Nala lividipes (Dufor) Dermaptera: Labiduridae N (Main entry in Chapter 4 Maize.)
Pest status on sorghum: Minor, widespread, regular. Damage: Nymphs and adults feed on germinating seed, but cause most damage when they attack the roots of young plants and the prop roots of older plants. Monitoring and control: See Chapter 4.
Life cycle on sorghum: Adults emerge from the soil during spring and early summer, and larvae are found from about 2 months later until the next spring. Larvae remain in the vicinity of germinating seeds while the top layer of soil remains moist, but follow moisture below the root zone as the soil dries. Risk period: The risk from larvae is highest for early planted crops, whereas the risk from adults is highest in summer. Damage: Larvae feed on newly germinated seeds and the growing points of plants. Adults damage young plants by surface-feeding or cutting off the plant at, or near, ground level.
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Damage is severe when germination and early plant growth are retarded by dry conditions, particularly if seeds are sown into a dry seed bed, or when the weather is cool and wet. Monitoring: Germinating seed baits detect both adults and larvae. During periods when wireworms are active in the crop, they may be found near the most-recently injured plants or approaching adjacent undamaged ones. Action level: Treatment decisions depend on the numbers of insects found at germinating seed baits. Control larvae if more than 25 larvae are found around 20 germinating seed baits, and adults if more than 60 are found around 20 baits. Chemical control: Chemical controls are costeffective and must be applied at planting. Cultural control: Prepare the ground so that germination is as even and rapid as possible. The use at planting of press wheels set at 2 kg cm–1 width after planting rain and up to 4 kg in dry soil, reduces damage. Rapid germination and emergence is promoted by shallow sowing into a warm, moist seed bed. Clean cultivation during summer dries out the topsoil and eliminates weeds that provide food for the adults. Conservation of natural enemies: Natural enemies provide very little control.
Cutworms Agrotis spp. Lepidoptera: Noctuidae N (Main entry in Chapter 2 Cereals.)
Pest status on sorghum: Minor, widespread, irregular. Damage: Cutworm larvae feed on leaves and stems of young plants, which may wilt and die if stems are cut at or below ground level. Cutworms may be in the ground at seeding if the crop is sown into a previously weedy field. Slow plant growth during dry or cold periods increases the risk period. Monitoring and control: Cut plant parts on the ground are indicative of cutworm feeding. Larvae may be detected at evening or night. Spot treatment of affected areas when larvae are active is usually effective. Clean-fallowing prior to planting and weed control around the field perimeter at least 1 month before planting reduces risk of damage.
G ROW T H
Corn aphid Rhophalosiphum maidis (Fitch) Hemiptera: Aphididae Distribution: On all continents and in Australia wherever sorghum is grown. Pest status on sorghum: Minor, widespread, regular. Identification: Up to 2 mm long, light to dark olive-green with a purple area at the base of small tube-like projections at the rear of the body (Fig. 2.9). Adults are generally wingless but may be winged. Nymphs are similar but smaller in size. May be confused with: Although other species of aphids may be present on sorghum, the corn aphid is the most common and numerous. Host range: Sorghum, maize, winter cereals and many grasses. Life cycle on sorghum: Corn aphids breed throughout the summer on sorghum with a life cycle of about 1 week. There can be up to 13 generations on a sorghum crop and 30 generations per year. Risk period: All stages of the crop are attacked, but the most serious damage occurs when high populations infest heads. Damage: Adults and nymphs suck sap and produce honeydew. Very high numbers may turn plants yellow and to appear unthrifty. High populations on heads produce sticky grain and clog harvesters. Water-stressed dryland crops lose yield. Monitoring: Estimate percentage of plants infested and percentage of leaf area covered by aphids. Action level: The action level in the vegetative stage is 100% of plants with 80% of the leaf area covered by aphids. On the heads it is 75% of heads with 50% of the head covered by aphids. Chemical control: Chemical control is costeffective. Host-plant resistance: Hybrids with open heads are less infested than tight-headed hybrids. 299
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a
Pest status on sorghum: Minor, irregular occurrence. The northern armyworm occurs in northern NSW and Qld, the common armyworm is widespread in southern sorghum areas, and the dayfeeding armyworm is recorded as a minor pest of NT and Qld sorghum. Damage: Young plants may be defoliated or killed by larval feeding. Older plants may outgrow damage but seed yield may be reduced. Sorghum crops may be invaded by large larvae moving from adjacent cereal crops.
b
Signs of armyworm damage are chewed leaf margins and faecal pellets around the base of young plants or in the throats of older plants. The northern and common armyworms usually feed during evenings and at night, and during the day they hide on the under surface of the lower leaves, in the throats of mature plants or under plant debris or vegetation on the soil surface. Dayfeeding armyworms are active during the day. Monitoring: Monitor during the early growth stage and again during head emergence and flowering. Egg-laying is often associated with heavy rainfall, so checks for larvae should be made several weeks after rainfall. Action level: Treatment decisions should be made on whether the crop can compensate for damage, the prevalence of natural control agents, the value of the crop and the cost of treatment.
Fig. 10.1. (a) White collared ladybird, Hippodamia variegata Goeze, first discovered in Australia in 2000 (Franzmann 2002), and (b) striped ladybird, Micraspis frenata, include aphids in the larval diet. (K. Power)
Conservation of natural enemies: Wasp parasitoids Lysephlebus testaceipes, ladybirds (Fig. 10.1) and hoverflies are effective natural enemies.
Chemical control: Chemical treatment of common armyworms may be improved by applying late in the day when larvae are feeding.
F LOW E R I N G , H E A D A N D S E E D D E V E LO PM E N T
Corn earworm Armyworms Lepidoptera: Noctuidae Northern armyworm, Leucania separata (Walker) Common armyworm, Leucania convecta (Walker) Dayfeeding armyworm, Spodoptera exempta (Walker)
Helicoverpa armigera (Hübner) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Pest status on sorghum: Major, widespread, regular. Identification: See Chapter 3. May be confused with: Sorghum head caterpillar and yellow peach moth.
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Life cycle on sorghum: Adult moths emerge from overwintering diapause in spring and may complete five or six generations in sorghum crops. On an individual crop, one generation can occur on leaves in the vegetative stage, and another generation on the heads. The life cycle occupies 4–6 weeks. Small caterpillars (less than 10 mm long) feed mainly on pollen sacs in the flower head where they cause little damage, but larger caterpillars mainly feed on the developing seeds. Risk period: Larvae do not cause economic damage before flowering, even though foliage may appear ragged. Eggs laid on heads just prior to flowering from December to March produce larvae that cause economic damage.
Fig. 10.2. Adult Rutherglen bug. (K. Power)
Damage: Corn earworm larvae feed on developing seed. Each larva destroys about 2.4 g of grain.
plants (Fig. 10.2). Seed heads may be damaged by feeding, resulting in reduced yield and quality.
Monitoring: Count larvae on heads, which have just finished flowering by spinning (rotating the head stalks in the palms of the hands) into a bucket. Aim to detect small larvae about 10 mm long.
Monitoring: Monitoring from ‘soft dough’ to ‘hard dough’ stage of seed development , especially during hot, dry periods, should aim at detecting adults and nymphs in the head.
Action level: The level varies with factors such as commodity price and cost of insecticide. The level can be calculated using the factor of 2.4 g of grain destroyed per larva. As a guide, one to two larvae per head may result in economic damage. Biocidal control: A commercially formulated nuclear polyhedrosis virus (NPV) is cost-effective. Cultural control: Plant hybrids with open head type. Host-plant resistance: Hybrids with open heads are less infested than tight-headed hybrid varieties. Conservation of natural enemies: With the use of NPV, natural enemies (mainly parasitic wasps) are conserved.
Rutherglen bug Nysius vinitor Bergroth Hemiptera: Lygaeidae N (Main entry in Chapter 5 Oilseeds.)
Pest status: Minor, widespread, irregular occurrence. Damage to sorghum: Numerous immature and adult bugs may sometimes feed on sorghum
Action level: An indicative action level is 85% of heads containing more than 50 bugs per head sampled from five consecutive heads in six locations in the crop. Chemical control: Patches can be spot-treated. Chemical control may be cost-effective.
Sorghum midge Contarinia sorghicola (Coquillett) Diptera: Cecidomyiidae Distribution: The sorghum midge is on all continents where sorghum is grown. It is recorded from eastern Australia and NT, but not presently in WA. Pest status: Major, widespread, regular occurrence. Identification: A mosquito-like, orange fly, 1.5 to 2 mm long, with very long antennae (Fig. 10.3). Females are larger than males and have a slender ovipositor on the rear of the abdomen. Host range: Grain sorghum, forage sorghum, Johnson grass, Columbus grass, but not Australian native Sorghum spp. 301
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and moisture stress, but sorghum midge damage may be distinguished by the empty white pupal cases protruding from the tips of glumes and emergence holes of parasitoids through the sides of glumes. Monitoring: Count adult midge on flowering heads at about mid-morning. Repeat daily. Action level: The level varies with resistance level of the hybrid and other factors such as commodity price and cost of insecticide. The level can be calculated using the factor of 1.4 g of grain destroyed for each egg-laying adult. On susceptible hybrids the level is usually about one adult per head.
Fig. 10.3. Adult sorghum midge on a sorghum head. (DPI&F Qld: C. Freebairn)
Life cycle on sorghum: Adult midge emerge from overwintering diapause in spring and, after one to two generations on Johnson grass, attack grain sorghum for five or six generations. Eggs are laid into the developing flower spikelets and larvae feed on the developing ovary, preventing normal seed development. The life cycle takes about 3 weeks. Under optimum conditions, eggs hatch within 3 days and larval development is completed within 9–11 days. Adults emerge from the pupae within 3 days and rarely live for more than 1 day. Female sorghum midges may lay up to 120 eggs and up to 20 eggs in an individual flower spikelet. In autumn, most larvae enter diapause for overwintering and may remain in diapause for up to 5 years. After temperatures increase in spring, rainfall triggers the commencement of development with adult midges emerging about 2–3 weeks after rain. Risk period: During flowering from December to March. Damage: Midge larvae destroy developing seed. One larva is enough to prevent seed development. On midge-susceptible hybrids, the progeny of each egg-laying adult on a head destroy 1.4 g of grain. High populations on susceptible hybrids can completely destroy the crop (Franzmann 1993; Franzmann and Butler 1993). Other causes of head damage include heat
Chemical control: Chemical control may be cost-effective but more than one application of insecticide may be needed on susceptible hybrids. Cultural control: Early planting, removal of alternative hosts, cultural practices to ensure even flowering and resistant hybrids. Interstate quarantine: Transport of grain containing diapausing larvae believed to be the main method of spread and movement of sorghum midge into WA is presently restricted. Host-plant resistance: Resistant hybrids are available and widely used. Conservation of natural enemies: With the use of cultural controls (particularly resistant hybrids) insecticide use is rare and natural enemies are conserved. Natural enemies are three small black wasp parasitoids: Eupelmus sp. Eupelmidae, Tetrastichus sp. and Aprostocetus sp. Eulophidae, whose presence may be recognised by their small round emergence holes in the spikelet.
Sorghum head caterpillar Cryptoblabes adoceta Turner Lepidoptera: Pyralidae Distribution: In coastal and tropical areas of north-eastern Australia and NT. Pest status: Minor, restricted, irregular. Identification: Larvae (caterpillars) grow to 13 mm long and are yellow–brown to grey–
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Host-plant resistance: Hybrids with open heads are less infested than tight-headed hybrids. Conservation of natural enemies: Natural enemies (mainly parasitic wasps) provide some biological control.
Yellow peach moth Conogethes punctiferalis (Guenée) Lepidoptera: Pyralidae Distribution: In Australia in coastal and tropical areas and in South-East Asia. Pest status: Minor, restricted, irregular. Fig. 10.4. Sorghum head caterpillars in a sorghum head surrounded by frass. The largest larva is 13 mm long. (DPI&F Qld: D. Ironside)
green. They have a dark line running along each side of the back (Fig. 10.4). May be confused with: Corn earworm and yellow peach moth. Host range: Sorghum, maize, adzuki beans. Life cycle: One generation on a sorghum crop. Risk period: Eggs are laid on heads during flowering from December to March. Larvae eat the grain between ‘soft dough’ stage and harvest. Damage: Larvae feed on developing seed, webbing clusters of seed together. Each larva destroys about 0.5 g of grain. Monitoring: Signs of caterpillars are webbing covering clusters of seeds and the whole head covered in webbing, incorporating small pink or white particles of excreta. Count larvae on heads with seed in the milky stage by spinning (rotating the head stalks in the palms of the hands) into a bucket. Action level: The level varies with factors such as commodity price and cost of insecticide. The level can be calculated using the factor of 0.5 g of grain destroyed per larva. The threshold usually works out to be 5–10 larvae per head. Chemical control: Chemical control is costeffective. Open-headed sorghum varieties allow penetration of insecticides.
Identification: Larvae (caterpillars) grow to 20 mm long and have a dark head and are grey– pink with a dark spot each side of the midline of each body segment (Fig. 4.9). May be confused with: Corn earworm and sorghum head caterpillar. Host range: Sorghum, maize, cotton. Life cycle on commodity: There is only one generation on each sorghum crop. The larval stage occupies about 3 weeks and the pupal stage about 2 weeks. Risk period: Eggs are laid on heads after flowering from December to March. Damage: Larvae feed on developing seed. Each larva destroys about 1 g of grain. Monitoring: Count larvae on heads with seed in the milky stage by spinning (rotating the head stalks in the palms of the hands) into a bucket. Action level: The level varies with factors such as commodity price and cost of insecticide. The level can be calculated using the factor of 1 g of grain destroyed per larva. The threshold usually works out to be two to five larvae per head. Chemical control: Chemical control is costeffective. Host-plant resistance: Hybrids with open heads a less infested than tight-headed hybrids. Conservation of natural enemies: Natural enemies (mainly parasitic wasps) provide some biological control.
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Sources of information Franzmann, B.A. (2002). Hippodamia variegata (Goeze) (Coleoptera: Coccinellidae), a predaceous ladybird new in Australia. Australian Journal of Entomology 41: 375–377. Franzmann, B.A. (1993). Ovipositional antixenosis to Contarinia sorghicola (Coquillett) (Diptera: Cecidomyiidae) in grain sorghum. Journal of the Australian Entomological Society 32: 59–64. Franzmann, B.A. and Butler, D.G. (1993). Compensation for reduction in seed set due to sorghum midge (Contarinia sorghicola) in midge-susceptible and midge-resistant sorghum (Sorghum bicolor). Australian Journal of Experimental Agriculture 33: 193–196.
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11 SUGARCANE M.N. Sallam, P.G. Allsopp, K.J. Chandler and P.R. Samson N Sugarcane, Saccharum spp. hybrids Poaceae (Graminaceae)
A north Qld sugar farm with both newly planted and maturing cane. Surrounding rainforest offers a variety of food sources for adult scarab pests.
planting germination growth flowering (arrowing) harvest
Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec Jan Phenological stages of sugarcane.
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Canegrowing areas (not to scale) Mossman
Cairns Gordonvale Townsville Ayr
Proserpine Mackay
Queensland Rockhampton
Ord
Bundaberg Gin Gin Childers Maryborough Nambour Rocky Point
Brisbane Broadwater
NSW
Harwood
Grafton
The sugarcane season, for Burdekin, central and northern Qld regions, where cane is harvested 12–16 months after of planting. In cooler areas (mainly NSW), the crop grows for 16–24 months before harvesting.
PEST (major pests in bold) Germination symphylans wart-eye mite termites crickets: black field cricket, mole cricket sugarcane wireworms Stenocorynus weevil sugarcane bud moth Ratooning cicadas soldier flies ratoon shootborer Growth sugarcane spider mite
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PEST (major pests in bold) locusts aphids ground pearls pink sugarcane mealybug planthoppers sugarcane froghopper linear bug black beetles canegrubs (19 species) leaf beetles sugarcane butt weevil sugarcane weevil borer whitefringed weevil sugarcane and maize stemborer armyworms and loopers funnel ant
Areas of crop in Australia: The total area of sugarcane harvested in 2002 in Qld and NSW is 435 884 ha (BSES 2003) and 3143 ha for WA in 2001. Origin: New Guinea. Crop cycle: Sugarcane is a perennial crop in tropical and sub-tropical Australia. A ratoon crop is grown in the following year(s) from the root mass and stubble remaining in the soil from the previous crop. Crop cycles are usually restricted to four to five ratoons, but more are possible. Generally, growers start planting in April–May, but autumn-planting in southern Qld is usually earlier (February–April), and then spring-planting usually takes place between August and October. Growth of the first is interrupted by low temperatures, while the second allows plough-out/re-plants or long fallows/break-crops. The crop is harvested when sugar content is at its maximum, between June and December.
G E R M I N AT I O N
Symphylans Hanseniella spp. Symphyla: Scutigerellidae Distribution: The genus Hanseniella is distributed worldwide. In Australia, symphylans are present in all cane districts and on most soil types.
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Pest status: Minor, widespread, irregular. Identification: Symphylans are 5–10 mm long, white or cream in colour, and look like small centipedes (Fig. 11.1). They have two long, slender antennae and 12 pairs of legs when mature. May be confused with: Unlikely to be confused with any other pests of cane. Hosts: Symphylans feed on the roots of a wide range of plants and prefer soils rich in organic matter. Life cycle in sugarcane: Symphylans are present all year round. Females lay groups of 10–20 eggs. Young symphylans look like the adults but have fewer legs. Numbers increase rapidly when rotting vegetable matter, such as a legume cover crop, is ploughed in. Symphylans are most active in loose soil or naturally cracking loams. Risk conditions: Severe damage is mostly found when autumn planting is followed by a cold, wet winter. Damage can last into spring months causing poorly stooled crops with few stalks. Damage: Symphylans actively dig cylindrical pits of 0.5–1.0 mm diameter in growing roots. Root growth stops when the growing point is damaged causing branching, with the side branches often stubbed (Fig. 11.2). Damage results in a small root-ball, and damaged plants wilt easily in dry weather and roots are usually very slow to grow. Damage is common in young 307
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Fig. 11.1. Adult symphylan, 10 mm long. (BSES)
cane, but mature crops can sometimes be attacked. In Qld, damage has occurred from Mossman to Babinda, at Innisfail, Tully, Herbert Valley, and at Mackay and Gin Gin. Monitoring: Symphylans live in soil cracks and can be difficult to find; digging is the only method to monitor populations. Action level: Preventative chemical treatment at planting is recommended in prone areas in anticipation of damage. Chemical control: Insecticides routinely used at cane-planting significantly reduce infestation. Cultural control: Spring plantings are not normally damaged. Rapid germination and root growth seems to outstrip the ability of symphylans to stunt the roots. In hot, dry spring months, damaged plant cane recovers sooner if soil is hilled-up around the shoots. Natural enemies: Unknown.
Fig. 11.3. A node on sugarcane stem showing healthy shoots (top) and a wart eye shoot (arrowed). The stem is about 30 mm diameter. (BSES)
Wart-eye mite Unidentified mite species. Distribution: These mites have been recorded on sugarcane in central and far north Qld. Pest status: Minor pests, restricted to some areas and occurring irregularly. Identification: The mites are 1 mm long and are best identified by damage symptoms. May be confused with: Unlikely to be confused with any other pests of cane. Life cycle in sugarcane: Unknown. Risk period: Between planting to germination. Damage: The germinating eye, instead of growing into a pointed shoot, swells and becomes broad and flat (Fig. 11.3). Its surface becomes rough and looks like a wart. Damaged eyes fail to germinate. Monitoring: No established methods. Action level: Preventative chemical treatment at planting is recommended in prone areas in anticipation of damage. Chemical control: Chemicals added to the fungicidal dip or sprayed over the setts and soil during planting, including insecticides applied at planting to control wireworms, are not known to control this mite.
Fig. 11.2. A sugarcane sett with most of the roots damaged by symphylans. (BSES)
Cultural control: Good quality planting material should be used to offset wart-eye damage. Natural enemies: Unknown.
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Termites
Black field cricket
Isoptera: Mastotermitidae
Orthoptera: Gryllidae
Giant termite, Mastotermes darwiniensis Froggatt
Teleogryllus oceanicus (Le Gilliou) and Teleogryllus commodus (Walker)
Other minor species such as Coptotermes spp.
N (Main entry in Chapter 13 Pastures—summer
Distribution: The giant termite only occurs in Australia, and damages cane only in the Burdekin region, Qld, in sandy or sandy loam soils. Other termite species, such as Coptotermes spp., are found in all cane-growing areas in Australia. Pest status: Minor, widespread, irregular.
rainfall.)
Distribution: Both species are indigenous to Australasia, but Teleogryllus oceanicus was introduced into Hawaii (Zuk et al. 1995; 2001). In Australia, both species are found in NSW, Qld, SA, Tas. and Vic., while Teleogryllus commodus is also found in New Zealand.
Identification: Adult termites may be workers, soldiers or reproductives. Workers are white and 10–12 mm long. Soldiers are slightly longer and have powerful mouth parts. Breeders are wingless females (queens). Winged males and females swarm to make new nests. The wingless queens are about 15 mm long. Winged giant termite adults have a body length of about 18 mm, and including the wings are about 35 mm long.
Pest status: Minor, restricted, irregular.
May be confused with: Termites are distinct insects but superficially may be confused with ants (Hymenoptera).
May be confused with: Field crickets (gryllids) are generally distinct and are not confused with other insects. The two mentioned species are sibling species and it is difficult to distinguish them morphologically.
Hosts: Termites have a wide host range, especially crops growing in dry areas. Life cycle in sugarcane: Termites form colonies with the main nest underground, and workers access cane fields via underground tunnels up to 1 m deep. Nests are hard to find, as underground tunnels can be 1 m deep and may run more than 100 m from the nest to the field. Risk period: Planting to germination. Damage: Termites hollow out germinating setts, causing poor strike. They hollow out standing cane stalks over the whole length. This leaves only a shell, which may show few outer signs of damage. Monitoring: No methods available. Action level: Unknown. Chemical control: Control is carried out at the nest with registered products. Cultural control: In cane fields, the only practical control is to remove logs and trees from the edge of fields. Natural enemies: Unknown.
Identification: Adults and nymphs are shining black or brown with jumping hind legs. There is a strongly patterned area of venation on top of the folded forewing. Antennae are long and slender. Females have a long ovipositor protruding behind the tip of the abdomen (Fig. 13.32).
Hosts: In addition to sugarcane, Teleogryllus commodus is a pest of pastures and other field crops. Life cycle in sugarcane: Field crickets live in soil cracks in damp spots near drains and water channels, and in loam soils which form deep cracks. In wet areas, crickets live in burrows and cracks in soil along banks of channels and drains. Under dry conditions, crickets burrow deeply into the soil. After rain they are found near the surface. When furrows in planted cane are wet, the crickets can be found under loose soil crumbs in the hilled-up portion of the interspace during the day. They emerge to feed at night. Nymphs behave like adults. Risk period: Planting to germination. Damage: Signs of damage are found mainly in wet areas or in heavy soil where setts are planted close to the surface. Adults and nymphs eat out swelling eyes on setts, leaving a shallow, cleanly scooped-out hole. They also eat young shoot spikes as they emerge aboveground. Severe 309
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damage to buds causes gaps in stands that may need re-planting. Damaged shoots usually sideshoot with little yield loss. Significant damage has been recorded at Babinda and near Clare in the Burdekin region, Qld.
They may bore into the sett causing a clean-cut circular hole. Damage usually occurs before mole crickets are seen. Mole crickets are usually pests only in poorly drained soils.
Monitoring: Adult crickets can be found by tracking the calling songs of the males.
Monitoring: Mole crickets can be found more readily by following the burrows rather than by digging around damaged setts.
Action level: Unknown.
Action level: Unknown.
Chemical control: Not cost-effective.
Chemical control: No insecticides are registered.
Cultural control: Soil covering of setts reduces damage by crickets to eyes. Natural enemies: Unknown.
Cultural control: Damaged setts should be replaced. Natural enemies: Unknown.
Mole cricket Gryllotalpa sp. Orthoptera: Gryllotalpidae Distribution: Gryllotalpa is a cosmopolitan genus. It occurs in all cane-growing areas in Australia. Pest status: Minor, restricted, irregular. Identification: The adult is a strong-bodied insect, 25–30 mm long, covered with dull velvety dark brown to grey–brown short hairs The head is blunt and broadly convex. Wings are folded flat on the back of the body, and the front wings stretch only about half-way down the abdomen. Legs are short and thick, and are usually held close to the body. The first pair of legs are flattened and shaped for digging. Appendages about 5 mm long stick out from the abdomen. Nymphs are similar, except they are smaller and do not have wings (Fig. 15.2). May be confused with: Unlikely to be confused with any other pests. Hosts: Mole crickets have a wide range of hosts, but the host range of the species found in sugarcane has not been studied. Life cycle in sugarcane: Mole crickets make deep burrows in the soil. Eggs are laid in a chamber in the soil and the nymphs take several months to develop to adults. Mole crickets feed at night and adults are attracted to lights. Risk period: Planting to germination. Damage: Mole crickets burrow underground and chew at eyes and young shoots on sett, causing germination failure and dead heart.
Sugarcane wireworms Coleoptera: Elateridae Agrypnus variabilis (Candèze) = Lacon variabilis Candèze -Heteroderes spp. Conoderus spp. and other species (Samson and Calder 2003) Distribution: Native to Australia. Recorded in NSW, Qld, SA, Tas., Vic. and New Zealand. Agrypnus variabilis is found in wetter, poorly drained soils in central Qld and at Ingham. It is sometimes a pest in poorly drained parts of cane fields in north Qld. Pest status: Major, widespread, regular. Identification: Several species of wireworm damage sugarcane, mainly belonging to the genera Agrypnus, Conoderus and Heteroderes. The whole group requires taxonomic revision, but Agrypnus variabilis is the species nominated as ‘sugarcane wireworm’ (Samson and Calder 2003). Beetles of Agrypnus variabilis are up to 15 mm long and grey–brown. The larva grows to 20 mm long and has a firm, slightly flattened, shiny creamy-white segmented body with a hard yellow head and hard yellow forked rear end. Heteroderes spp. wireworms are larger than the sugarcane wireworm and more orange in colour. May be confused with: It is difficult to distinguish larvae of the different species of wireworms in cane fields, but only true wireworms (click beetle larvae) are pests of cane. Larvae of true wireworms possess a
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flattened rear end, while false wireworms (family Tenebrionidae) do not.
compensate for damage. Avoid planting susceptible varieties in poorly drained soils.
Hosts: Sugarcane, maize (Zea mays), Sorghum (Sorghum spp.), onions (Allium spp.), carrots (Daucus sp.), sunflower (Helianthus spp.) and cotton (Gossypium sp.). Wireworm larvae are omnivorous and will eat other soil insects, but damage caused to cane is believed to outweigh any beneficial activity.
Natural enemies: Unknown.
Life cycle in sugarcane: Beetles of sugarcane wireworm emerge during November–December and lay eggs in the soil. Eggs hatch in about 8 days. Larvae need wet soil during the first 3 weeks after hatching, but can handle dry weather afterwards. Larvae feed and grow over a 10-month period. The pupal stage lasts about 2 weeks. In central Qld, the sugarcane wireworm is mainly a pest of autumn plant-cane. Damage is less likely in spring plant-cane when larvae are almost fully fed and about to pupate. Risk period: Planting to germination. Damage: Wireworms bore into the eyes of germinating setts or ratooning stubble, or into the growing point on young shoots. Entry is by a small (< 2.5 mm) circular hole. Symptoms of damage are poor or patchy germination or ratooning and dead spindle leaves: ‘dead hearts’. Large shoots may compensate for damage by producing basal secondary shoots. Poor stooling varieties are probably more prone to wireworm damage. The impact of wireworms can be significant when ratoons are under stress in dry weather. Monitoring: Digging up the setts and checking for damage and presence of larvae is a reliable method to detect infestation, but a monitoring protocol has not been developed (Samson & Robertson 1996). Action level: Based on the known history of damage in each locality, preventive at-planting insecticide should be applied routinely to avoid germination failure.
Stenocorynus weevils Stenocorynus spp. Coleoptera: Curculionidae Distribution: Australian native. North and central Qld. Pest status: Minor, restricted, irregular. Identification: Grubs are short, fat, legless and up to 12 mm long. They range from white to yellow with pale yellow heads and black mouth parts. The adult weevil is 12 mm long. It is light brown with darker brown stripes along the head and body with the back smooth and convex. May be confused with: Adults may be confused with the sugarcane weevil borer, but they are light brown with darker brown stripes along the head and body. The back is smooth and convex. Larvae live in soil; whereas cane weevil borer larvae tunnel in stems. Hosts: Sugarcane. Adult weevils feed on nutgrass, Cyperus spp., flannel weed Sida cordifolia and common sida, Sida rhombifolia. Life cycle in sugarcane: Little is known about this species. Larvae have been seen damaging cane in August and begin pupating in October. During October–November, larvae, pupae and beetles were present in the soil. Larvae had all pupated by the end of November and beetles emerged from the soil in mid-January following heavy rain. Risk period: Prior to sett germination. Damage: Larvae chew the rind and the root band of germinating setts or ratoons, killing young roots and eyes. If the eye survives, root damage causes poor germination and weak growth. Larvae may also bore into setts, where damage impacts on growth and milling quality.
Chemical control: Preventative treatment of plant cane with insecticides is cost-effective and widely used. In ratoons, shoots killed by wireworms are usually compensated for by excess numbers of shoots produced by the cane plant.
Control: The pest occurs rarely, and no control measures are suggested.
Cultural control: Avoid autumn planting of cane in prone areas. Use vigorous cultivars to
Opogona glycyphaga (Meyrick) Lepidoptera: Tineidae
Sugarcane bud moth
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Distribution: Native to Australia. Occurs throughout Qld and northern NSW. Pest status: Minor, most frequent damage to cane occurs in the Herbert and Tully regions of Qld, irregular. Identification: Larvae are dull yellow with dark red–brown heads, up to 16 mm long and have dark blotches towards the head and small dark spots on the sides. The head and body are covered in hairs. Larvae live behind the leaf sheath base on mature stems. They make rough shallow feeding scars on the inside of leaf sheaths and on the rind and dig small entry holes at the tip of the bud. Pupae are found under leaf sheaths in silken cocoons covered with debris. Adult moths are 8 mm long with a shining purple head and thorax and yellow wings with purple tips. Moths rest with wings close to the body and antennae straight out in front. May be confused with: Both moth and larva are distinct. Damage to the eyes of the cane is characteristic of the sugarcane bud moth, but larvae living on cane before planting stay underground causing ‘dead heart’, a symptom that can be confused with those caused by wireworms, large mothborer, ratoon shootborer or black beetles. Hosts: Sugarcane, bananas, Musa spp. and granadillas, Passiflora spp. Life cycle in sugarcane: Moths probably breed all year, but large numbers of moths and larvae occur during autumn when the most damage to eyes occurs. Risk period: Planting to germination.
Cultural control: Stripping trash from plant sources may reduce damage to eyes (Jarvis 1927; Allsopp et al. 1993). Natural enemies: Unknown.
R ATO O N I N G
Cicadas Hemiptera: Cicadidae Brown sugarcane cicada, Cicadetta crucifera (Ashton) Yellow sugarcane cicada, Parnkalla muelleri (Distant) Green cicada, Cicadetta multifascia (Walker) Distribution: All native to Australia. Mainly found in NSW and Qld. Cicadetta multifascia is also recorded in SA. Both brown and yellow cicadas are common in all regions of coastal Qld. Brown cicada occurs at Mossman, Mulgrave, Tully, Abergowrie, Proserpine and Fairymead. Yellow cicada has damaged crops at Mulgrave and in the Burdekin region. Green cicada has damaged crops at Gin Gin. Pest status: Minor, restricted, irregular. Identification: The first four nymphal stages of brown and yellow species are white. Fully grown female nymphs of both species are 15 mm long, and males are up to 12 mm long. Mature nymphs of green cicadas have a green abdomen. Brown and green cicada adults are 14–18 mm. Brown cicada males are darkcoloured and the female is pale bodied (Fig. 11.4). The yellow cicada has a black
Damage: Larvae bore and kill bud eyes, ruining sources of planting material in susceptible varieties and causing germination failures if infested stalks are planted. Damaged buds are hollow. Larvae also bore into prop roots at the base of stools. Larvae living on cane before planting stay underground on the setts and bore into shoots and cause ‘dead heart’. Monitoring: Damage can be detected in the field by inspecting the buds. Setts are inspected prior to planting. Chemical control: Chemical control is not practical.
Fig. 11.4. A brown cicada nymph exposed in the soil around cane roots. (BSES)
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Z-shaped mark in the outer part of each front wing. Both sexes of the yellow cicada are cream– yellow. May be confused with: Unlikely to be confused with other soil insects. Hosts: Blady grass (Imperata cylindrica), crowsfoot grass (Poaceae), nutgrass (Cyperus spp.) and sugarcane. Life cycle in sugarcane: Most research has been done on the brown cicada, but all species have similar life cycles. Eggs are laid into midribs of sugarcane leaves. The brown cicada oviposits on the lower side of the midrib on fully expanded green leaves, while the yellow cicada lays into midribs of dead leaves. Eggs of the brown cicada hatch 8–11 weeks after laying (early March–April), and small (2 mm) nymphs drop to the ground and enter soil cracks. Fully grown nymphs emerge after about 8 months (between November– February) to develop into adults. Adults live for up to 7 days (Chandler 1982). Risk period: At ratooning, especially in old ratoons. Damage: Cicada nymphs tunnel in the soil beside the roots where they suck the sap and complete their growth. Heavily infested fields fail to ratoon, while lightly infested stools ratoon poorly. Small ratoon shoots sometimes become yellow and withered at the leaf tips. Areas of complete failure are surrounded by zones of poor growth. Monitoring: Nymphs are found by digging, and nymph skins can be seen on the soil surface if they have already emerged. Adults can be seen laying eggs on leaves. Action level: Light damage occurs with as few as 20 nymphs per stool. Moderately damaged stools commonly have 40–60 nymphs per stool. Heavily damaged (dead) stools have 80–300 nymphs per stool; however, no remedial action is feasible at time of infestation.
Natural enemies: Up to 40% of brown cicada eggs are killed by larvae of small unidentified wasps, and nymphs are attacked by the fungus Cordyceps heteropoda Kobayasi. Attack by the coastal brown ant is a major biological control factor of the brown cicada in sugarcane (Allsopp et al. 1993).
Soldier flies Diptera: Stratiomyidae Sugarcane soldier fly, Inopus rubriceps (Macquart) Yellow soldier fly, Inopus flavus (James) Distribution: Both species are native to Australia. Inopus rubriceps has been introduced to USA and New Zealand. In Australia, it is found in NSW, Qld, SA, Tas. and Vic. It occurs as a pest of cane in red volcanic soil at Innisfail, in silty loam soil at Macknade (Herbert River), and in mill areas from Proserpine south to Harwood in NSW. It is also a pest of pasture and sod in New Zealand. Inopus flavus occurs at Ayr, Proserpine, Mackay and Sarina (Qld). Another unidentified species has been found in sugarcane at Giru (Qld). Pest status: Major, restricted, generally irregular, but can be regular in certain areas in the central district of Qld. Identification: Inopus rubriceps: the adult female has a black body and an orange–red head. The adult male has dark grey–brown body and head. Larvae are legless maggots with tough-ribbed skins (Fig. 11.5). They are oval in cross-section and tapered at both ends, white to brown, and up to 14 mm when fully grown. Eggs are white, long and thin with rounded ends, and 1 mm long. Inopus flavus: the adult female is about 12 mm long with lightly smoky wings and small black eyes. Its body, head and legs are orange–yellow.
Chemical control: No insecticide is registered for controlling cicadas. Cultural control: Summer fallow free of sugarcane volunteers breaks the breeding cycle. Plough-out/re-plant of infested areas leads to increased cicada damage.
Fig. 11.5. Soldier fly larva, 14 mm long. (BSES)
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The male is about two-thirds the size of the female and has large black eyes that cover most of the head. Its body is black, head brown–black and legs yellow. Immature stages are similar to I. rubriceps. May be confused with: Unlikely to be confused with other pests. Conical pits made by larvae in roots are unlike the cylindrical holes made by symphylans. Hosts: Larvae feed on the roots of many plants other than sugarcane. Native grasses are their natural food, and infestations may develop quickly when grasslands are first brought under cane. They also feed on introduced grasses, cereals, legumes (such as Dolichos and cowpea (Fabaceae)), and a wide range of vegetable crops (Allsopp and Robertson 1988). Life cycle in sugarcane: Most larvae mature into flies in 1 year, while the rest take 2 years. Sugarcane soldier flies emerge from March to July, depending on the weather. Numbers peak usually in May. The yellow soldier fly emerges earlier with the peak usually in April. Adults emerge during the morning. Females usually mate and lay eggs on the day of emergence, and live for only a few days. Eggs are laid in clumps of up to 200 about 10 mm below the soil surface, or between the soil and dead plant matter. They hatch in 1–3 weeks depending on temperature. The larvae then feed on roots. The pupal stage lasts about 1 month. Pupal cases can be found on or just beneath the soil surface for several months after adult emergence. Allsopp (1990) analysed data collected over 23 years and suggested that outbreaks in southern and central Qld followed years with relatively low minimum temperature and rainfall in mid–late summer. In southern Qld, outbreaks also followed years with lower May minimum temperatures, higher June rainfall and higher July–August minimum temperatures. In central Qld, there was also a relationship with October rainfall. Risk conditions: Planting to germination, and from harvesting to ratooning. In southern and central Qld, damage is more likely in years with lower January–February minimum temperatures and lower February rainfall than normal. In southern Qld, outbreaks followed years with lower May minimum temperatures, higher June rainfall and higher July–August minimum
temperatures, while dry Octobers presage damage in central Qld. Damage: Larvae feed on roots, including sett roots. Small larvae cut hairs from the roots, and, when larger, burrow into the roots. Larvae can cause a poor strike if high numbers occur at planting. Larvae often cause poor ratooning, with underground buds failing to germinate after harvest. Affected stools produce few or no shoots resulting in gaps in the field (Fig. 11.6). Stools on the edge of cane blocks often look healthy. Monitoring: Larvae are usually found within 15 cm of the soil surface along rows throughout the year. Larvae are difficult to find in winter as most are very small. Some large larvae may still be found as they will become adults after 2 years in the soil. Larvae are most easily found by digging around roots from about October through to April. Action level: Unknown. Chemical control: No chemical control recommended. Cultural control: The following can be done: • harvest affected blocks as early as possible in the season • have a grass-free break from cane, e.g. a herbicide fallow after spray-out of the old ratoon under trash, or a rotation with another non-grass crop such as soybeans • plant late, after the flight period (after June) • plant with minimum tillage following the herbicide fallow • choose varieties that ratoon strongly; some varieties may also help to limit the rate of population increase
Fig. 11.6. Empty patches are indicative of root-feeding by soldier fly larvae. (BSES)
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• harvest plant and early ratoon crops when ratooning conditions are good • do not plough-out/re-plant or plant early in autumn after a badly infested ratoon. Natural enemies: Soldier flies are attacked by parasitic diapriid wasps (Neurogalesus sp. Hymenoptera: Diapriidae), the fungus Metarhizium in the larval stage, and by predators, such as wireworms that eat the pupae (Robertson 1987).
Ratoon shootborer Ephysteris promptella (Staudinger) Lepidoptera: Gelechiidae Distribution: Africa, Mediterranean Europe and Asia, introduced into Australia probably from Indonesia (Jarvis 1927). Pest status: Minor, restricted, irregular.
Monitoring: The larval pin-hole entrance to the tunnel can be seen by removing the bottom leaves of the shoot. Action level: Unknown. Chemical control: No insecticides are known to control this insect, probably because larvae are well hidden and most damage has been done by the time the insect is found. Cultural control: No cultural control methods have been developed. Natural enemies: Jarvis (1927) stated that no natural enemies are recorded on this pest perhaps because it is an introduced species.
G ROW T H
Sugarcane spider mite
Identification: Eggs are pale green–yellow, flecked with shiny blue and gold–pink, and are 0.7 mm by 0.4 mm. Larvae are yellow and about 5 mm long. Pupae are boat-shaped, about 4 mm by 1 mm in a frail silk cocoon covered with frass. Adult moths are 3–4 mm long with a wingspan of 7–10 mm; males are smaller than females. The wings are dark grey with white flecks on females and black dots on males.
Oligonychus zanclopes Beard and Walter Acarina: Tetranychidae
May be confused with: Larvae could be confused with larvae of the large mothborer. ‘Dead heart’ symptoms can be confused with those caused by wireworms, large mothborers or black beetles.
May be confused with: Adults and nymphs may be confused with other mites, although the key of Beard et al. (2003) is diagnostic.
Hosts: Sugarcane. Reported to attack maize and sorghum in South Africa (Drinkwater 1986).
Life cycle in sugarcane: Adults and nymphs live on the undersides of leaf blades, where they produce extensive webbing. Details of development periods are not known.
Life cycle in sugarcane: Eggs laid on the cane stalk probably take a few days to hatch. Larvae tunnel into the stalk and eat the juicy bottom part. Pupae occur on the soil surface or in dead twisted leaves. Adults emerge 10–14 days after pupation. Risk period: At ratooning, especially during dry periods. Damage: Larvae bore into young shoots, often killing them and causing ‘dead hearts’. Damage is only in ratoons, especially in crops harvested mid-season.
Distribution: Native to eastern Qld. Pest status: Minor, irregular. Identification: Mainly identified from the damage symptoms. Adults can be separated from other species of Oligonychus using the key of Beard et al. (2003).
Hosts: Sugarcane, rice (Oryza sativa).
Risk period: Summer–autumn, particularly during dry periods. Damage: Feeding causes white to brown blotches on the leaves, which become very obvious in some cultivars. The undersides of leaves also have extensive webbing. Monitoring: No established methods. Action level: No established thresholds. Natural enemies and heavy rainfall appear to reduce populations dramatically. 315
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Chemical control: No chemical control is registered for spider mites in sugarcane. Cultural control: Some cultivars suffer less damage from spider mites than others. Natural enemies: Unidentified coccinellid beetles feed on adults and nymphs.
Locusts Orthoptera: Acrididae N (Main entry in Chapter 18 Locusts and grasshoppers of
pastures and rangelands.)
Australian plague locust, Chortoicetes terminifera (Walker) Migratory locust, Locusta migratoria (Linnaeus) Spur-throated locust, Austracris guttulosa (Walker) = Nomadacris guttulosa (Walker) Yellow-winged locust, Gastrimargus musicus (Fabricius) Pest status on sugarcane: Minor, widespread, irregular. Hosts: Locusts are polyphagous and will feed on a very wide range of plants. Life cycle in sugarcane: Locusts usually live as scattered individuals. Sometimes, they build up into large numbers forming hopper bands and adult swarms. When in swarms or bands they do economic damage. Adult Australian plague locusts sometimes appear in early summer, carried in on a cold front. They can go through two to three generations each summer. Eggs stay in the soil over winter. Infestations in sugarcane usually last only one to two generations before dying out. Migratory locust swarms usually originate on the Central Highlands. They have three or four overlapping generations each year. The hopper stage lasts only 4 weeks in summer. Adults lay soon after maturity and continue to do so for 1–2 months. Young hoppers can move into and feed on cane. Swarms of adult spur-throated locusts may appear in coastal areas during October– November, and have only one generation per year. Eggs are scattered in cultivated fields or grass, but many are laid in fallow fields under legume cover. Hoppers hatch in early to mid-
February, and usually eat grasses in weedy fields before feeding on cane. The nymphs grow to adults by mid-April. Adults then overwinter in large cane and scrub before laying eggs in October–November. The yellow-winged locust has two generations per year. In the summer, eggs hatch in 17 days to produce hoppers in November–December. These mature to adults in February–March. The eggs laid by these adults hatch from August onwards. This second generation is mature in November– December. Egg pods contain about 60 eggs and are laid in cultivated fields, hard bare soil or the forest floor. Risk period: Summer–Autumn. Damage: Adult locusts and hoppers chew the leaves of sugarcane, and in heavy infestations only leaves and midribs remain. Weeds then grow because the cane canopy is lost. All locusts are pests from time to time, usually coming from outside cane areas. Australian plague locust can occur in southern districts, migratory locusts can occur in northern and central areas, spur-throated locusts can occur as far south as Bundaberg, and yellow-winged locust can occur in all districts. Chemical control: Generally, locusts are not worth trying to control with insecticides. The long-term damage they do to cane is usually small and infestations die out within a couple of generations. Because hoppers usually group together, they are easier to spray than adults. Cultural control: Good kills of eggs can be achieved by cultivating the egg beds before the hoppers hatch. Natural enemies: A wide range of parasitoids and predators attack different locust stages (Baker 1983). Egg parasitoids are extremely active in northern sugarcane regions. The fungus Metarhizium anisopliae is used successfully against Locusta migratoria in Australia (Hunter et al. 1999). Different strains of the same fungus are virulent to the other species of locust.
Aphids Hemiptera: Aphididae Sugarcane aphid, Melanaphis sacchari (Zehntner)
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Corn aphid, Rhopalosiphum maidis (Fitch) Hemiptera: Pemphigidae Oriental grassroot aphid, Tetraneura nigriabdominalis (Sasaki) Distribution: Sugarcane aphid is probably indigenous to Australia, but occurs in North and South America, Africa and Asia. Corn aphid is cosmopolitan in distribution and found all through Australia. Oriental grassroot aphid is probably native to Australia and occurs on most of the mainland. It is recorded in South America, Asia and Southern Europe. Pest status: Minor, restricted, irregular. Identification: Sugarcane aphid is oval and light-coloured, ranging from yellow–white to green- or brown-tinged. The corn aphid is larger, longer, and darker, ranging from pale olive to yellow–green. The base of each siphunculus has an olive–brown blotch. The oriental grass root aphid is found only on roots and has small cone-like siphunculi. May be confused with: The sugarcane aphid and corn aphid look similar but can be easily separated by morphological features. Hosts: The sugarcane aphid attacks grasses of the genera Saccharum, Sorghum, Oryza, Echinochloa, Panicum and Pennisetum (Mead 1978), while the corn aphid mainly attacks corn (maize) but has a very wide host range that includes several gramineous and legume crops. Life cycle in sugarcane: Aphids may lay eggs or produce live nymphs. There are winged and wingless forms. Risk period: Aphids do not generally pose a high risk in sugarcane. Damage: The sugarcane aphid feeds in colonies on the undersides of leaves. Heavily infested leaves become spotted and dry up. Sooty mould also grows on the honeydew produced by of the aphids. This species does not transmit viruses. The corn aphid is not common in sugarcane, but is usually found in grasses such as wild sorghums. This aphid spreads sugarcane mosaic virus (Fig. 11.7). The oriental grass root aphid feeds on roots, sometimes producing a red–purple colour in the leaves.
Fig. 11.7. Sugarcane mosaic virus symptoms on leaves (left) and cane (right). The disease is transmitted by corn aphids and other aphid species. (BSES)
Many other aphid species can be found in sugarcane crops. Some of these can spread sugarcane mosaic virus disease, even though they feed on sugarcane for only short periods. Monitoring: Visual inspection under the leaves. Action level: Control of aphids is usually not cost-effective. Chemical control: No insecticides are registered for the control of aphids in sugarcane. Natural enemies: Ladybirds and syrphid flies usually control dense infestations.
Ground pearls (Margarodids) Hemiptera: Margarodidae Pink ground pearl, Eumargarodes laingi Jakubski White ground pearl, Promargarodes australis Jakubski Distribution: The pink ground pearl may be an exotic species, while the white ground pearl is indigenous to Australia. Both species occur in NSW and Qld. Pest status: Minor, restricted, irregular. Identification: Adult female ground pearls resemble mealybugs, but do not have a white powdery covering. Pink and white ground 317
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females are often found on the soil surface in the late morning. After a few days, the female forms a small hole in the soil, and lays up to several hundred eggs and then dies. Eggs hatch in 2–4 weeks, producing tiny white nymphs with legs and antennae. In 2–3 weeks, the nymphs settle beside a root, change to cysts and lose their legs and antennae. The cysts feed through a fine tube. Risk period: Damage is mostly seen in autumn months.
pearls are named from the colour of the adult female (Fig. 11.8). No adult males are known. The young of ground pearls are enclosed for most of their lives in a round, shiny cyst which crumbles like mica when rubbed. The cyst of the pink ground pearl is creamy to brown (Fig. 11.9). Its skin is tough and not shiny. The cyst turns pink in colour just before the adult is ready to emerge. The cyst of the white ground pearl is harder and shiny white to yellow.
Damage: Pink ground pearls occur mainly on red volcanic soils and some sandy soils around Bundaberg, while white ground pearls occur on all soil types. The nymphs of all species form cysts (round shiny sacks) in the soil and attack roots through long feeding tubes. Pink ground pearls cause poor growth patches which expand as the insects spread. At first, cane is stunted and the leaves yellow, and the stool eventually dies. Poor patches often spread by cultivation along rows. Yield losses of about 75% are common. White ground pearls can cause similar effects but may also result in little or no yield loss even when numbers are high.
May be confused with: Unlikely to be confused with other insects in cane.
Monitoring: Cysts can be found by digging under cane plants.
Hosts: Sugarcane. Species of ground pearls are recorded as minor pests on the roots of turf grass.
Action level: Not available.
Fig. 11.8. White (left) and pink (right) ground pearls (larger individuals are about 4 mm long). (BSES)
Life cycle in sugarcane: The pink ground pearl has a 1-year life cycle, while the white ground pearl takes 2, 3 or 4 years to become adult. Adults emerge mostly during October–January but also September and February. Many ground pearl
Chemical control: No chemical is registered for this species. Cultural control: Some cultivars are more tolerant to pink ground pearls than others. In Bundaberg, growing Q147 and Q135 can minimise the economic impact of the pest. Infested blocks should be ploughed out early and fallowed for 12 months. Ploughing or rotary hoeing several times during the fallow will reduce pest numbers. It is also very important that machinery working in infested areas be cleaned thoroughly before moving to other fields, especially in spring–summer when adults, eggs and nymphs can be transported easily in moist soil stuck to implements and vehicles. Natural enemies: Unknown.
Mealy bug Fig. 11.9. Pink ground pearls (cysts) attached to sugarcane roots. There are about 15 cysts in the picture, the largest of which is 4 mm diameter. (BSES)
Pink sugarcane mealybug, Saccharicoccus sacchari (Cockerell) Hemiptera: Pseudococcidae
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Distribution: Introduced to Australia. Occurs in North and South America, Africa and Asia. In Australia it occurs in all cane areas.
North-western Australia: Perkinsiella thompsoni Muir
Identification: The adult is a soft, oval, wrinkled, wingless insect up to 5 mm in length, covered with white powder. Nymphs are similar to adults but smaller. Colonies are usually found on stalks behind leaf sheaths, but can also occur on the underground stems.
Distribution: Both species are probably native to Papua New Guinea. Perkinsiella saccharicida is widespread in Asia and was introduced to North, Central and South America and the Hawaiian Islands. It is also recorded from Mauritius, Reunion, Madagascar and South Africa. In Australia, it is common in NSW and Qld (White et al. 1995).
May be confused with: Ground pearls, but these do not have a white powdery covering.
Pest status: Major only as a vector of Fiji leaf gall virus, widespread, irregular.
Hosts: In Australia, it attacks only sugarcane.
Identification: Adults are about 5 mm long, brown–black in colour, and taper towards the rear from a broad head. When approached, adults resting on a cane stalk will retreat rapidly to the other side of the stalk. Adults will often move sideways, giving them the common name of ‘sidewinder’. Nymphs are plump, wingless mottled-brown insects, and have five nymphal stadia.
Pest status: Minor, restricted, irregular.
Life cycle in sugarcane: Mealybugs survive harvests on underground parts of the plant. Crawlers reappear aboveground in spring following the formation of stalks. As new nodes are formed, colonies start behind the leaf sheaths. Numbers are highest in February– March and then drop due to lower temperatures and parasite and predator activity. Growth from eggs to adults takes about 4 weeks. Movement between fields may be on planting material, by wind or by ants. Damage: Mealybugs usually do not damage cane, but very high numbers are usually associated with stressed or weakened plants. Sooty mould also grows on honeydew produced by the pest.
May be confused with: Other delphacids associated with sugarcane, especially Eumetopina spp., which occurs on the Torres Strait islands and in Bamaga, and is considered an important quarantine issue for Australia. Perkinsiella saacharicida is morphologically similar to Perkinsiella thompsoni Muir from the Ord (WA), but can be distinguished by the male genitalia and DNA technology.
Monitoring: Visual inspection of plant stalks.
Hosts: Sugarcane.
Action level: Not available.
Life cycle in sugarcane: Adults prefer the upper stalk where they hide and feed in the young leaf axils. Banana-shaped eggs are laid mainly in the upper leaf midrib near the leaf axil. Upon hatching, young nymphs move down the stalk and behind the leaf sheaths. Older nymphs are found on or near the leaf spindle. Planthopper numbers increase rapidly in summer with nymph numbers peaking in January–February and adults in February–March. Adults can swarm in late summer and are strongly attracted to lights. Planthoppers are not common in fields between May and November.
Risk period: Autumn months.
Chemical control: Chemical control is not required. Cultural control: Tighter-trashing varieties are more prone to attack. Natural enemies: The parasitic wasp Anagyrus saccharicola Timberlake (Hymenoptera: Mymaridae), the fly Cacoxenus perspicax (Knab) (Diptera: Drosophilidae:) and the fungus Aspergillus parasiticus Speare help to control numbers, particularly in late summer.
Planthoppers Hemiptera: Delphacidae Eastern Australia: sugarcane planthopper, Perkinsiella saccharicida Kirkaldy
Risk period: Late summer–autumn. Damage: Nymphs and adults suck sap from the leaves, causing yellowing and mottling. Sooty mould grows on the honeydew produced by planthoppers. Most importantly, Perkinsiella 319
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Fig. 11.11. Sugarcane froghopper (5 mm long) that causes sugarcane blight. (BSES) Fig. 11.10. Symptoms of Fiji leaf gall virus, which is transmitted by sugarcane planthopper. (BSES)
saccharicida is the carrier of Fiji leaf gall virus (Fig. 11.10), which was a major disease in all canegrowing districts south of Mackay (central Qld). The disease still occurs from Maryborough south. Nymphs can acquire the virus within a few days of hatching. Older nymphs spread the virus easily and this persists as they change to adults. Monitoring: Visual inspection of the underside of leaves when the crop is waist to shoulder high. The presence of the disease in only a few plants is enough to deem the whole block as diseased. Action level: No action is usually taken against this pest. Chemical control: No chemicals are registered for the control of Perkinsiella in cane fields. Cultural control: Resistant sugarcane varieties provide good control of Fiji disease. Natural enemies: The egg predators Tytthus spp. (Hemiptera: Miridae) and the parasitoids Anagrus sp. (Hymenoptera: Mymaridae) and Ootetrastichus beautus Perkins help reduce planthopper numbers in summer.
Sugarcane froghopper Euryaulax carnifex (Fabricius) = Eoscarta carnifex (Fabricius) Hemiptera: Cercopidae Distribution: Native to Australia. Occurs in Qld where it has been found on cane grown the Herbert valley, the Tully district and at Gordonvale. The genus Euryaulax was wrongly synonymised with the oriental genus Eoscarta (Liang and Fletcher 2002). Pest status: Minor, restricted, irregular.
Identification: Adults are 5 mm long, found on the top of cane leaves. They have an orange head and body, and dark purple–black front wings with central orange patches (Fig. 11.11). Nymphs are yellow or pink and live on roots inside foam that resembles spittle. May be confused with: Unlikely to be confused with other cane pests. Hosts: Sugarcane. Life cycle in sugarcane: Adults lay eggs in the soil. Eggs laid in autumn do not hatch until the following summer, usually in December. Nymphs live underground in cracks, at the surface under cane trash or among roots at the base of sugarcane stems. Nymphs suck sap from cane roots but can be difficult to find. Adults suck sap from leaves and inject saliva that slows water and nutrient flow. Affected tissues turn yellow and die. Heavily infested fields have lower growth, due to loss of healthy leaf area. There are usually three and sometimes four generations of froghoppers each year, although later generations overlap. Risk period: Autumn months. Damage: Infested fields look ‘scorched’. Fresh damage appears as short yellow and red streaks in the top leaves. Yellow chlorotic streaks extend and develop red central stripes. Khaki-coloured dead areas occur where streaks reach leaf edges. Advanced symptoms are seen on fifth and older leaves. Symptoms were first described as ‘froghopper blight’ by Magarey et al. (1988). Monitoring: Visual inspection of plants. Action level: No method of control is practised against this pest in sugarcane. Chemical control: Not cost-effective. Cultural control: Unknown.
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Natural enemies: Unidentified assassin bugs, wasps and fungal diseases affect adult froghoppers. Nymphs are attacked by larvae of an unidentified hover fly.
Linear bug Phaenacantha australiae Kirkaldy Hemiptera: Colobathristidae
Black beetles Coleoptera: Scarabaeidae African black beetle, Heteronychus arator (Fabricius) = Heteronychus sanctaehelenae Blanchard Black beetle, Metanastes vulgivagus (Olliff) = Pentodon australis Blackburn
Distribution: The linear bug is thought to be native to Australia, but it may be an introduced species as it appeared suddenly in 1917 (Allsopp et al. 1993). Phaenacantha australiae occurs through northern and central Qld.
Distribution: Heteronychus arator originated in sub-Saharan Africa, and it was introduced to Australia. In Australia, it is recorded from NSW, Qld south of Maryborough, SA and WA. Metanastes vulgivagus is native to Australia, recorded from NSW, NT, Qld and SA.
Pest status: Minor, restricted, irregular.
Pest status: Minor, restricted, irregular.
Identification: The adult is long and thin, about 9 mm in length, and orange–brown to dark green–brown. Nymphs are similar to adults but are wingless and orange–yellow. Nymphs go through five growth stages.
Identification: Beetles of two species are shining black above and red–black underneath. The upper surface of the back is smooth and the wing covers have several parallel fine ribs along their length. Larvae of both species grow to 25–30 mm long. The soft part of the body is blue to grey–white, and the head is rough-surfaced and dark red–brown. They do not have a hair pattern like canegrubs.
May be confused with: Unlikely to be confused with other pests of sugarcane. Hosts: Sugarcane, wild grasses. Life cycle in sugarcane: Phaenacantha australiae occurs throughout the year on grass or cane, but numbers peak in spring. Eggs are laid on the soil surface around the roots of grasses. Young nymphs feed in protected areas near the base of plants. Linear bugs are usually found near grassy fields. Risk period: Spring–summer, but it rarely causes significant damage. Damage: Linear bugs feed on the under-side of leaves, with the leaves turning yellow and the tips drying. Where the affected leaves face the sun, they can turn red–purple. Damaged leaves are more prone to fungal attack. Monitoring: Adults are seen easily by checking the under-side of the leaves. Action level: Not available. Chemical control: Not cost-effective. Cultural control: Weed control in cane fields reduces the infestation significantly. Natural enemies: There are a number of unstudied natural enemies that impact on linear bugs populations.
May be confused with: Beetles of two species are very similar in appearance. Metanastes vulgivagus is usually about 15 mm long, but can vary from 12–17 mm long, while Heteronychus arator is slightly smaller (9–15 mm long). The African black beetle has two rows of grooves on the back of the last segment beneath the rear of the wing covers; these are not present in the black beetle. Hosts: Sugarcane, maize, pasture, perennial ryegrass (Lolium perenne), clover (Trifolium spp.) and Paspalum spp. Life cycle in sugarcane: Both species have a 1-year life cycle. The beetles mostly emerge from mid-summer until the end of autumn and then rest during winter. Adults become very active in spring, when they damage cane. Egg-laying begins early in September and continues until February. Eggs of both species are laid singly in soil. They hatch in about 2–4 weeks depending on temperature. Both species have three larval instars that take about 3 months in total. The pupal stage then lasts 2 weeks. Over the spring– autumn period there is usually a range of insects at different growth stages present at the one time. Grubs feed mainly on organic matter in the soil. Older grubs may feed on roots of grass. 321
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Risk period: Spring–summer and autumn.
Canegrubs (white grubs)
Damage: Beetles of both species damage plant and ratoon cane. They chew ragged deep holes into young shoots causing dead hearts. A shoot attacked near the base cannot recover, while shoots attacked above growing points may sideshoot. Therefore, it is not unusual in northern Qld for a satisfactory stand of cane to develop after a heavy attack. In southern Qld, attack by either species is much more common and heavier in spring than in autumn. Damage by black beetle in northern Qld occurs as much in autumn as in spring. At Innisfail, damage caused by larvae can sometimes be just as severe as that caused by adults. In southern Qld, damage from African black beetle is most common when cane is newly planted into old pasture country.
Coleoptera: Scarabaeidae
Monitoring: Beetles and feeding symptoms can be seen by inspecting the dead hearts. Chemical control: No insecticides are registered for control of black beetles in Qld but some may be registered in NSW. Cultural control: The following can be done: • avoid planting susceptible varieties • repeat ploughing of pasture areas before planting • long-fallow, preferably with a legume (bean) • autumn-plant rather than spring-plant • observe strict quarantine against transfer of contaminated material or live Heteronychus arator specimens to north of Maryborough. Natural enemies: Unknown.
Soil insects: preventive management Preventative treatment against soil-inhabiting pests needs to be anticipated prior to the risk period. It is usually not possible to apply insecticides against soil pests during the growing season. In this chapter, ‘action level’ is not given for soil insects that require preventive management. Usually, a history of soil-insect damage in an area indicates the need for preventive management.
Species of Antitrogus, Dermolepida, Lepidiota and Rhopaea. General introduction Nineteen species of canegrubs, all native to Australia, damage sugarcane crops in NSW and Qld. Most canegrub species are major pests of cane and inflict frequent high yield losses. The greyback canegrub (Dermolepida albohirtum) is a key pest from Mossman (far north Qld) to Sarina (central Qld), with estimated annual losses of up to $10 million. Other major species are Lepidiota frenchi, L. negatoria and Antitrogus parvulus. Annual yield losses and cost of pesticide application against canegrubs far outweigh that caused by all other pests combined, including vertebrates and nematodes. Canegrubs are larvae of scarab beetles, and all species have three larval instars. The adults lay their eggs in the soil and hatchlings feed on fine roots and soil organic matter. Second- and third-instar grubs move closer to the roots and feed on the root mass. Some species have a typical 1-year life cycle, such as Dermolepida albohirtum, Lepidiota grisea, Lepidiota squamulata and Antitrogus consanguineus. Other species , such as Lepidiota frenchi, Lepidiota negatoria, Lepidiota crinita and Antitrogus parvulus, show a typical 2-year life cycle. The remaining species show a variation from the typical life cycle, where a proportion of the species exhibits a 1-year life cycle while the other will exhibit a 2-year life cycle. Identification: Larval stages can be identified using the raster pattern on the ventral surface of the last abdominal segment (Fig. 11.13). The adults of all cane beetles are medium to large (2–3 cm), light to dark brown beetles. Some have a covering of fine hairs (Fig. 11.14), whereas others are covered with white scales (Fig. 11.15). May be confused with: Larvae of some species have an almost identical hair pattern, and are therefore hard to tell apart in their larval stage, and can be confused with other white grub species that are not pests of cane. DNA techniques may be used to separate these species. Adults are morphologically distinguishable, and most can be identified by the naked eye or by using a magnifying glass.
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SPECIES (major pest in bold)
GEOGRAPHICAL DISTRIBUTION
Bundaberg canegrub (Fig. 11.13a) caudata canegrub (see Fig. 11.13c) Childers canegrub (Fig. 11.13b) consobrina canegrub (Fig. 11.13c) French’s canegrub (see Fig. 11.13c) Froggatt’s canegrub (Fig. 11.13d) grata canegrub (Fig. 11.13e) greyback canegrub (Fig. 11.13f) grisea canegrub (Fig. 11.13g) Nambour canegrub (Fig. 11.13h) negatoria canegrub (Fig. 11.13i) noxia canegrub (Fig. 11.13j) picticollis canegrub (Fig. 11.13k) planiceps canegrub (no image) rhopaea canegrub (Fig. 11.13l) Rothe’s canegrub (Fig. 11.13m) sororia canegrub (Fig. 11.13n) southern one-year canegrub (Fig. 11.13o) squamulata canegrub (Fig. 11.13p)
southern Qld north Qld southern Qld north Qld north Qld–southern Qld far north Qld throughout Qld north–central Qld north Qld southern Qld–northern NSW central Qld–northern NSW southern Qld southern Qld northern NSW southern Qld–northern NSW north Qld and NT north Qld southern Qld north Qld–southern Qld and WA
Hosts: Canegrubs attack a wide range of gramineous crops such as cane, maize and sorghum, and a range of wild grasses and pasture such as Guinea grass (Panicum maximum), blady grass (Imperata spp.) and Brachiaria spp. Depending on the grub species, crops such as bananas, pawpaw, peanuts, pineapples and taro are also attacked. Dermolepida albohirtum adult beetles feed on palm trees, bananas, wattle and figs (Ficus spp.). Life cycle in sugarcane: Cane beetles fly and lay eggs between September and January, depending on weather and the species. Cane is damaged after the grubs enter their third and final instar. In those species with a 1-year life cycle, this occurs in late summer or early autumn, 2–4 months after the eggs are laid. Species with a 2-year life cycle spend about 10 months in the first two instars and reach the third instar in the spring after beetle flight. All species pupate deep in the soil before turning into beetles. Typical 1-year and 2-year life cycles are discussed more fully under greyback and French’s canegrubs, respectively. Risk conditions: Depending on the species, damage may be seen during late summer or early autumn. One-year cycle grubs typically damage semi-mature crops in autumn, while
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2-year cycle grubs typically damage young plants or ratoons in spring. Hence, the risk period in the case of canegrubs commences well before damage is seen. Damage: Larvae in the soil destroy the roots of sugarcane, which starves the cane of soil moisture and nutrients. In light infestations, leaves turn yellow and grubs cause lodging of the plants. Heavier infestations, especially by major species, will rob the plant of all roots and stools and will cause plant ‘tipping’, and ultimately plants will die or be pulled out during harvest, which creates gappy ratoons. The extent of damage and lodging depends on the number of grubs present, grub species, the variety of cane, the age of the crop and the growing conditions. The time at which damage becomes noticeable depends on the life cycle of the species involved. Monitoring: The following can be done: • digging up of cane stools and counting the number of grubs per stool • aerial photography of damage • satellite imaging • monitoring beetle activity on feeding/ aggregation trees 323
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• visual inspection of damage prior to harvest
a
• setting light traps that attract adult beetles and can be used to roughly estimate population densities. Action level: Depends on the species present, soil type, climatic conditions and cane variety. Preventative treatment needs to be applied at planting (or at fill-in stage) ahead of time and prior to risk period. For 1-year type grubs, it is not feasible to take action after damage appears during the same crop cycle. Risk assessment is required to avoid over- or under-estimation of infestation. However, curative treatment is possible with 2-year species. The best method for early warning is to dig up a number of stools in areas where grubs are suspected. If grubs are encountered frequently, then a ratoon treatment is recommended around September–October. Chemical control: Controlled-release and liquid formulations of insecticides are available for application in plant cane, either at planting or at fill-in stage, and in ratoons. Liquid and granular insecticides are available for treating ratoons. Fungal biocides are also available for application in plant cane only to control the greyback grub. Cultural control: Cultural methods such as well-timed deep ploughing, trap cropping and manipulation of planting and harvesting dates can be used to minimise the impact of grubs. Natural enemies: Canegrubs are attacked by digger wasps (Campsomeris spp. Hymenoptera: Scoliidae) and sometimes by robber flies (Family Asilidae). Wireworms also eat the eggs and larvae, but do not give economic control. Grubs are also prone to a number of diseases, such as that caused by the protozoan Adelina sp., which is an efficient mortality factor against greyback grubs in certain areas in far north Qld. The green muscardine fungus, Metarhizium anisopliae, infects grubs in soil (Fig. 11.12); natural infections exert some control. A commercial formulation is available as a natural biocide. Milky disease caused by the bacterium Paeaibcillus popilliae may reduce numbers of canegrubs towards the end of their larval stage. Nematodes are sometimes found in canegrubs and may cause high death rates, but the cost of producing them as biocides is the main factor limiting their use. Mammals such as bandicoots,
b
Fig. 11.12. (a) Canegrub larva and (b) adult greyback canegrub infected with green muscardine fungus, Metarhizium anisopliae. (CSIRO: R. Milner)
and birds, such as ibis, prey upon grubs in soil, but they are generalist feeders and their impact on the overall population is minimal. The table on page 323 lists all canegrub species found in all sugarcane areas and their status.
Bundaberg canegrub Lepidiota crinita Brenske = Lepidiota trichosterna Lea Distribution: Southern Qld. Damage to cane is only at Bundaberg and Gin Gin, usually in forest loam and sandy soils. It is often found with negatoria canegrubs in forest loams and with Childers canegrubs in clay loams. Pest status: Minor, restricted, irregular. Identification: The larva has a hair pattern of two parallel rows, each with about 15 short hairs. There is no clear gap between the two rows (Fig. 11.13a). Adult beetles are about 25 mm long, bright red–brown with the upper surface coarsely but evenly punctured; each puncture
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(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
(i)
(j)
(k)
(l)
(m)
(n)
(o)
(p)
Fig. 11.13. Canegrub larvae showing end views of rasters. Raster patterns are useful for identifying common pest species of canegrubs: (a) Bundaberg canegrub; (b) Childers canegrub; (c) consobrina, caudata and French’s canegrubs have a similar raster pattern; (d) Froggatt’s canegrub; (e) grata canegrub; (f) greyback canegrub; (g) grisea canegrub; (h) Nambour canegrub; (i) negatoria canegrub; (j) noxia canegrub; (k) picticollis canegrub; (l) rhopaea canegrub; (m) Rothe’s canegrub; (n) sororia canegrub; (o) southern one-year canegrub; (p) squamulata canegrub. (BSES)
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are about 25 mm in length (slightly larger than beetles of French’s canegrub). May be confused with: It is difficult to distinguish caudata larvae from consobrina (Fig. 11.13c) and French’s canegrubs. Adults are morphologically distinguishable. Hosts: Sugarcane is the main host, also a common pasture pest on the eastern Atherton Tableland.
Figure 11.14. Adult Bundaberg canegrub (25 mm long) (left); Childers canegrub (20 mm long) (right). (BSES)
has a long, thin white scale about as long as the puncture (Fig. 11.14). May be confused with: The raster pattern of this species is fairly distinct, but superficially resembles that of Froggatt’s canegrub (Fig. 11.13d). Their distributions do not overlap. Hosts: Sugarcane. Life cycle in sugarcane: Lepidiota crinita has a typical 2-year cycle with beetles emerging after rain in November–December. Beetles do not feed and males are strongly attracted to lights. Risk period: Spring or early summer.
Life cycle in sugarcane: Lepidiota caudata has a 2-year life cycle, but it develops faster than the typical 2-year cycle pattern. Damage is a combination of 1-year and 2-year symptoms. Adults usually fly following rain in October– November. Grubs become third-instar by March, when severe damage is first seen. Third-instar grubs feed until December or later, although most are fully fed by then. Risk period: The first and most severe damage occurs from mid-summer to autumn and then continues the following spring.
Childers canegrub Antitrogus parvulus Britton. Formerly referred to as Pseudholophylla furfuracea (Burmeister)
Damage: Bundaberg canegrubs cause typical 2-year type damage in spring or early summer. Damage commonly occurs in young ratoon cane, but also in young re-plant and fallow-plant crops. Larvae feed on the roots causing poor growth and sometimes death of plants.
Distribution: Southern Qld. It is common around Childers and also at Woongarra, Bingera and Gin Gin. Grubs are found mainly in red volcanic clays or clay loams, but low numbers can be found in heavy alluvial soils. Childers grubs can occur with Bundaberg and negatoria canegrubs on light clay loams.
Caudata canegrub
Pest status: Major, restricted, irregular.
Lepidiota caudata Blackburn Distribution: North Qld in both alluvial and red volcanic clay loam soils. It lives near rainforest close to food trees. Pest status: Minor, restricted, irregular. Identification: The hair pattern of caudata canegrubs is pear-shaped. It usually has three to four rows of 27–44 hairs on each side. Each side meets at the base. Beetles have shiny brown backs with very small scales. There is an almost bare area in the middle of the ventral abdominal segments. They
Identification: Beetles are yellow–brown to nearly black and 18–23 mm long (Fig. 11.14). May be confused with: Larval raster pattern of Childers canegrub (Fig. 11.13b) is similar to that of noxia (Fig. 11.13j), negatoria (Fig. 11.13i) and French’s canegrubs. Hosts: Sugarcane. Life cycle in sugarcane: Childers canegrubs have a typical 2-year cycle, with beetles usually emerging in November–December. Eggs hatch by January, and grubs are small sized until March–April. Second-instar grubs may usually be found until September. Third-instar grubs
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may develop as early as July, but some develop as late as December, often under green cane trash blanketing. Second- and third-instar grubs can be common in the upper soil during winter. Female adults are poor fliers and usually mate on the ground near where they emerge; hence, infestations often spread out from a central point. Risk period: Childers canegrubs cause typical 2-year type damage with feeding by third-instar grubs during winter, spring and summer.
Consobrina canegrub
Beetles fly and mate after rain from September to January. One-year types at Mossman tend to fly early (September–October), while two-year types fly later (November–January). Feeding, mating and egg-laying is similar to French’s canegrub. Grub development of the 1-year type north of Cairns is similar to that of greyback canegrub. Grub development of the 2-year type south of Cairns is similar to that of French’s canegrub. Risk period: Damage symptoms are typical 1-year or 2-year type, depending on the race present. The risk period is mid-summer to autumn or in spring.
Lepidiota consobrina Girault Distribution: A 1-year life cycle type of this species is the most common from Mossman to Cairns, while a 2-year type of the species occurs in pockets between Cairns and Gordonvale (far north Qld), where mixed groups occur with French’s canegrub. Most infestations occur in dark-coloured sandy loam soils. Pest status: Minor, restricted, irregular. Identification: The raster pattern has about 50 hairs arranged in four or more rows on each side with a clear path between the two. The pattern is pear-shaped, similar to that of French’s canegrub, except that it is slightly more angular towards the posterior end and tapers to a distinct point at the anterior end where there are two single rows of hairs (Fig. 11.13c). Beetles are dark brown with oval white scales scattered evenly across the back. They are slightly larger than French’s beetles. May be confused with: Consobrina larvae are difficult to distinguish from French’s and caudata grubs by details of their raster. In the case of the adult, more than half the length of each ventral segment on the abdominal surface is devoid of scales in consobrina adults, while on French’s beetles these bare areas appear as narrow bands less than half the length of each body segment. Hosts: Sugarcane. Life cycle in sugarcane: This species has two distinct races that have different life cycles and cause typical 1-year-type damage in autumn or 2-year-type damage in spring. At Behana, near Gordonvale (far north Qld), a mixture of both races occur.
French’s canegrub Lepidiota frenchi Blackburn Distribution: The most widespread canegrub in northern districts of Qld and at Mackay, but it is a minor species at Bundaberg. It commonly infests sandy loam soils, but also occurs in red volcanic loams, red schist soils and some alluvial loams in areas which once grew open eucalypt woodland. Pest status: Major, widespread, irregular. Identification: Lepidiota frenchi has a pearshaped hair pattern, pointed at the bottom and with a clear path between the rows. Each side has about 50 dark hairs in three or four loosely arranged rows. Beetles are about 25 mm long, dark brown with rounded white scales on the back. There is a dense mass of white hairs on the underside around the bases of the legs. May be confused with: Lepidiota frenchi can be confused with caudata and consobrina (Fig. 11.13c) canegrubs in the north, negatoria canegrub (Fig. 11.13i) at Mackay, noxia (Fig. 11.13j), negatoria and Childers canegrubs (Fig. 11.13b) at Bundaberg. Hosts: Sugarcane and pastures. Life cycle in sugarcane: Lepidiota frenchi typically has a 2-year life cycle. Beetles emerge at dusk after early summer rain, usually between November and January. After a short flight over cane, female beetles land on low objects and plants, and mate. Beetles then fly to feeding trees, such as Moreton Bay ash, bloodwoods and eucalypts, but also can eat leaves of other plants including sugarcane. Feeding trees are not 327
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needed as beetles can eat leaves of other plants like sugarcane. Beetles return to a suitable site and burrow into the soil at dawn. After about 1 week, females lay a loose batch of about 30 eggs 17–25 cm deep in the soil. Eggs hatch in 16 days. In good conditions, beetles survive for up to 8 weeks after they emerge and females can lay smaller batches of eggs. Young larvae feed on organic matter and cane roots for up to 9 weeks. Medium-sized grubs (second instars) feed mainly on living roots. Initially, second-instar grubs are scattered throughout the root zone, but then concentrate about the stool during late summer and autumn, removing enough roots to loosen the stool but not causing obvious damage. During May or June, second-instar grubs stop feeding and move down into the subsoil 30–60 cm depending on soil type. Here they construct a small chamber, become yellow and overwinter. In spring, grubs change into the third instar and move towards the surface where they start feeding. Grubs are normally fully fed by March. During April to June, fully fed grubs (third instars) burrow 25–60 cm deep and form chambers in which they prepare to pupate. Pupation usually occurs in October and the pupal stage lasts 1 month. Fully grown beetles remain in the chambers until suitable rainfall and temperature triggers emergence. This completes a full 2-year life cycle. Risk period: Late spring–summer.
Froggatt’s canegrub Lepidiota froggatti Macleay Distribution: Far north Qld, in or near rainforest. Pest status: Insignificant, restricted, irregular. Identification: The raster pattern of Froggatt’s canegrub has 12–23 thick hairs on each side which fan out toward the posterior end. Hairs are arranged in single rows at the bottom end with a second, inner row of slightly smaller hairs at the top (Fig. 11.13d). Grubs are very large with shiny, dark brown heads. Beetles are 30–38 mm long and are a dull felted brown colour due to the body covering of yellow–brown hairs. May be confused with: Picticollis and rhopaea grubs.
Life cycle in sugarcane: Appears to be a 2-year life cycle, as it causes typical 2-year type damage to ratoon cane in spring. Beetles fly after rain in October and November, usually at the same time as caudata and greyback beetles. Following a flight in October, well-grown small grubs were found in plant cane during December. Secondinstar grubs were present in April, and partially fed third-instar grubs were found in ratoons during October. Fully grown grubs are two to three times the size of a greyback grub. A single grub can remove all roots from a single plant and severely damage the underground stems as well. Risk period: Spring.
Grata canegrub Lepidiota grata Blackburn Distribution: Lepidiota grata occurs throughout Qld, mostly in light sandy soils. Damage is most common in the Central, Burdekin and Herbert areas. Pest status: Minor, widespread, irregular. Identification: The raster pattern of grata canegrubs has two slightly curved sides, each with a single row of 18–26 short, thick hairs (Fig. 11.13e). Grubs are small. The number of hairs on each side will tell them apart. Greyback canegrubs normally have 20–28 hairs in each row. Beetles are fairly small, being 20 mm long. Backs are dark brown and are evenly covered with round white scales. May be confused with: Grubs are often confused with small greyback canegrubs because of similar hair patterns. Hosts: Sugarcane, pineapples and turf. Life cycle in sugarcane: Both 1-year and 2-year types. Beetles fly after rain in November and December. With good crop growth, over 90% of grubs reach third instar by April and they feed until June. Most grubs stop feeding in winter but some may still be feeding until August. Fully fed grubs pupate in October and beetles emerge in November, completing a 1-year cycle. Remaining grubs and those from small, droughted or grub-damaged cane develop more slowly and complete a 2-year cycle. Maturing crops are often stunted, but damage rarely causes lodging.
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Risk period: Plant cane may be damaged but damage is more likely in ratoon crops. Ratoons fail in spring if grubs have a 2-year feeding habit. With a 1-year cycle, ratoons grow vigorously in spring and early summer but further growth stops in April when the next generation of grubs reaches third instar.
Greyback canegrub Dermolepida albohirtum (Waterhouse) Distribution: Found in all northern and central districts of Qld. Pest status: Major, widespread, regular. Identification: The raster pattern of greyback canegrub larvae has two almost straight sides, each with a single row of 20–28 short hairs. Sides curve slightly inward at both ends and have a naked path between the (Fig. 11.13f). Beetles are large (35 mm) and coloured grey by a coating of body hairs (Fig. 11.15). Beetles develop dark brown patches as hairs wear away. The abdomen has a central, bare, dark area. May be confused with: Larvae are easily separated from most species by the raster pattern. First and second instars may be confused with grata canegrub. Hosts: Larvae attack sugarcane, corn, sorghum, bananas, peanuts (Fabaceae), pawpaw (Asimina triloba), Guinea grass and blady grass (Imperata cylindrica). Adults aggregate on a range of trees such as figs, wattle and palms and feed on the leaves. Life cycle in sugarcane: Greyback canegrubs have a 1-year life cycle. Beetles emerge after rain between October and February. Beetles mate
and feed on trees, particularly figs, wattles, palms, some eucalypts and bananas. Egg-laying starts after 10–14 days. About 28 oval, creamcoloured eggs 3–4 mm in diameter are laid in a single chamber 22–45 cm deep in the soil. Beetles can lay up to three clutches of eggs, and eggs hatch after 2 weeks. Hatchlings feed on fine roots and organic matter for 4 weeks. Second instars aggregate under cane stools and feed on roots for 5 weeks. Third instars feed on the roots and stool and grow rapidly in the first 2 months when they cause most damage. After 3–4 months, fully fed grubs burrow down and form chambers. Pupae are formed between July and October, and beetles develop within 1 month. Beetles remain in chambers until rainfall triggers emergence. Risk period: March–June; older ratoons are generally more prone to damage. Damage: Greyback canegrubs cause 1-year type damage between February and June. Infestations are common on loam soils along river systems in a wide range of soil types. Root pruning causes plants to fall, and stools to pull free of the soil, preventing further ratooning. In less severe cases, the crop remains upright but grows poorly and the leaves yellow. However, stools are pulled from the soil during harvesting resulting in gappy ratooning.
Grisea canegrub Lepidiota grisea Britton Distribution: Qld, mainly found at Yule Point near Mossman and Behana near Gordonvale. The Grisea canegrub is thought to damage cane at Ayr and Ingham. It seems to prefer sandy soils, and usually occurs with other damaging species such as sororia and squamulata canegrubs. Pest status: Minor, restricted, irregular. Identification: Grisea canegrubs have a hair pattern of 26–34 short, thick, dark hairs arranged in single straight rows (Fig. 11.13g). Beetles are 22–27 mm long with large white scales over the entire back. They are very similar to squamulata beetles.
Figure 11.15. Greyback canegrub adult, 35 mm long. (Graphic Science: © Denis Crawford)
May be confused with: Grisea can be confused with greyback and squamulata canegrubs, both of which have 1-year life cycles and occur in the 329
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same areas. Grisea canegrubs are of a similar size to French’s and squamulata canegrubs.
Distribution: Negatoria canegrubs occur from Proserpine in Qld south to NSW.
Hosts: Sugarcane and pasture.
Pest status: Major, restricted, irregular.
Life cycle in sugarcane: Grisea canegrub has a 1-year life cycle. Little damage has been noticed due to this species.
Identification: Lepidiota negatoria larvae have the same hair pattern as Lepidiota frenchi. Beetles are dark red–brown with large white scales on both upper and lower surfaces (Fig. 11.13i).
Risk period: March–April.
Nambour canegrub Antitrogus rugulosus (Blackburn) = southern populations of Antitrogus mussoni (Blackburn) = Rhopaea rugulosa Blackburn Distribution: Nambour canegrub occurs at Didillibah and Bli Bli in the Moreton district, at Rocky Point Qld, and in northern NSW, usually in sandy soils. Pest status: Minor, restricted, irregular. Identification: Larvae have a hair pattern of two single convex rows which come together at each end. There are 19–31 short, thick hairs on each side, but most commonly 23–26 (Fig. 11.13h). Grubs look similar to southern one-year canegrub. They sometimes occur with negatoria canegrubs, but are slightly smaller and have a different hair pattern. Beetles are bright red–brown and without scales, but with short hairs on their back. May be confused with: Southern one-year grub. Also with grata and Rothe’s, but separated geographically from those two species.
May be confused with: Negatoria canegrubs have a pear-shaped raster pattern similar to that of French’s and noxia canegrubs. Adult males can be distinguished by means of the shape of the aedeagus. Hosts: Sugarcane. Life cycle in sugarcane: Negatoria canegrubs have a typical 2-year life cycle with beetles emerging after heavy rain, usually in November–December. Males and females both feed on the foliage of trees and cane, and are attracted to light. Lepidiota negatoria is one of the main grub species in Bundaberg, and is one of only two of importance at Maryborough. Grubs are found in forest loam and light clay soils. Negatoria canegrubs cause typical 2-year-type damage in spring or early summer. Damage most commonly occurs in young ratoon cane, but also in young re-plant and fallow-plant crops. Risk period: Spring–summer.
Noxia canegrub
Hosts: Sugarcane.
Lepidiota noxia Britton
Life cycle in sugarcane: Nambour canegrubs have a 1-year life cycle with non-feeding beetles usually emerging in October–November following rain. Some beetles emerge as late as February in dry years. Third-instar grubs move into the rows in mid-summer and feed through autumn, winter and early spring. Late springfeeders may be the offspring of late-flying beetles and could take 2 years to complete their life cycle.
Distribution: Noxia canegrubs occur in Qld north of the Burnett River in the Bundaberg area, in red and brown forest loams and scrub soils at Gooburrum, Sharon, Bingera and Yandaran and in coarse alluvial soils at Waterloo. They also occur at Nikenbah and Pialba at Hervey Bay.
Risk period: Autumn–early winter.
Identification: Noxia canegrubs have a hair pattern similar to those of negatoria and French’s canegrubs and often occur with those species (Fig. 11.13j). Beetles are dark red–brown with a few oval white scales.
Negatoria canegrub Lepidiota negatoria Blackburn = Lepidiota deceptrix Blackburn = Lepidiota mungomeryi Britton
Pest status: Minor, restricted, irregular.
May be confused with: Negatoria and French’s canegrubs.
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Hosts: Sugarcane.
Planiceps canegrub
Life cycle in sugarcane: Noxia canegrubs have a 2-year cycle with beetles emerging in September–October following storm rain. Grub growth is complete by the end of the second autumn and beetles remain in the pupal cells until spring rains.
Antitrogus planiceps (Blackburn)
Risk period: Summer–early autumn.
Picticollis canegrub Lepidiota picticollis Lea Distribution: In Qld, picticollis canegrubs damage sugarcane in the Bundaberg area at Yandaran, Elliott Heads and Kolan, and in the Isis area at Logging Creek and Goodwood. They always occur in very sandy soils. Pest status: Minor, restricted, irregular. Identification: Picticollis canegrubs have a hair pattern of two parallel rows, each with 29–40 short thick dark hairs (Fig. 11.13k). Some grubs have a short second row of a few hairs at the posterior end. They are the largest grub found in southern cane growing areas, and they sometimes occur with southern one-year canegrubs.
Distribution: Damage reported from Harwood and Broadwater districts of NSW. Pest status: Minor, restricted, irregular. Identification: Planiceps canegrubs have a pearshaped hair pattern with two rows each with 32–39 long thin hairs. There are two rows from the posterior end to the centre and one row at the anterior end. Grubs with fewer hairs lack part of the anterior portion of the rows. Beetles are tan to black, without scales, and the male antennae have five long segments. May be confused with: Consobrina canegrub, but they have different geographical distribution. Hosts: Sugarcane. Life cycle in sugarcane: The detailed life history of this species is not known. Third-instar grubs are present in the field in early summer. Beetles fly in October–December, but autumn flights have also been recorded. Risk period: Summer.
Rhopaea canegrub
Beetles are large (range 25–32 mm) and shiny yellow–brown to deep chestnut with backs bordered in dark brown or black. Most have an orange–red patch on either side of the body behind the head.
Rhopaea magnicornis Blackburn
May be confused with: Rhopaea grub. Also the raster pattern is similar to picticollis grub, but the two species are geographically separated.
Pest status: Minor, restricted, irregular.
Hosts: Sugarcane. Life cycle in sugarcane: Grubs take 15 months to develop, pupate late in the second summer, and spend the second autumn and winter as beetles underground. The beetles usually emerge in September–October following storm rain, but some may emerge in summer or autumn. Damage by Lepidiota picticollis is similar to damage from southern one-year canegrub. One grub per stool is capable of causing economic damage. Risk period: Late summer–early autumn.
Distribution: NSW and Southern Qld. Damage usually occurs at Nambour, Rocky Point and in northern NSW, especially in the Tweed Valley.
Identification: Rhopaea canegrubs have a hair pattern of two straight rows, each with about 20 short hairs (Fig. 11.13l). Beetles are dark brown, about 21–30 mm long, and have a coat of short, fine, semi-erect hairs. Males have large antennae with clubs made up of eight segments. May be confused with: The raster pattern may be similar to that of picticollis canegrub. Hosts: Sugarcane. Life cycle in sugarcane: Rhopaea canegrubs have a 1-year or 2-year life cycle depending on the weather, but mostly cause typical 1-year type damage. Some grubs take 2 years to develop following cool autumns, and these damage 331
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recently harvested ratoon crops. After a warm autumn, most grubs will pupate in August and emerge as beetles in late October or early November.
at the posterior end. Each side has three to four rows with about 45 long hairs (Fig. 11.13n). Beetles are relatively small (22 mm long), and light brown with white markings on the abdomen.
Risk period: Depends on the life cycle type, but generally damage is seen in autumn months.
May be confused with: Larvae could be confused with other white grubs that are not pests of sugarcane which have a similar ‘horseshoe’ raster pattern.
Rothe’s canegrub Lepidiota rothei Blackburn = Lepidiota bovilli Blackburn = Lepidiota koebeli Blackburn = Lepidiota parva Blackburn Distribution: NT and Qld. Pest status: Insignificant, restricted, irregular. Identification: Rothe’s canegrubs have a hair pattern with two single, slightly curved rows, each with 10–12 long thin hairs. The hairs almost meet across the path between the two sides (Fig. 11.13m). Beetles are smaller (range 15–19 mm) and are darker-coloured than French’s canegrub beetles, and do not have hairs on the ventral side of the abdomen. The scales on the back are rounded.
Hosts: Sugarcane. Life cycle in sugarcane: The sororia canegrub probably has a 1-year life cycle, but the details are not clear. Beetles fly as early as August, but more commonly in November and December, and often without a need for rainfall. In one year, first-, second- and third-instar grubs were present in December, with second-instar grubs being the most common. Almost all grubs were third-instar by early February. Grubs were fully fed in March and pupated in May. Sororia canegrubs cause 1-year type damage, and are often found with grata and squamulata canegrubs in damaged cane at Ingham, but, in one instance, sororia was the only species causing damage.
May be confused with: Grata and southern one-year grub.
Risk period: March–April.
Hosts: Sugarcane.
Southern one-year canegrub
Life cycle in sugarcane: This species has an unusual 1-year life cycle. Beetles fly at dusk in early summer, and mate on low trees and shrubs. Second-instar grubs are present through winter. These change into the third-instar grubs in spring, and feed until October or November. Pupation occurs in November and beetles are formed in 24 days.
Antitrogus consanguineus (Blackburn) = northern populations of Antitrogus mussoni (Blackburn) = Rhopaea consanguinea Blackburn
Risk period: Rothe’s canegrub is not usually a pest of sugarcane, but grubs are very common in weedy fields and when grassy fallow fields are ploughed for planting.
Pest status: Major, restricted, irregular.
Sororia canegrub Lepidiota sororia Moser = Lepidiota wilsoni Britton Distribution: North Qld. Pest status: Minor, restricted, irregular. Identification: Sororia canegrubs have a horseshoe-shaped hair pattern, with the opening
Distribution: Southern Qld. The southern population of the species (once confused with A. mussoni) is now known as Nambour canegrub.
Identification: Southern one-year canegrubs have a hair pattern of two single convex rows of 19–31 (commonly 23–26) short, thick hairs, coming together each end (Fig. 11.13o). The species can occur with negatoria canegrubs, but the southern one-year canegrub is smaller and has a different hair pattern. Beetles of southern one-year canegrubs are bright red–brown and without scales, but with short hairs on the back. Male antennae have 5½ large segments. The southern one-year canegrub occurs in the Bundaberg and Maryborough areas in sandy alluvial or yellow podsol soils, especially wallum country.
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May be confused with: Nambour grub, but geographically separated. Hosts: Sugarcane. Life cycle in sugarcane: Southern one-year canegrubs have a 1-year life cycle with beetles emerging in September–October following rain. Third-instar grubs are found in the stool shortly after Christmas and continue feeding until late May. Grubs then move to about 25–40 cm into the soil and pupate in late winter. Risk period: Late summer–autumn. Southern one-year canegrubs are commonly found in autumn-planted crops as well as ratoon crops.
Squamulata canegrub Lepidiota squamulata Waterhouse = Lepidiota darwini Blackburn = Lepidiota leai Blackburn = Lepidiota rugosipennis Lea Distribution: Northern Australia including WA. The insect is commonly found with other grubs, mainly grata, grisea, sororia and French’s, and it is found only in sandy soils. Pest status: Minor, restricted, irregular. Identification: Squamulata canegrubs have a hair pattern of two straight rows each with 28–40 short, thick hairs. The space between the rows widens at the anterior end (Fig. 11.13p). Beetles are 22–32 mm long and slightly larger than French’s canegrub. They are dark-coloured with oval white scales along the sides and on the abdomen. May be confused with: Squamulata canegrub can be confused with grisea and greyback canegrubs. Hosts: Sugarcane, pineapples, also common as lawn pests in the north. Life cycle in sugarcane: The squamulata canegrub has a typical 1-year life cycle. Beetles fly after heavy rains in November–January and feed on eucalyptus leaves before laying. Eggs hatch in 15 days. Grubs grow from small to second instar in a total of 8–12 weeks. Third-instar grubs occur between February and May and are fully fed and ready to pupate as early as May. All grubs pupate by October, and beetles are present in soil chambers from October onwards. Risk period: Autumn.
Leaf beetles Coleoptera: Chrysomelidae Black leaf beetle, Rhyparida morosa Jacoby = Rhyparida nitida Clark Sugarcane leaf beetle, Rhyparida dimidiata Baly Distribution: Both species are indigenous to Australia, recorded from NSW and Qld. Pest status: Minor, restricted, irregular. Identification: Larvae of both species have a yellow–grey body and a shiny red–brown head. They are about 9 mm long when fully grown. Rhyparida morosa adults are about 6 mm long, dark shining black, and have a rounded outline, while Rhyparida dimidiata adults are 6–7 mm long, brown coloured with straight sides. May be confused with: Adult beetles can be confused with other types of chrysomelid and scarabaeid beetles associated with cane, but can be easily separated based on the morphology. Damage may be confused with that caused by the ratoon shootborer, large mothborer and wireworms. Hosts: Sugarcane. Life cycle in sugarcane: Larvae are active in spring and change to adults by December– February. Rhyparida morosa adults frequent taller grasses and are often found on cane leaves in large numbers, while Rhyparida dimidiata adults often shelter on or under the bark of Moreton Bay ash or blue gum. Risk period: Spring–early summer. Damage: In September–November small grubs bore into the bases of shoots causing ‘dead hearts’. Damage is common in ratoons but can occur in plant cane. Heavy attack may kill all shoots and also the stool. A less severe attack may only weaken the stool and kill only a few shoots. Damage more often occurs in grassy fields. Adult beetles eat sugarcane leaves giving them a tattered appearance, but cause no economic damage. Monitoring: Plants can be dissected to inspect the presence of larvae at the base of the shoots. Action level: By the time damage is seen it is too late to attempt control. 333
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Chemical control: No insecticides are known to control these pests. Cultural control: Avoid planting susceptible varieties. Natural enemies: Unknown.
Sugarcane butt weevil Leptopius maleficus Lea Coleoptera: Curculionidae Distribution: Native to Australia. Damage occurs only in far north Qld. Pest status: Minor, restricted, irregular. Identification: Larvae are legless and slightly curled with small heads. Their bodies taper slightly towards the rear end. They are cream– yellow with a pale head and black mouth parts, and are 20–24 mm long when fully grown. Larvae are found in the soil or boring in setts, stubble or shoots (Fig. 11.16). Adult weevils are grey or reddish with many rounded lumps on their bodies. The mouthparts are on a long snout. The body is much broader at the rear end of the wing covers than at the front end. Weevils are 16–21 mm long, and males are smaller than females. May be confused with: The larvae can be confused with larvae of weevil borers, but the latter have a red–brown head and an abdomen which widens towards the rear and then ends in a sharp point. Hosts: Adult weevils feed on common sida (Sida rhombifolia), pink flannel burr and senna leaves, but do not usually feed on cane leaves.
Life cycle in sugarcane: The larval stage lasts several years. Large grubs have been kept in the laboratory for at least 1 year with little change in body size. Large grubs leave plants and move into the soil by January, probably to pupate. In infested fields, large numbers of weevils usually emerge in spring of the year following damage to cane. Weevils are found in cane fields between August and March, usually feeding and mating on leaves of both young and mature rattlepod (Crotalaria spp.) Up to 45 mature eggs are ready to be laid by February. Risk period: Damage to cane is seen between June and December. Damage: Damage has been recorded in alluvial loam at Mundoo, in red volcanic loam at Mena Creek, in red schist loam at Moresby, and dark sandy loam at Kurrimine, far north Qld. Both plant and ratoon canes are damaged. Grubs hollow out ratoon stubble and setts, make long channels into and destroy the rind, and eat eyes and root buds. They also tunnel into and make large holes in the base of new shoots, causing ‘dead hearts’ and wilted shoots (Fig. 11.18). Moderately damaged shoots can recover if growing conditions are good. Monitoring: Plants can be dissected to inspect the presence of larvae at the base of the shoots. Action level: Butt weevil damage is rare and of minor importance. Chemical control: No chemicals are registered for butt weevils. Cultural control: Control of weeds that weevils feed on will reduce insect numbers in cane. Natural enemies: Unknown.
Sugarcane weevil borer (New Guinea weevil borer) Rhabdoscelus obscurus (Boisduval) Coleoptera: Curculionidae
Fig. 11.16. Sugarcane butt weevil larva (24 mm extended length) with damage to the butt of a cane. (BSES)
Distribution: Native to Papua New Guinea. Introduced to Australia, Hawaii, Fiji and other Pacific Ocean islands, although the population in Hawaii may be a sibling species (Giblin-Davis et al. 2000). Pest status: Minor, restricted, irregular.
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Identification: Larvae are legless and cream with an oval red–brown head and red–brown mouth parts. The tail bells-out and then comes to a point, and each larva lives in its own tunnel. Fine sawdust-like frass is packed tightly into the tunnel behind the larva. Pupae are wrapped in tightly bound fibre cocoons. Adult weevils are dark-coloured, 12–15 mm long with distinctly curved rostrum and red patterns on each wing cover. May be confused with: Adults can be confused with other weevils associated with sugarcane. The banana weevil borer, Cosmopolites sordidus (Germar), is very similar to this species, but its wings lack the red colouration. The larvae may be confused with butt weevil larvae, which damage cane around Innisfail (far north Qld), but they have a cream–yellow head and their body tapers slightly towards the rear end of the body, unlike weevil borer larvae that have a larger, red–brown head and a posteriorly enlarged abdomen tapering sharply to a point. Hosts: Larvae attack palm trees (Arecaceae), as well as sugarcane. The pest is common in palm nurseries and has been collected in palms transported to weevil-free areas such as Bundaberg. Life cycle in sugarcane: Adult weevils start appearing in cane fields by October–November. They chew a small cavity into the rind or split tissue and lay a single, long, curved egg into each hole. Larvae hatch and burrow into the inner tissue, and are fully grown in about 10 weeks. They then chew small exit holes in the rind as an escape route for the adult. They pupate in fibre cocoons and emerge as adults in about 9 days. Under good conditions, weevils complete four generations a year, and adults can survive for 6 months. Weevil borers can infest young ratoons by boring down into the stool if there is no other food around. Billets or whole stalks left after harvest allow borers to breed and to re-infest young cane in February and March. Risk period: Infestation can start as early as December but damage is seen later in March– April until harvest. Damage: Rhabdoscelus obscurus mainly lowers the Cane Commercial Sugar (CCS) value. Heavily bored crops have lost up to two units of CCS value. The larvae bore into and tunnel through
the soft tissue inside mature and semi-mature stalks leading to red rot and other diseases which reduce sugar content. Damaged stalks also have higher fibre content and higher dextran levels in juice. Damage is generally restricted to the bottom sections, which are attacked when the stem is young and soft, but stems which have been split and twisted in cyclonic wind or those that have been attacked by rats are prone to heavy damage. Weevils may also enter underground parts of stems. Although the borer is common north of the Herbert Valley, heavy damage is found on the drier soils in the super-wet belt from Tully to Cairns (far north Qld). Recently, damage has been reported from central Qld. Monitoring: Split cane traps are a cheap method to monitor borers, where pieces of split cane are wrapped in black plastic and placed under cane plants at the rate of eight traps per hectare. Pheromone trapping is also available, and the use of water pheromone traps have been successful in far north Qld. Damage is best monitored through sampling 175 cane stalks randomly, splitting the stalks and counting the number of tunnels per internode (Sallam et al. 2001). Action level: Damage is difficult to predict based on adult monitoring, but an average damage level of 0.3–0.4 internodes per stalk will cause significant damage in susceptible varieties (Sallam et al. 2004). Chemical control: Chemical control is available and should be applied as cane forms its first millable internode. Cultural control: Area-wide integrated pest management is necessary to control the pest, using tolerant varieties, trash management, farm hygiene and pheromone mass-trapping of adults (Robertson and Webster 1995; Sallam et al. 2001). Natural enemies: The fly Lixophaga sphenophori (Villeneuve) (Diptera: Tachinidae) was introduced from New Guinea in 1911. It has established in Australia, but is not economically effective.
Whitefringed weevil Naupactus leucoloma (Boheman) = Graphognathus leucoloma (Boheman) Coleoptera: Curculionidae 335
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N (Main entry in Chapter 15 Pastures—winter rainfall.)
Pest status: Minor, widespread, irregular.
Pest status: Minor, restricted, irregular.
Identification: Fully grown larvae are 35–40 mm long with a purple or pink tint along the back, and small black spots along the body. The head is red-tanned (Fig. 9.24).
May be confused with: Adults may occur with other weevils associated with sugarcane. Larvae may be confused with those of weevil borers, but the latter have a red–brown head and an abdomen which widens towards the rear, then ends in a sharp point; also weevil borer larvae do not live in the soil. Life cycle in sugarcane: All whitefringed weevils in Australia are females. Adults are present throughout summer and autumn. Eggs are laid during this period in batches of 12–60 attached to plant stems, dead leaves or stones. Each female can lay up to 1500 eggs. Eggs hatch in 2–4 weeks in summer and autumn but take up to 3 months in winter. Grubs take 6–18 months to develop depending on food supply. Adults emerge from the pupal chamber after rain. Adults cannot fly, but can move easily into nearby crops. Insects rarely persist for more than one generation in weedfree cane fields. Risk period: Spring. Damage: Adult weevils chew the leaves of cane but rarely cause any economic damage. The larvae attack the roots of germinating setts or ratoons, weakening or killing the plants. Damage often follows legume cover crops or fallows with large numbers of legume weeds. Monitoring: Plants can be dissected to inspect the presence of larvae at the base of the shoots. Action level: Damage by whitefringed weevil is rarely of economic importance. Chemical control: No insecticide is registered for use against this pest in sugarcane. Cultural control: Removal of leguminous weeds assists in reducing numbers.
Sugarcane and maize stemborer (large mothborer) Bathytricha truncata (Walker) Lepidoptera: Noctuidae Distribution: Native to Australia, recorded from NSW, Qld and Tas.
Pupae are about 19 mm long. They are found in tunnels in large cane, and in the trash or behind leaf sheaths in young cane. Moths are dull-coloured with small dots along outer edges of wings and in a group in the middle of the wing. May be confused with: ‘Dead heart’ symptoms maybe confused with those caused by wireworms or black beetles. Weakened young cane often breaks in strong wind, and this may be mistaken for climbing rat damage. Feeding symptoms maybe confused with those caused by weevil borers, but tunnels are filled with coarse, loose, wet frass, often foul-smelling, which is different from the fine, fairly dry and tightly packed frass left in tunnels by weevil borers. Hosts: Sugarcane, maize, rice (Oryza spp.) and wheat (Triticum aestivum). It also attacks a wide range of wild grasses such as Paspalum spp., crowsfoot grass (Poaceae), Rhodes grass (Chloris gayana), Natal grass (Rhynchelytrum repens) and Guinea grass (Panicum maximum). Life cycle in sugarcane: Eggs are laid in small clusters during spring months under the edge of leaf sheaths of grasses and young sugarcane shoots. Crowsfoot grass, guinea grass, red natal grass and rhodes grass are preferred as egg-laying sites over sugarcane shoots. Moths lay 500–800 eggs, and eggs hatch in 8 days. Young larvae feed inside the leaf sheath for several days before boring through the rind of the plant, then move from one shoot or stem to another as the tissues begin to rot, damaging several shoots and stems during their life. Young larvae can bore directly into grass and young sugarcane shoots, but not into stems of large sugarcane. Larvae go through five or six growth stages which take about 6 weeks in total. The pupal stage lasts for 12 days in summer, and moths continue breeding until February. Risk period: Spring–summer.
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Damage: The large mothborer damages cane in all sugarcane regions. It usually infests cane in fairly dry, well-drained areas close to grass. The larvae bore into the growing points of both small and large cane causing ‘dead hearts’. They also tunnel inside young internodes, often completely circling the rind at each notch, promoting stalk rots which reduce sugar content. Bored stalks sucker and side-shoot, and heavily infested fields sometimes fail to recover, resulting in gappy plant or ratoon crops. Monitoring: Dead heart symptoms can be recognised during autumn to early summer. Larvae can be found in the growing point by dissecting the plant. Action level: No economic injury level studies are available for this pest. Chemical control: Bathytricha truncata is a fairly minor pest and no chemical control is required. Cultural control: Managing grass weeds, which are egg-laying sites, and food for young larvae helps reduce the damage. In northern regions, the damage can be significant in fields where couch grass is a weed and in blocks with overgrown grassy headlands. Some cultivars are also less susceptible to damage. Natural enemies: Larvae and pupae are attacked by at least four small wasps and a fly. Cotesia sp. (Hymenoptera: Braconidae) the most active parasitoid (Fig. 11.17), is the only species identified. Coastal brown ant, Pheidole
megocephala (Fabricius, Hymenoptera: Formicidae) eats larvae.
Armyworms and loopers Lepidoptera: Noctuidae Nightfeeding sugarcane armyworm, Leucania loreyi (Duponchel) Sugarcane armyworm, Leucania stenographa Lower Leucania loreyi is an exotic species found in Europe, Africa and Asia. Distribution in Australia is not well studied, but it possibly occurs Australia wide. Leucania stenographa is a synonym of Leucania loreyimima Rungs, which is known throughout mainland Australia (Edwards 1992).
Common armyworm, Leucania convecta (Walker) Leucania convecta (previously assigned to the genus Mythimna) is native to Australia and widely distributed in the continent.
Northern armyworm, Leucania separata (Walker) Mythimna separata is native to Australia but distributed in Asia, West Africa and New Zealand.
Day-feeding armyworm, Spodoptera exempta (Walker) Spodoptera exempta probably originated in Africa, but currently occurs in North America, Australia and New Zealand. In Australia, it occurs throughout the mainland.
Sugarcane looper, Mocis frugalis (Fabricius) Mocis frugalis is native to Australia and occurs in NSW, NT and Qld. An old world species from Africa and Asia was formerly considered one with this species, but was recently separated and designated as Mocis proverai (Zilli 2000). Pest status: All are minor pests, but can be important in restricted areas, widespread, irregular. All species occur in all sugarproducing regions of Australia. Fig. 11.17. Cotesia sp. parasitoids on larva of sugarcane and maize stemborer (caterpillar is about 35 mm long). (BSES; UnivAd)
Identification: Night-feeding armyworms: larvae of night-feeding armyworms hide in rolled
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spindle leaves or under leaf litter during the day, and feed mostly at night. Pupae form under trash or 1 cm underground in small chambers. Adults of the night-feeding armyworms, Leucania loreyi (Fig. 2.40), Leucania convecta (Fig. 2.32) and Leucania separata (Fig. 2.38) all have uniform pale yellow-coloured forewings with obvious veins. There is a line of small dark specks on the outer part of the wing and a single white speck near the centre. Hindwings of Leucania loreyi are mostly white, whereas Mythimna convecta and Mythimna separata have a large dark area on the outer half of the hindwing. Day-feeding armyworms: young larvae are bright green with tan heads. When about 20 mm long, they have dark green, almost black, stripes down the body. On the front of the head is an upside down Y-shaped white line. Fully-grown larvae are 25–35 mm long. Larvae move about during daylight and feed on leaves. Pupae are found in small chambers just under the soil surface. The moth is small, about 14 mm long, and has darkcoloured forewings with small white lines in the central area. The hindwings are pale with a light pink tint and a dark edge. Sugarcane loopers: Larvae are long, thin, green or yellow–brown with two black bands across the front part of their body. They have three pairs of stubby legs at the rear end. Loopers move in a different way to other larvae. They grab on with their front legs, let go with their hind legs and slide their rear section forward. Larvae make a tube-like cocoon by rolling the opposite edges of a leaf tip together. The moth is pale yellow with a dark stripe running at an angle down the forewing.
Life cycle in sugarcane: Night-feeding armyworms: rotting plant matter attracts moths to lay eggs. Flooded fields with silt-covered leaf litter and newly harvested ratoons with a fermentation smell are often targets. Larvae feed on grassy weeds and cane. In a field, there is usually only one major infestation per season. Sometimes a second wave of attack can follow the first, but this is rare. Day-feeding armyworms: larvae feed day and night and move as a band from one food source to another. Between December and February, large numbers start off in grass, weedy cane fields, lawns, headlands and creek banks, before moving into cane. It takes 4 weeks to go from egg to caterpillar to moth in mid-summer. Sugarcane loopers: larvae start off in grassinfested cane or in grass-infested legume cover crops, and move into cane. Risk period: Spring–summer. Damage: Armyworm larvae eat cane leaves and in cases of heavy infestation leave only a bare leaf midrib (Fig. 11.18). Monitoring: Larvae can be found under trash, or hiding at the base of the leaf. Inspection of symptoms offers a good monitoring option. Action level: Spraying may be cost-effective at most larval densities in Southern Qld (Liu and Allsopp 1996) but not in far north Qld, where natural enemies may effectively control these species. Chemical control: Sometimes cost-effective in southern Qld.
May be confused with: Common and northern armyworms occur in similar areas and are difficult to tell apart. In mixed groups on sugarcane, the northern armyworm is found in higher numbers than the common armyworm. Larvae range from pale olive-green to red– brown with stripes of white, red-brown and black running down the body. The head is a mottled tan without any white markings. Fullygrown larvae are over 40 mm long. Hosts: Armyworms feed on a wide range of gramineous plants and field crops.
Fig. 11.18. Armyworm defoliation of young sugar plants, allowing weed growth. (BSES)
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Cultural control: Some cane cultivars are less susceptible than others. Natural enemies: A number of flies and wasp parasitoids attack the larval and pupal stages of armyworms. Larvae are also attacked by several viral and bacterial diseases. The abundance of natural enemies in far north Qld is thought to be responsible for minimising the pests’ impact on the crop.
Funnel ant Aphaenogaster pythia Forel Hymenoptera: Formicidae Distribution: Native to Qld. Pest status: Minor, restricted, irregular. Identification: Worker ants are honey-coloured and about 5 mm long. Sexual stages are darkercoloured and have wings; males are 5.5 mm long and females 11 mm long. Funnel ants have two backward pointing spines in the middle of their backs. This sets them apart from other yellow ants. Workers are found by digging into the mounds, but are hard to find during the day. Funnel ants form mounds that are up to 25 cm wide and 20 cm high, with cone-shaped sides and a funnel-shaped opening at the top (Fig. 11.19). May be confused with: Workers may be confused with other ants, but mound building is fairly characteristic. Hosts: Sugarcane, blady grass (Imperata cylindrica). Funnel ants naturally live in areas of wet eucalypt forest near rainforests. They are probably omnivorous, feeding on bodies of dead insects and honeydew secretions. Life cycle in sugarcane: Sexually mature ants swarm from nests during November. Mating takes place in the air, and females return to the ground to start new colonies. It takes 49–60 days to make new workers. Tunnels 7 mm in diameter are dug under the stool and into the subsoil. In deep sandy subsoils, tunnels can be deeper than 2 m. Ants do not tunnel into clay subsoil. Oval chambers 5 cm × 2 cm are built for rearing
Fig. 11.19. Funnel ant mound. (BSES)
young. Ants live in shallow chambers in the stool zone during the wet season and deeper chambers during the dry. Risk period: Summer–early autumn. Damage: Funnel ants make small cuts in roots and eat root sap. Several species of planthoppers that share ant tunnels also feed on cane roots. Plants also suffer moisture stress in dry conditions, and in cases of heavy infestations the plants are not firmly anchored in soil and are prone to being pulled out during harvest. Funnel ants are pests of sugarcane in areas between Mossman and Tully. Monitoring: The number of mounds per metre of cane row can be counted during February– March. Action level: Up to 3500 mounds per hectare will not reduce yield (about one mound per 2 m of cane row), but a mound density of more than 4500 per hectare is likely to damage susceptible varieties. Chemical control: No chemical control is registered for funnel ants in sugarcane. Cultural control: Some varieties suffer less damage from funnel ants than others. Trials in Tully on alluvial soils found Q124 and Q138 to be less affected than varieties like Q107. Factors other than ant activity may initiate poor growth conditions under which ant populations thrive. Natural enemies: Unknown.
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Sources of information Allsopp, P.G. (1990). Use of weather data to forecast outbreaks of soldier fly, Inopus spp. (Diptera: Stratiomyidae) in Queensland sugarcane. Agriculture, Ecosystem and Environment 30: 61–70. Allsopp, P.G., Chandler, K.J., Samson, P.R., and Story, P.G. (1993). Pests of Australian Sugarcane. Bureau of Sugar Experiment Stations, Brisbane. 90 pp. Allsopp, P.G. and Robertson, L.N. (1988). Biology, ecology and control of soldier flies Inopus spp. (Diptera: Stratiomyidae ): a review. Australian Journal of Zoology 36: 627–48. Baker, G.L. (1983). Parasites of locusts and grasshoppers. Agfacts AE 2, 16 pp. Beard, J.J., Walter, D.E. and Allsopp, P.G. (2003). Spider mites of sugarcane in Australia: a review of the grass-feeding Oligonychus Berlese (Acari: Prostigmata: Tetranychidae). Australian Journal of Entomology 42: 51–78. BSES (2003). Mill Area Statistics—2002 season. BSES Publication ST03001. Chandler, K.J. (1982). Cicadid biology and control in Queensland sugarcane fields. MSc Thesis, University of Queensland. Drinkwater, T.W. (1986). Ephysteris promptella (Staudinger) (Lepidoptera: Gelechiidae), a new pest of maize and grain sorghum in South Africa. Phytophylactica 18: 221. Edwards, E.D. (1992). A second sugarcane armyworm (Leucania loreyi (Duponchel)) from Australia and the identity of L. loreyimima Rungs (Lepidoptera: Noctuidae). Journal of Australian Entomological Society 31: 105–108. Giblin-Davis, R.M., Gries, R., Crespi, B., Robertson, L.N., Hara A.H., Gries G, O’ Brien C.W. and Pierce H.D. (2000). Aggregation pheromones of two geographical isolates of the New Guinea sugarcane weevil, Rhabdoscelus obscurus. Journal of Chemical Ecology 26: 2763–2780. Hunter, D.M., Milner, R.J., Scanlan, J.C. and Spurgin, P.A. (1999). Aerial treatment of the migratory locust, Locusta migratoria (L.) (Orthoptera: Acrididae) with Metarhizium anisopliae (Deuteromycotina: Hyphomycetes) in Australia. Crop Protection 18: 699–704. Jarvis, D. (1927). Notes on insects damaging sugarcane in Queensland. BSES Entomology Bulletin 3, 94pp. Liang, A.P. and Fletcher, M. (2002). Morphology of the antennal sensilla in four Australian spittlebug species (Hemiptera: Cercopidae) with implications for phylogeny. Australian Journal of Entomology 41: 39–44. Liu, D.L. and Allsopp, P.G. (1996). QCANE and armyworms: to spray or not to spray, that is the question. Proceedings of the Australian Society of Sugarcane Technologists 18: 106–112. Magarey, R.C., Chandler, K.J. and Croft, B.J. (1988). Eoscarta carnifex (F.) (Homoptera: Cercopidae) implicated in the first Australian record of ‘froghopper blight’ in sugarcane crops. Journal of the Australian Entomological Society 27: 238. Mead, F.W. (1978). Sugarcane aphid, Melanaphis sacchari (Zehntner) -Florida- New Continental United States Record. Cooperative Plant Pest Report 3(34): 475. Robertson, L.N. (1987). The effects of natural enemies on the population dynamics of sugarcane soldier fly, Inopus rubriceps (Dip.:Stratiomyidae). Agriculture, Ecosystem and Environment 19: 343–363. Robertson, L.N. and Webster, D.E. (1995). Strategies for managing cane weevil borer. Proceedings of the Australian Society of Sugarcane Technologists 17: 90–96.
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Sallam, M.N., Garrad, S.W. and Oehlschlager, A.C. (2001). Aggregation pheromones for the management of weevil borers. Proceedings of the Australian Society of Sugarcane Technologists 23: 204–211. Sallam, M., MacAvoy, K., Puglisi, G. and Hopkins, A. (2004). Can economic injury levels be derived for sugarcane weevil borer, Rhabdoscelus obscurus (Boisduval) (Coleoptera: Curculionidae), in far-northern Queensland? Australian Journal of Entomology 43: 66–71. Samson, P.R. and Calder, A.A. (2003). Wireworm (Coleoptera: Elateridae) identity, monitoring and damage in sugarcane. Australian Journal of Entomology 42: 298–303. Samson, P.R. and Robertson, L.N. (1996). Development of a decision-support plan for wireworm management. Proceedings of the Australian Society of Sugarcane Technologists 18: 62–70. White, W.H., Reagan, T.E. and Sosa, O. Jr. (1995). The sugarcane delphacid (Homoptera: Delphacidae) extends its North American range into Louisiana. Florida Entomologist 78: 617– 619. Zilli, A. (2000). African–Arabian and Asian–Pacific ‘Mocis frugalis’: Two distinct species (Lepidoptera: Noctuidae). European Journal of Entomology 97: 419–426. Zuk, M., Rotenberry, J.T. and Simmons, L.W. (2001). Geographical variation in calling song of the field cricket Teleogryllus oceanicus: The importance of spatial scale. Journal of Evolutionary Biology 14: 731–741. Zuk, M., Simmons, L.W. and Rotenberry, J.T. (1995). Acoustically orienting parasitoids in calling and silent males of the field cricket Teleogryllus oceanicus. Ecological Entomology 20: 380–383.
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12 TOBACCO S.G. De Faveri and M.B. Malipatil N Tobacco, Nicotiana tabacum (Solanaceae). Origin: South America
(a) Transplanted seedlings (VDPI: G. Baxter)
(b) Maturing crop (VDPI: G. Baxter)
seedling transplant & growth topping & growth progressive harvest
Jul Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun Growth of summer tobacco in Vic. and southern Qld. In north Qld, tobacco was usually grown as an autumn crop, transplanted in April, with harvest during July to September. Tobacco seedlings are generally grown in a nursery before transplanting to the field. All nurseries in north Qld were outdoors. In Vic., seedlings may be produced outdoors or in greenhouses.
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PEST (major pests in bold) Pre-emergence (nursery) seedharvesting ants Seedling and growth stages two-spotted mite spur-throated locust aphids common brown leafhopper green vegetable bug thrips African black beetle false wireworms cutworms tobacco stemborer potato moth cluster caterpillar cape gooseberry budworm corn earworm native budworm looper caterpillar Post harvest tobacco beetle tobacco moth
RECORDED AS A PEST IN:
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Tobacco in Australia is currently grown in Qld and Vic. Tobacco-growing in north Qld was centred around Mareeba and Dimbulah but by 2004 production had ceased, leaving a small growing area in southern Qld. The main production area is now in Vic., around the town of Myrtleford. Leaf quality is critical to commercial tobacco production. Leaf quality may be directly affected by insects or indirectly by the diseases they vector. A number of pests are common to both Qld and Vic., but others are pests only in one state.
Risk period: Seed beds are at risk at sowing and germination. Damage: Seedharvesting ants cause damage by carrying surface-sown seed to their granaries, while some species cut and remove the leaves of seedlings soon after germination. Monitoring: These ants produce mounds which indicate the likelihood of seed loss. Chemical control in Qld: Spraying mounds is usually not necessary. Cultural control in Qld: Seed loss may be prevented by covering the surface of the bed with fine sand that has been sterilised and sieved.
P R E- E M E RG E N C E ( N U R S E R Y )
Seedharvesting ants Pheidole spp. including Pheidole anthracina Forel Hymenoptera: Formicidae Pest status and distribution: Seedharvesting ants are minor, widespread and irregular pests of tobacco in north Qld.
S E E D L I N G A N D G ROW T H S TAG E S
Two-spotted mite Tetranychus urtice Koch Acarina: Tetranychidae N (Main entry in Chapter 3 Cotton.)
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Pest status: Two-spotted mites are minor, widespread and irregular pests of tobacco in Qld.
especially in seed beds. The risk period is for the duration of the crop for virus transmission.
Risk period: Mites may be induced by broadspectrum pesticides used against other tobacco pests. They are also favoured by hot dry weather.
Large concentrations of aphids may develop under the lower leaves of field plants, especially in the cooler months. Feeding is associated with the production of honeydew. Cured leaves lack body and are of low quality.
Damage: Mites feed on the under-side of the leaf and damage is seen as a mottling of the leaf colour. Severely affected leaves become yellow and may fall from the plant. Monitoring in Qld: Examine the leaves on the bottom half of the plant. Count mites on both leaf surfaces. Examine five consecutive plants in the row at six widely scattered positions throughout the crop (total of 30 plants). Action level in Qld: Spray when the number of all mobile stages exceeds 20 per leaf. Chemical control in Qld: Chemical control may be cost-effective when indicated by monitoring. Two-spotted mites are resistant to a number of insecticides and miticides, and populations increase after applications of pesticides to which they are resistant.
Spur-throated locust Nomadacris guttulosa (Walker) Orthoptera: Acrididae N (Main entry in Chapter 18 Locusts and grasshoppers of
pastures and rangelands.)
Spur-throated locusts are minor, widespread and irregular pests of tobacco in North Qld. Both adults and nymphs damage leaves by their chewing. Chemical control may be necessary when numbers and damage begin to increase.
Aphids Including Myzus persicae (Sulzer) Hemiptera: Aphididae Pest status: A number of aphid species cause minor damage to tobacco crops in north Qld and Vic., where they are widespread and damage is irregular. Damage: Aphids are vectors of potato virus Y (PVY). Transmission occurs rapidly following the commencement of feeding. Small numbers of aphids may infect a large number of plants,
Monitoring in Qld: Sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants). Examine lower leaves. Action level in Qld: Treat if aphid numbers exceed 300 on 30 plants. Chemical control in Qld: Chemical application has not been demonstrated to reduce virus transmission but may reduce feeding damage. Cultural control in Qld: Maintain seedbed and field crop areas free of solanaceous weeds and crops. Destroy crop residues promptly.
Common brown leafhopper Orosius argentatus (Evans) Hemiptera: Cicadellidae Distribution: Africa, East and South-East Asia, some Pacific Islands, and Australia, where it is common in all states. Pest status: The common brown leafhopper is a major disease vector in Vic. tobacco crops, where it is widespread and regular in occurrence. Identification: Adult females are 3.2–3.5 mm long, males are 2.0–3.0 mm long. The head is pale yellow with brown markings and brown eyes. The thorax and wings have a characteristic brown pattern (Fig. 12.1). Eggs are 0.8 × 0.3 mm and may be seen as a small glistening lump in plant tissue. Newly hatched nymphs are 1.8 mm long with bright red eyes. The colour of nymphs varies from light cream, orange to dark grey. Wing pads are visible in third-instar nymphs. Fully grown (fifth-instar) nymphs are 2.8–3.4 mm long. May be confused with: Due to their distinctive colouration, adults are unlikely to be confused with other common leafhoppers in tobacco. Several species of leafhoppers in the genera Balclutha Kirkaldy and Zygina Fieber and the species Limotettix incertus Evans are capable of transmitting virus particles. These leafhoppers 345
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into a slit cut into plant tissue by the ovipositor. Under favourable conditions, the five nymphal stages are completed in 25 days. Mating occurs soon after the adult moult and the female begins to lay eggs 3–7 days later and is capable of laying up to six eggs per day for several months. The eggs laid in February give rise to the summer generation. The third overwintering generation appears during March in Vic. (Helson 1942).
Fig. 12.1. Common brown leafhopper adult (3 mm long). (SARDI: G. Caon)
are coloured green to yellow and are smaller and more delicate looking than common brown leafhoppers. Host range: These leafhoppers are found on many non-grass hosts, most of which are regarded as weeds. The range of hosts on which they can breed (and on which nymphs may be found) is more limited. Autumn–spring annual hosts include some medics, clovers, capeweed, mustard, common crowfoot and wild radish. Summer annual hosts (growing after summer rain, or under irrigation) include some Solanaceae, Chenopodiaceae, Boraginaceae and Portulacaceae. Perennial hosts include some mallows (Malvaceae). Common brown leafhoppers are not able to breed on tobacco, which appears to be an accidental host, but adults can survive long periods on tobacco and other non-preferred hosts (Bowyer and Atherton 1971; Helson 1942; Thomas and Bowyer 1980). Life cycle: The common brown leafhopper has three generations per year in Vic. Adults and fifth-stage nymphs survive the winter on nongrassy pasture and weeds surrounding tobacco areas and during spring they actively feed on these plants, some of which are reservoirs of phytoplasmas. Young nymphs do not move far from where they hatched. As these spring hosts die, adult leafhoppers disperse, with adults flying up to 7 m above the ground during evenings at temperatures between 22°C and 37°C with light to moderate air movement (Hosking and Danthanarayana 1988). On suitable host seedlings, adults insert eggs singly
Disease transmission: Tobacco yellow dwarf virus (TYDV) is Australian in origin and is transmitted persistently by the leafhoppers. Common brown leafhoppers do not undertake exploratory probing of a food source but insert their stylets directly into plant tissue. Stylet insertion into plant tissue appears to be random, and a matter of chance whether phloem tissue is penetrated (Day et al. 1952). After feeding on a plant infected with TYDV, the minimum latent period of the pathogen in the leafhoppers is 24–48 hours, and the insects remain infective for at least 21 days. Individuals that acquire the pathogen in the nymphal stage remained infective as adults. Risk period: From seedling to maturity. Tobacco plants are susceptible to the virus during seedling and growth stages. Damage: The common brown leafhopper is a vector of TYDV (Geminiviridae: Mastrevirus), one of the most significant causes of lost production in Vic. tobacco if left untreated (Fig. 12.2). Monitoring: No practical monitoring technique is available that is sufficiently sensitive to detect small numbers of leafhoppers. Action level in Vic.: Any infective leafhoppers can cause economic damage.
Fig. 12.2. The tobacco plant on the left shows symptoms caused by tobacco yellow dwarf virus, compared with a healthy plant of the same age. (Tobacco Co-operative of Victoria: C. Parker)
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Chemical control in Vic.: Applications of insecticides aimed at preventing any leafhopper probing or feeding on tobacco plants (Paddick and French 1972). Cultural control in Vic.: Weeds, including wild radish, are reservoirs of the virus. Control of weeds in the crop and surrounding areas may be important in reducing transmission to tobacco (van Rijswijk et al. 2004).
problem in Vic. tobacco crops but could cause major damage if western flower thrips, Frankliniella occidentalis (Pergande), became established in production areas.
African black beetle Heteronychus arator (Fabricius) Coleoptera: Scarabaeidae N (Main entry in Chapter 13 Pastures—summer rainfall.)
Green vegetable bug Nezara viridula (Linnaeus) Hemiptera: Pentatomidae N (Main entry in Chapter 7 Pulses—summer.)
Pest status: Minor, widespread and regular pest of tobacco in north Qld and a minor and irregular pest in Vic. Identification: See Chapter 7. The green vegetable bug is unlikely to be confused with other insects on tobacco and may be present at any time after germination. Damage: Feeding causes wilting of leaves and buds and stunting of plants. Monitoring in Qld: Sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants). Examine crop margins if these are next to tropical legumes. Action level in Qld: Spray if five bugs or fresh bug damage is observed. Chemical control in Qld: Chemical control is cost-effective when indicated by monitoring. Cultural control in Qld: Destroy crop residues promptly and avoid planting next to tropical legumes, tomatoes and capsicums. Maintain headlands and irrigation rows weed free.
Thrips Thysanoptera, including onion thrips Thrips tabaci Lindeman
Pest status: Adults are minor, widespread and irregular pests of young tobacco plants in Vic. Adults fly into new plantings during spring and chew the young plants. Adults may also be found in soil adjacent to plants. Damage may be prevented by pre-planting insecticide application.
False wireworms (Gonocephalum spp) Coleoptera: Tenebrionidae N (Main entry in Chapter 5 Oilseeds.)
Pest status: False wireworms are minor, widespread and irregular pests of tobacco plants in north Qld and Vic. Risk period: Plants are at risk between transplanting and establishment. Damage: Damage is mainly caused by larval feeding which severs stems of small seedlings at and below the soil surface. Adults feed mainly at, or just below, the soil surface. Monitoring in Qld: Sample plants in 2-m lengths of row at six randomly selected positions. Examine roots of damaged plants and surrounding soil to a depth of 10 cm for larvae. Action level in Qld: Spray if one or more larvae are found in total. Chemical control in Qld: Pre-plant insecticide application may be cost-effective. Cultural control in Qld: Prompt destruction of crop residues and weed-free headlands and irrigation rows reduces pest populations.
Pest status: Thrips are minor, widespread and irregular pests of Vic. tobacco.
Cutworms
Damage: Some thrips species transmit tomato spotted wilt virus, which is occasionally a minor
Agrotis spp., including Agrotis munda Walker Lepidoptera: Noctuidae 347
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N (Main entry in Chapter 2 Cereals.)
Pest status: Cutworms are minor, widespread and irregular pests of tobacco seedlings in north Qld and Vic. Risk period: Risk of damage occurs in seed beds and at transplanting until plants are established. Damage: Cutworms feed at night on the base of plant stems, which may be severed. Symptoms of dead seedlings may be confused with damage caused by false wireworms. Monitoring in Qld: In seed beds, examine the plants surrounding bare patches of seed beds for damage. Dig to a depth of 5 cm around the base of damaged plants. In the field, sample plants in 2-m lengths of row at six randomly selected positions. Examine roots of damaged plants and surrounding soil to a depth of 10 cm for larvae (Elder et al. 1992). Action level in Qld: In seed beds, spray if a single larva is found. In the field, spray if one or more larvae are found in total. Chemical control in Qld: Chemical control may be cost-effective when indicated by monitoring.
Tobacco stemborer Scrobipalpa heliopa (Lower) Lepidoptera: Gelechiidae Distribution: A native of Australia, now in Asia and Africa. Pest status: A major pest in the Mareeba and Dimbulah growing areas of north Qld, where it is widespread and occurs regularly. Identification: Eggs are 0.5 mm long, oval– cylindrical and pearly white when laid, turning lemon-yellow prior to hatching. Larvae are 1 mm long after hatching and grow through five stages (Patel and Chari 1978) to 12 mm and are creamy white. Adults are coppery red in colour with a wingspan of 1 cm.
Fig. 12.3. Tobacco stemborer damage on older tobacco. The tissue beyond the stem mine is dead. (DPI&F Qld: S.G. De Faveri)
Life cycle: The life cycle is completed in 28 days in summer. Risk period: All stages of growth of the tobacco plant are susceptible, but seedlings and transplants are most vulnerable. Damage: Symptoms of damage in seedlings and young plants include stem-galling and stem discolouration, wilting, and rosetting of the terminal leaves, shrivelling of older leaves (Fig. 12.3), and suckering of the plant and small lateral leaf mines near leaf midribs. Stemboreraffected plants are more prone to wilting under hot conditions than are healthy plants. Monitoring in Qld: Seed beds should be sprayed regularly so monitoring is not necessary. In the field, sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants). Check for leaf mines and bud damage on plants. Examine leaf tips for mines after topping (Elder et al. 1992).
May be confused with: Leaf mines may be confused with those of potato moth, Phthorimaea operculella (Zeller) Lepidoptera: Gelechiidae.
Action level in Qld: In seed beds, spray seedlings weekly. In the field, on plants less than 25 cm in height spray when the population exceeds one larva on 30 plants. Re-plant damaged plants. On plants more than 25 cm in height, spray when the population exceeds more than one larva on 30 plants. Do not re-plant damaged plants. After topping, spray if the number of mines exceeds one per 30 plants.
Host range: Some species of wild tobacco, and eggplant.
Chemical control in Qld: Chemical control is cost-effective when indicated by monitoring.
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Chemical control protects the plant but will not control established larvae. Cultural control in Qld: Crop residues and old seed beds should be destroyed promptly. Damaged seedlings should be culled before planting. Plants from the outside of the bed should not be used as they are more likely to be attacked.
Potato moth Phthorimaea operculella (Zeller) Lepidoptera: Gelechiidae Distribution: Cosmopolitan. Pest status: Major widespread and regular in Qld. Identification: Eggs are minute and pearly in colour. Eggs may be laid on the plant or on soil. Most found on the plant are pressed firmly between the globular hairs of the lower leaves. Larvae vary in colour from pink to grey–green after hatching. They are 12 mm long when fully grown and are creamy white in colour. Adults are small with a wingspan of 12 mm and are brownish grey in colour. May be confused with: Tobacco stemborer leaf mines. Host range: Potato, tomato, eggfruit, and solanaceous weeds such as blackberry nightshade, tree tobacco, devil’s apple and cape gooseberry. Life cycle: The life cycle is completed in 30 days during summer. Risk period: During the entire cropping season. Damage: Larvae tunnel directly into the plant tissue after emerging from the eggs (Fig. 12.4). On very small plants larvae may travel between plants in tunnels constructed of soil particles, debris and silk. Initially, damage occurs on lower leaves with progressive development of attack higher up the plant. Larvae sometimes destroy the supporting plant tissue in stems with damaged plants likely to blow down in moderate winds. Yield is reduced in heavily mined leaves. Monitoring in Qld: Preventive spraying for tobacco stemborer in seed beds negates the need
Fig. 12.4. Potato moth mines (white areas) on a tobacco leaf. (DPI&F Qld: K.H. Halfpapp)
for monitoring. In the field, sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants). Action level in Qld: Treatments for tobacco stemborer in seed beds will control potato moth. On plants less than 25 cm in height, spray if there is more than one larva on 30 plants. On plants 25–100 cm high, spray if there are more than 10 larvae on 30 plants. Spraying after picking has commenced is seldom warranted (Elder et al. 1992). Chemical control in Qld: Chemical control is cost-effective when indicated by monitoring. Potato leafminer is resistant to some chemical groups. Cultural control in Qld: Control of solanaceous weeds is important. Avoid growing potato and tomato crops adjacent to tobacco. Crop residues and old seed beds should be destroyed promptly.
Cluster caterpillar Spodoptera litura (Fabricius) Lepidoptera: Noctuidae N (Main entry in Chapter 3 Cotton.)
Pest status: Minor, widespread and irregular pests of tobacco in north Qld.
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Damage: Young larvae are gregarious and in groups skeletonise lower leaf surfaces, leaving only the veins. Older solitary larvae chew out large pieces of leaf. Monitoring in Qld: Sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants). Action level in Qld: When plants are 10–100 cm high, spray if there is more than one larva in 30 plants. When plants are more than 100 cm high, spray if there are more than six larvae on 30 plants. Chemical control in Qld: Chemical control may be cost-effective if indicated by monitoring.
ground. On large expanded leaves, the damage caused is proportional to the number of larvae present. A high population can strip leaves to the midribs and main veins. In some cases, larvae will attack the midribs causing wilting, sunburning or fracturing of the leaf. Sometimes the stem is attacked also, causing collapse or reduced growth of the plant above that point. Damage by larvae to expanding leaves results in leaf tissue losses. In nursery seedlings, this damage makes the plant unsuitable for transplanting. In young field plants, if the growing point is also destroyed by larvae, the plant produces profuse sucker growth.
Cultural control in Qld: Destroy tomato, capsicum and tobacco crop residues promptly. Maintain headlands and irrigation rows weed free.
Monitoring in Qld: Treatments for tobacco stemborer in seed beds negate the need for monitoring. In the field, sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants).
Helicoverpa spp. caterpillars
Action level in Qld: Spray if one or more larvae are found in 30 plants.
Lepidoptera: Noctuidae Cape gooseberry budworm, Helicoverpa assulta (Guenée) Corn earworm, Helicoverpa armigera (Hübner) Native budworm, Helicoverpa punctigera (Wallengren) N (Main entry in Chapter 3 Cotton.)
Corn earworm and native budworm are major pests of tobacco in north Qld, widespread in tobacco crops and regular in occurrence. Native budworm is a major pest of tobacco in Vic., where it is widespread and regular in occurrence. Cape gooseberry budworm is a minor pest of north Qld tobacco. Risk period: Tobacco crops are at risk during the growing season. Corn earworm larvae predominate in Qld from July to September, while native budworms are the more numerous between September to October (Titmarsh et al. 1990). In Vic., the risk period is late November to January. Eggs are laid equally on both upper and lower leaf surfaces (Broadley 1978). Helicoverpa assulta larvae also occur on tobacco in north Qld. Damage: Corn earworm and native budworm will feed on any part of the plant above the
Chemical control: Chemical control may be cost-effective when indicated by monitoring. Corn earworm in Qld is resistant to a number of chemical groups.
Looper caterpillar Chrysodeixis argentifera (Guenée) Lepidoptera: Noctuidae Distribution: A native of Australia. Pest status: Major in Qld, minor in Vic., widespread and irregular in both areas. Identification: Eggs are round, slightly flattened, white, 5 mm in diameter and are visible to the naked eye. They are usually laid singly on the under-surface of larger leaves, within 1 cm from the edge. Moths lay eggs mainly from mid-October to early November in Qld (Broadley and Swain 1981) and November to December in Vic. At emergence, the larva is 2 mm long, has a black head with a thin colourless body bearing prominent black hairs. Larvae are mostly found on the under-side of tobacco leaves (Broadley
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and Swain 1981). The larva takes on its typical looper shape and translucent green colour at the second instar. The prominent hairs are less noticeable at this stage. The older larvae are grass-green with two white stripes running along the back. They are smooth and slender and grow to a length of 4 cm. The adult moth is predominantly bronzy brown in colour with distinct silver markings on the forewing. The hindwings are brown with a coppery sheen. They are stoutly built with a 3 cm wingspan. May be confused with: The larva or its damage are unlikely to be confused with other tobacco insects. Host range: Some species of wild tobacco, potatoes, blackberry nightshade, tomatoes and beans. Life cycle: The life cycle is completed in 28 days in summer. Risk period: During the warmer months on all crop stages. Damage: Young larvae eat the under-side of leaves (3 mm diameter), leaving a translucent upper surface. The larva eats a number of these areas, leaving a windowed appearance on the leaf. Subsequent growth stages are not restricted to the lower surface and many move to leaves in the middle of the plant where, if large numbers are present, only the stems and larger leaf veins will be spared. Monitoring in Qld: Sample five consecutive plants in the row at six widely spaced intervals throughout the crop (total of 30 plants). Action level in Qld: Spray when numbers of medium-sized loopers (10 mm long) exceed one on 30 plants for plants less than 25 cm high, and 20 on 30 for plants more than 25 cm high. Chemical control in Qld: Chemical control may be cost-effective when indicated by monitoring. Cultural control in Qld: Crop residues and old seed beds should be destroyed promptly. Damaged seedlings should not be planted out. Early planting may allow plants to grow before the main egg-laying period.
POST HARVEST
Tobacco beetle or cigarette beetle Lasioderma serricorne (Fabricius) Coleoptera: Anobiidae N (Main entry in Chapter 2 Cereals.)
Distribution: Cosmopolitan, but most abundant in the tropics. In cold climates, it is troublesome only in heated buildings. Pest status in tobacco: Major, widespread, regular. Identification: The adult is small, 3–4 mm long, oval, stout, brownish-red, with a head strongly deflexed under the prothorax and with long saw-like (serrate) antennae. The larva is a small, white, hairy, C-shaped grub. May be confused with: Drugstore beetle and hairy fungus beetle, which do not have the curved under-side to the wing covers. Host range: The tobacco beetle is a major pest of store tobacco and a pest of cereals, herbs and spices such as paprika, red pepper and curry powder. It also occurs in dried flower arrangements and potpourris. Life cycle: Adult beetles live 2–4 weeks. During this period, a single female lays up to 100 eggs, loosely on the commodity. On hatching, larvae burrow in to the commodity and feed on it. After four to six instars, the larva pupates in a cell made of fragments of food bound with silk. Adults do not feed and can readily fly. Under optimal conditions of 30°C and 70% relative humidity, the life cycle takes up to 26 days. Maximum population growth rate per month can be up to 20 times. Risk period: Summer months. Damage: Larvae feed on stored tobacco causing loss of quality by perforating the leaf and contamination by cocoons, frass and dead insects. Adults feed very little. Larvae avoid light so populations may develop unseen within stored tobacco rather than on its surface. If not controlled, huge populations build up quickly and cause considerable damage. Both larvae and
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adults can chew through the packaging materials such as plastic, paper and wooden containers to gain entry to the food source.
Pest status: A minor pest of stored tobacco in Vic., where the adult moths have been collected on sticky traps in warehouses.
Monitoring: This beetle pest can be monitored by using both light and pheromone-based traps.
Identification: The caterpillar is initially dark, and later becomes yellow, with a dark line down its back, and a dark brown head. It lives in a loose web spun among its food. When one caterpillar encounters another, they each produce a small amount of light brown liquid from their mouths, and this causes them to walk apart: a mechanism that appears to prevent overcrowding of the caterpillars. The adult moth is brown, and has a wingspan of about 2 cm.
Action level: When detected. Chemical control: Fumigation in sealed storage. For small quantities of infested material, heating to 60°C is effective. Cultural control: General hygiene including vacuuming of shelves and disposal of infested containers. Destroy all stored tobacco residues, switch lights off and keep windows and doors of warehouse closed when not in use.
Tobacco moth
Host range: Vegetable products, including cereals, and cocoa.
Ephestia elutella Hübner Lepidoptera: Pyralidae
Monitoring: Use sticky traps in warehouses and other areas where tobacco is stored.
Distribution: Cosmopolitan species.
Cultural control: See tobacco beetle.
Sources of information Broadley, R.H. (1978). Distributions of Heliothis spp. (Lepidoptera: Noctuidae) eggs and first instar larvae on pre-flowering, flue-cured tobacco plants in north Queensland. Queensland Journal of Agricultural and Animal Sciences 35: 73–76. Broadley, R.H. and Swain, A.J. (1981). Chrysodeixis argentifera (Guenée) (Lepidoptera: Noctuidae) egg and larval distributions on flue-cured tobacco plants in north Queensland. Queensland Journal of Agricultural and Animal Sciences 38: 139–142. Bowyer, J.W. and Atherton, J.G. (1971). Summer death of French bean: new hosts of the pathogen, vector relationship, and evidence against mycoplasmal etiology. Phytopathology 61: 1451–1455. Day, M.F., Irzykiewicz, H. and McKinnon, A. (1952). Observations on the feeding of the virus vector Orosius argentatus (Evans) and comparisons with certain other Jassids. Australian Journal of Scientific Research Series B 5: 128–142. Elder, R.J., Brough, E.J. and Beavis, H.S. (Eds)(1992). Managing Insects and Mites in Field Crops, Forage Crops And Pastures. Department of Primary Industries, Queensland. 143pp. Helson, G.A.H. (1942). The leafhopper Thamnotettix argentata Evans, a vector of tobacco yellow dwarf. Journal of the Council for Scientific and Industrial Research 15: 175–184. Hosking, J.R. and Danthanarayana, W. (1988). Low level flight activity of Nesoclutha pallida (Evans), Orosius argentatus (Evans) and Zygina zealandica (Myers) (Hemiptera: Cicadellidae) in southern Victoria. Journal of the Australian Entomological Society 27: 241–249. Paddick, R.G. and French, F.L. (1972). Suppression of tobacco yellow dwarf with systemic organophosphorus insecticides. Australian Journal of Experimental Agriculture and Animal Husbandry 12: 331–334. Patel, B.H. and Chari, M.S. (1978). Bionomics of tobacco stem borer (Gnorimoschema heliopa Low). Gujarat Agricultural University Research Journal 3: 82–85.
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Titmarsh, I.J., Storey, R.I., and Strickland, G.R. (1990). The species composition of Helicoverpa Hardwick (Heliothis Ochsenheimer) (Lepidoptera: Noctuidae) infestations on tobacco in far north Queensland. Journal of the Australian Entomological Society 29: 81–86. van Rijswijk, B., Rodoni, B.C., Revill, P.A., Thomas, J.E., Moran, J.R. and Harding, R.M. (2004). Analysis of variability in partial sequences of genomes of Tobacco yellow dwarf virus isolates. Australasian Plant Pathology 33: 367–370. Thomas, J.E. and Bowyer, J.W. (1980). Properties of tobacco yellow dwarf and bean summer death viruses. Phytopathology 70: 214–217.
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13 PASTURES—SUMMER RAINFALL R.J. Elder
Brahman cross cattle browsing Leucaena leucocephala inter-planted with green panic grass, Panicum maximum. (P. & J. Larsen) GROUP Pasture grasses and forbs snails and slugs grasshoppers and locusts planthoppers beetles
soldier flies caterpillars
ants Leucaena insects psyllids and scales beetles caterpillars
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common garden snail slugs See Chapter 18 pasture planthopper pasture scarabs crabgrass leaf beetle amnemus weevils rough brown weevil sugarcane soldier flies flatheaded pasture webworm and roundheaded pasture webworm Ebor grassgrub and Tindale’s grassgrub sod webworm (or grass caterpillar) buffel grass seed caterpillar common armyworm dayfeeding armyworm and lawn armyworm cutworms funnel ants
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leucaena psyllid long soft scale leucaena seed bruchid leucaena flower-eating caterpillar
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The summer rainfall pastures of northern Australia are composed of native grasses and annual forbs that support large insect communities, some species of which (such as grasshoppers, locusts and native budworm) cause economic damage when they migrate to cropping areas. However, insects rarely cause economic damage to rangeland pastures because of the pastures’ low economic value per unit area to the grazing animals (mostly sheep for wool and beef cattle). Insect damage often occurs during the wet season when pastures can rapidly replace any lost production without any shortfall for stock. In higher rainfall areas and under irrigation, pastures ‘improved’ with introduced grasses and legumes allow intensive grazing. Pest control in these pastures may be cost-effective, especially during the establishment phase to prevent loss of seed, seedlings and foliage, or where pasture plants are harvested for seed. Leucaena (Leucaena leucaecephala) is a legume browse tree planted in parts of Qld and WA as a component of both dryland and irrigated pastures. Leucaena is a genus of tropical legumes originating in central America. L. leucocephala was introduced into Australia in the 19th century. It is planted in rows 1.5 to 10 m apart, usually with pasture grasses grown in the interrow spaces in low frost areas where annual rainfall is above 500 mm per annum. It may also be an environmental weed. The insect species that leucaena supports may be regarded as pests when the trees are cultivated, but they also act as biological control agents and may prevent the spread of leucaena to unwanted sites such as creek lines.
PA S T U R E G R A S S E S A N D F O R B S
Common garden snail
been reported damaging clovers, kikuyu grass and ryegrass. Control may be cost-effective when high numbers threaten high-value seed crops.
Pasture planthopper Toya sp. Hemiptera: Delphacidae Pest status and distribution: A minor pest of para grass, Brachiaria mutica, occurring in most of Qld where the grass is grown. Infrequent in occurrence. Description: The adults vary in colour from light to dark brown with two dark brown or black stripes on the face of the head. They have smoky coloured wings and a length of 2 mm to nearly 4 mm. Two forms are usually present: short- and long-wing forms. They have sucking mouth parts that are held close to and underneath the body when not in use. They fly when disturbed, and in large numbers they appear as clouds. Risk conditions: Most damage occurs during mild and moist winters. Damage disappears in spring.
Pasture scarabs Coleoptera: Scarabaeidae N (Main entry in Chapter 15 Pastures—winter rainfall.)
Pest status: Minor pests in summer rainfall pastures, where damage is recorded mainly from high rainfall (750 mm) areas of Qld highlands (750 m) with a temperate climate. Damage: Larvae may damage mixed pastures of annual grass and clovers, lawns, parks and golf courses. Permanent damage to pasture is uncommon, but unfavourable pasture conditions the following spring and summer may accentuate larval damage. Maintenance of pasture vigour by reduced grazing and applying fertiliser and water may reduce damage.
Helix aspersa (Müller) Eupulmonata: Helicidae
Slugs
Crabgrass leaf beetle
Eupulmonata
Oulema rufotincta (Clark) Coleoptera: Chrysomelidae
Minor pests, with damage restricted to high rainfall areas. In summer pastures, they have
Pest status: A minor pest in coastal and subcoastal pastures with irregular occurrence.
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Description: Larvae grow to 5 mm in length, are creamy coloured with a dark brown head that may be partially retracted into the under-side of the body. The body is globular in shape and covered with slime. The larval stage lasts 2–3 weeks. The 3 mm long beetle may have metallic green wing covers, while a colour variant has two brown spots on the wing covers.
adults, are white when they first moult from the larval stage and slowly darken to the adult colour. Host range: Known hosts are Fabaceae including clovers, Trifolium spp., Desmodium spp. and Glycine spp., and to lesser extent siratro, Macroptilium atropurpureum.
Damage: Larval damage is recorded on woolly finger grass, Digitaria eriantha, and couch Cynodon spp. Larvae erode the upper surface of the leaf blade between the veins, leaving a thin layer of translucent tissue. This produces a series of parallel longitudinal striations extending down the leaf blade. The first sign of pasture damage is a small brown circular or irregularshaped patch that expands and may give the appearance of frost damage.
Life cycle in pastures: Annual life cycle. Adults occur in pastures from November to May and oviposition is from January to May. Eggs are inserted in hollow petioles of clover plants and in the hollow stems of grasses associated with tropical legumes or in dead runners of the legumes in the ground litter. The larvae feed through the autumn and winter on the roots of legumes. Pupation occurs in the soil between September and November and the adults emerge from November.
Amnemus weevils
Risk Period: Following germination during autumn and winter.
Coleoptera: Curculionidae Amnemus quadrituberculatis (Boheman) and A. superciliaris (Pascoe) Distribution: South-east Qld and northern NSW. Pest status: Major, widespread in legume pastures, frequent. Identification: Adults of both species of amnemus are large weevils about 10 mm in length with a short thick ‘snout’ (rostrum). They are dark brown in colour often covered by an adhering layer of soil (Fig. 13.1). Larvae of amnemus weevils are legless, similar in shape to white grubs, have a creamy white body and a light brown head. They can grow to 15 mm long. Pupae are similar in shape to
Fig. 13.1. Adult amnemus weevils, length 10 mm. (DPI&F Qld)
Damage: Most damage is by larvae that feed on and in the tap-root(s), forming large distinctive channels (Fig. 13.2) and may cut off the root(s) and kill the plant. Adults feed on the leaves and are only a problem to young seedlings. Action level: At first signs of seedling damage. Treatment is warranted when there is a rapidly increasing area of damaged germinating plants. Monitoring: Inspect the pasture 5 days after planting or as the legume seedlings emerge. Damage may be confined to small areas. If symptoms are detected, search for weevil adults at night when they are active. They hide in the soil during the day. Weevil activity is often confined to patches within a pasture.
Fig. 13.2. Larvae of amnemus weevil (longest is 15 mm) channelling roots. (DPI&F Qld)
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Chemical control: May be cost-effective, depending on current registration. Control should aim at killing adults and may be more effective when adults are active during evenings and at night than during the day, when they are hidden. Spot-spraying located patches may be sufficient. Cultural control: Maintain favourable growing conditions to enable the legume component of the pasture to compensate for loss of plants through weevil damage.
Rough brown weevil Baryopadus corrugatus Pascoe Coleoptera: Curculionidae Distribution: Northern Australia. Pest status: Major, restricted to areas north of Townsville, regular. Identification: The flightless adults are large weevils about 10 mm in length with a short thick ‘snout’ (rostrum). They are dark mottled-brown in colour but this is often hidden under an adhering layer of soil. Larvae are legless up to 15 mm in length, have a creamy white body and a light brown head. Pupae are similar in shape to adults, are white when they first moult from the larval stage and slowly darken to the adult colour. Host range: Adults (on foliage) and larvae (on roots) have been recorded feeding on the legume species Desmodium intortum, D. triflorum, D. uncinatum, Lablab purpureus, Macrotyloma axillare, Macroptilium atropurpureum, M. lathyroides, Phaseolus vulgaris, Vigna marina and Vigna sinensis. In the absence of preferred hosts, adults have also been recorded feeding on the foliage of Arachis hypogaea, Glycine max, G. wightii, Stylosanthes guyanensis and S. humulis.
Fig. 13.3. Larva of the rough brown weevil (15 mm long) in a feeding channel. (DPI&F Qld)
Damage: Larvae feed on and in the main roots leaving large channels (Fig. 13.3). This feeding can reduce plant vigour, causing wilting, leafyellowing, leaf-shedding and plant death. In severe infestations, complete loss of the legume from the paddock can occur, frequently in D. intortum and M. atropurpureum sward seed crops in the second and subsequent years of the crop’s life. Losses are also common in V. sinensis and L. purpureus (usually row) crops, where susceptible crops have been grown the previous year. Adult-feeding has destroyed seedling crops of G. wightii. Action level: One or more adults per 30 plants. Monitoring: Look for adults at the seedling stage by examining five consecutive plants at six widely spaced locations throughout the crop. Adults emerge from the soil with the first storm rains in October to early December. More than one emergence is possible. If greater than 12 mm of rain falls in the following month a further sample is required. Chemical control: Controlling adults after the early summer storms can save seedling stands.
Life cycle: One year life cycle. Adults emerge from earthen cells in the soil (at a mean depth of 100 mm) with the first storm rains of the wet season yielding approximately 50 mm in October–November. Larvae are present from late February to early October and pupae from March to November. All adults had emerged from their earthen cells by mid-December.
Cultural control: Avoid planting susceptible crops on areas planted to a susceptible crop in the previous year. Crops planted adjacent to susceptible crops should be checked for adults moving in after rain in October to early December. Adults cannot fly, so only areas adjacent to infested crops are subject to invasion.
Risk period: Adults damage seedling legumes at emergence and larvae attack the resultant legume stand on an ongoing basis.
Natural enemies: The pathogens Nomuraea rileyi, Aspergillus sp. and Metarhizium anisopliae have been recorded.
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Sugarcane soldier flies Inopus rubriceps (Macquart) Diptera: Stratiomyidae N (Main entry in Chapter 11 Sugarcane.)
Sugarcane soldier flies are minor pests of pastures in tableland areas of Qld, where they frequently occur. Larvae are recorded feeding on kikuyu, paspalum, narrow-leaf carpet grass Axonopus affinis, green and blue couch. Larvae feed by sucking on the outside of the root and stolon tissue below ground level. Large numbers have been recorded in pastures on the northern tablelands of Qld in the Atherton, Malanda to Millaa Millaa districts. No control is recommended.
Pasture webworms Lepidoptera: Hepialidae N (Main entries in Chapter 15 Pastures—winter
rainfall.)
Flatheaded pasture webworm, Oncopera mitocera (Turner) Roundheaded pasture webworm, O. brachyphylla Turner Ebor grassgrub, O. alboguttata Tindale Tindale’s grassgrub, O. tindalei Common Distribution: All species are native to Australia. Flatheaded and roundheaded pasture webworms are found on the Atherton Tablelands of north Qld. Ebor and Tindale’s grassgrubs are recorded from the northern tablelands of NSW. Pest status: Roundheaded and flatheaded pasture webworms: major, restricted to tablelands of north Qld, regular occurrence. Ebor grassgrub: minor, restricted to northern tablelands of NSW, irregular. Tindale’s grassgrub: minor, restricted, irregular. Identification: Roundheaded and flatheaded pasture webworms: Eggs are spherical, grey in colour, about 0.5 mm in diameter. Larvae moult up to 15 times before they reach full size when they may be up to 65 mm in length. They are greenishgrey in colour with a distinct yellowish tinge in older larvae. Pre-pupae, which do not feed, are
caterpillar shaped and cream in colour. Pupae have the legs, wings and antennae attached to the body, are brown in colour, cylindrical in shape with a nob on the tip of the head. Larvae, pre-pupae and pupae never completely leave their tunnel. The moths of the two species are similar in shape and appearance. Moths are up to 20 mm in length and hold their wings in an inverted Vshape over their body at rest. Males are smaller than females and the forewings are rich dark brown in colour often with variable white patterns either as dots or streaks. The hindwings are dark grey–brown in colour. Females have much smaller eyes and are overall speckled grey in colour. May be confused with: Flatheaded and roundheaded pasture webworms may be distinguished by their tunnels. The tunnel entrance of the flatheaded pasture webworm has a covered feeding and storage chamber at the surface, while that of the roundheaded pasture webworm opens directly onto the surface. Host range: All pasture species. Life cycle in pastures: The complete life cycle takes about 12 months. Adults mate in trees (flatheaded) or low vegetation (roundheaded). The females fly over pasture at dusk and dawn scattering eggs singly from January to April. Eggs take 16 days to hatch. The larvae commence burrowing and feed on debris and fresh plant material. The larval stage takes about 37 weeks for roundheaded larvae and 47 weeks for flatheaded larvae. Pasture webworms live in a vertical tunnel in the soil and come up at night to feed on any pasture and/or debris that is within a body-length of the burrow entrance. The area around the entrance, both horizontally and vertically, becomes cleared of all material. The material and faecal pellets are often not consumed directly but are placed in the storage chamber. The larvae feed on pasture, plant debris and recycle their faecal pellets. The pre-pupal stage is about 9 weeks for roundhead and 3 weeks for flatheaded pasture webworm. Risk period: By the beginning of the dry season (August), the larvae have reached a size where they can cause serious damage, which may continue to January for flatheaded pasture 359
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webworm or November–December for roundheaded pasture webworm. Damage: Runners of pasture legumes may be cut off and die, reducing the legume component of mixed pastures. Weeds invade the pasture after feeding activity is finished in November to January depending on species. Monitoring: Estimate numbers of tunnels in August–September. This may be done by removing all vegetation from a 30 × 30 cm area and covering for 2 nights with hard board, which is then removed and the number of opened tunnels counted. Repeat at 20–30 representative points. Action level: One hundred webworms per square metre, a common density, can reduce dry season production by 800 kg ha–1. The annual pasture carrying capacity is determined by the capacity during the dry season when pasture production is lowest. Chemical control: Chemical control may be cost-effective and is best applied in August– September if damage is apparent. Prior to August, the caterpillars are too small to cause significant pasture loss. A single application to patches of webworm is likely to provide control. Cultural control: Maintenance of a vigorous pasture by use of fertiliser and weed control. Natural enemies: A number of insect diseases may effectively reduce webworm numbers.
Sod webworm (or grass caterpillar), Herpetogramma chrysotricha (Meyrick and Lower) Lepidoptera: Pyralidae Distribution: An Australian native, widely distributed throughout Qld and northern NSW. Outbreaks in pastures are more common in southern areas. It has been recorded south to Batemans Bay, NSW. Pest status: Minor and infrequent pest of pastures but a major and frequent pest of turf. Identification: Eggs are round and greenish and overlap each other like fish scales. The larvae at first are almost white in colour but later they acquire a greyish-green tinge that is accentuated by bright, raised spots on each segment of the body. When fully grown, the larvae are spindle-shaped and approximately
30 mm long. Pupae are about 15 mm long and light brown in colour when first formed but darken with age. The moths have a wingspan of about 25 mm. The wings are light, greyishbrown in colour, with narrow darker, somewhat waxy markings, particularly towards the outer margins. The moths can occur in such large numbers around lights that they become a nuisance in country homes. Host range: Grasses, including Cynodon incompletus, broadleafed carpet Axonopus compressus, fescue Festuca spp., kikuyu Pennisetum clandestinum, paspalum Paspalum dilatatum, perennial ryegrass Lolium perenne, rice, Rhodes grass Chloris spp., rye Secale cereale, and wheat. Life cycle: Female moths lay about 250 eggs over a 2–3 week period. The eggs are deposited on the upper surface of leaf blades near the base of the leaf and along the midrib. They are laid singly or as groups often in a line on the grass near ground level, in groups of five to 15. Eggs take 4 to 6 days to hatch. The caterpillar stage lasts 2 weeks in summer (24°C). Feeding is usually at night and the caterpillars hide in webbing tunnels during the day. Larvae pupate in cocoons that are covered with soil particles. Pupal duration is about 8 days. The life cycle can be completed in as little as 4 weeks and there may be several generations per year. Risk Period: The main risk period is from January to April. Damage: Larvae feed on debris and leaf blades low in the pasture, with debris usually being eaten first. They latter eat pieces out of leaf blades. Eventually the grass stems (runners) are the only aboveground plant parts left and a lawn or pasture will appear thin and browned-off. Where very large numbers occur as in recent outbreaks in New Zealand, larvae may feed in fronts in a similar manner to armyworms. Monitoring: Signs of sod webworm activity are grass pieces, webbing and faecal pellets. Larvae can be detected by putting out wet bags overnight and inspecting underneath the next morning. Action level: For pastures, when 20% of the area is showing signs of damage. For turf, spray at the first sign of small feeding patches and when more than 20 caterpillars are found under each square metre of wet bag left overnight.
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Chemical control: Spraying may be costeffective in turf. The chemical should reach the lower layers of turf. Conservation of natural enemies: A number of wasps including Leptobatopsis indica Ichneumonidae, Antrocephalus sp. Chalcididae, and a fly, Actia sp. Tachinidae, parasitise larvae (caterpillars). They do little to reduce the damage in the current caterpillar generation as they kill pupae, but they can be important in preventing continuation of an outbreak into the next generation. The parasitoids can occur in noticeable numbers flying over the lawn/pasture surface. A number of diseases have been recorded killing sod webworm larvae. Cultural control: Couch grasses, Cynodon spp. (green/speedy and blue), paspalum and kikuyu are more susceptible than some other species (e.g. Indian bluegrass, Bothriochloa pertusa), particularly when heavily fertilised to achieve high nitrogen levels and hence rapid growth.
Buffel grass seed caterpillar Mampava rhodoneura (Turner) Lepidoptera: Pyralidae Distribution: Native to Australia. Pest status: Major where host grasses are grown, regular occurrence. Identification: The egg is small (0.4 mm long, 0.2 mm diameter), white, elongate and ovoid. The larva is white, up to 18 mm long, with a brown head. The pupa is brown and 12 mm long. The adult moth is white with a wingspan of 25 mm. The margin of the forewing is tinged with pink and there is a small brown spot located 5 mm from the apex. May be confused with: Unlikely to be confused with other caterpillars on buffel grass. Host range: Cenchrus spp., Setaria spp., and the native host, swamp foxtail Pennisetum alopecuroides. Life cycle: After emerging from the pupal cocoon, the female mates and then lays eggs into the seed head. Head emergence to flowering, seems to be the most attractive stage for egg-laying. After hatching, the larvae feed on the developing seed. In moving through the
head, the larvae form tunnels from webbing and dead florets. In green heads, these tunnels appear as a white spiral down the head. Later, the remains of seeds and white to brown grasslike material are webbed loosely together. Seed heads attacked by this pest have a more fluffy appearance than normal. The larva then spins a silken cocoon in which it pupates. This cocoon is readily seen as a dense tuft protruding from the buffel seed head. Depending on temperature, the complete life cycle can take from 4–6 weeks. Risk period: Damage is most common in the third and subsequent crops where a number of sequential harvests are made, or in late summer and autumn. Damage: Seed yield and viability are reduced. One caterpillar in a head will destroy all the seed in that head. The pest does not affect dry matter yields for grazing. Monitoring: Sample the crop 10 days after seed heads emerge. Examine five adjacent heads at six widely scattered positions. Check carefully for caterpillars and/or webbing. At this stage caterpillars will be small and difficult to find. Action level: If six out of 30 heads are infested, a yield loss of 20% is likely and control at this point may be cost-effective. Chemical control: It will usually pay to spray the second and subsequent seed crops in a series. Spray 10 days after heads emerge. Cultural control: Heavy grazing or burning may reduce carry-over of the pest, and eradication of host plants prior to planting a new crop may reduce pest pressure. Conservation of natural enemies: Being a native, the buffel seed caterpillar has a range of parasites including tachinid flies, predatory shield bugs and spiders, but they do not prevent economic damage.
Common armyworm Leucania convecta (Walker) Lepidoptera: Noctuidae Distribution: Australia. Pest status: Minor, widespread, irregular. 3 61
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Identification: Larvae, when newly hatched, are pale in colour and about 1 mm long. After the first moult, faint longitudinal body stripes develop. Fully developed larvae possess conspicuous white, pink and brown bands about 1 mm wide running the whole length of the body. Body colour is variable: larvae may appear pinkish, brown or green. Crowded larvae tend to be darker than solitary larvae. The head capsules of mature larvae are mottled-orange in colour with a prominent Y-shaped marking. Larvae grow to 35 mm in length, each possessing three pairs of thoracic legs and five pairs of fleshy prolegs on the abdomen (Figs 2.33 and 2.34). Common armyworm moths have wingspans of 30–40 mm (Fig. 2.32). Their forewings are speckled yellow–brown, with a black dot near the centre. There is a faint dark line running the length of the wing and through the black dot. The hindwings are light grey brown with a much darker area over the last third of the wing and extending along the outer edge of the wing. The wing veins are darker than the rest of the wing. May be confused with: Larvae of other armyworm species are difficult to distinguish in the field, and are reared to adulthood for species identification. Identification to species is not required for control operations. Host range: All pasture grasses, and crops such as barley, oats, wheat, sorghum, maize, millets and Panicum spp. are hosts. Dicotyledonous plants are not preferred hosts. Life cycle on summer pastures: The duration of life cycle varies from 28 days in spring to 100 days in winter. The small, shiny white, spherical eggs are cemented to each other, and to the host plant, by a translucent adhesive. Eggs are laid by the moth at night usually in leaf litter, on dead leaves at the base of the plant, in the folded blades or under the sheaths of the upper leaves. Incubation of the eggs depends largely on prevailing temperatures, and may be as little as 3 or 4 days. Larvae usually pass through five moults, and larger larvae hide in the leaf debris at the base of the plant or in cracks in the soil and emerge at night to feed. Younger larvae tend to aggregate, but older larvae disperse. The name ‘armyworm’ describes the spectacular movement of large numbers of larvae to new feeding areas once an area is eaten out. The
larvae consume about 90% of their total food intake in the last third of the larval development (length 10 to 35 mm). Mature larvae leave the plant and burrow below the soil surface, where they transform into pupae in earthen cells. Pupae are light green–brown at first, but rapidly tan into a chestnut brown colour. Risk period: North of the Tropic of Capricorn, common armyworms are likely to occur during autumn and winter, and during winter and spring to the south. Damage: Larvae eat patches of pastures and may cut off the seed head of grasses. Monitoring: Feeding larvae are best detected after sunset, as they spend most of the day concealed in host plants or in the soil. Larvae smaller than 10 mm are easily missed without careful inspection. Feeding larvae can often be detected by the presence of ragged leaves and ‘frass’ or larval excreta on the plants and soil. This semi-digested plant tissue often lodges near the plant. Action level: Most pastures can compensate for armyworm damage, and control is rarely necessary. Conservation of natural enemies: Natural enemies frequently give effective control and usually ensure that there is only one generation during an outbreak. Armyworm larvae are subject to fungal (Nomuraea rileyi) and nuclear polyhedrosis virus, the latter of which can be readily recognised as they cause larvae to hang in an inverted V from the plant, the larvae rupturing readily to release liquefied body contents. There are a number of wasp and fly parasitoids. The wasps include the endoparasite Apanteles sp. (Hymenoptera: Braconidae) that feeds inside the larvae and is seldom seen until 10–30 wasp larvae emerge from the armyworm. They spin their small oblong fluffy white cocoons near the armyworm often in a cluster. Ichneumonid wasps (Hymenoptera: Ichneumonidae) are also common. The larvae of the wasp grow to 10 mm when mature and can be found in the cell under the soil constructed for pupation by the armyworm larvae. Wasp cocoons are constructed of tightly woven black fibres, and are oblong in shape. The fly parasitoids belong to the Family Tachinidae and can be seen attached to the skin of the
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armyworm larvae. The maggot-like larvae of the fly can be found in the body of the armyworm larva and pupates within the armyworm pupa. Predators include birds, larger wasps and various species of sucking bugs.
Dayfeeding armyworm Spodoptera exempta (Walker)
Lawn armyworm S. mauritia (Boisduval) Lepidoptera: Noctuidae Distribution: Africa, Asia Australia. Pest status: Minor, widespread in northern Australia. Dayfeeding armyworms are irregular in occurrence while lawn armyworms occur regularly. Identification: Larvae appear similar to other armyworm species (Fig. 13.4). Dayfeeding armyworm moths have wingspans of 30 to 40 mm. Their forewings are speckled dark grey to black. The hindwings are very light grey with a narrow much darker area along the outer edge of the wing (Fig. 13.5). Host range: The dayfeeding armyworm mainly feeds on members of the grass family (Poaceae), including guinea Panicum maximum, kikuyu Pennisetum clandestinum, love Eragrostis spp., molasses Melinis minutiflora, para Brachiaria mutica, paspalum Paspalum spp., red natal Melinis repens, Rhodes Chloris gayana and summer Digitaria ciliaris grasses, and also sugarcane,
Fig. 13.4. Lawn armyworm (Spodoptera mauritia) larvae eating kikuyu grass. The fully grown larvae (largest 45 mm in length and 7 mm in width) have an inflated sausage-like appearance and black triangular marks above the lateral stripes. (DPI&F Qld)
Fig. 13.5. Lawn armyworm moths, wingspans 30–40 mm. (DPI&F Qld: J. Wessels)
maize and sorghum. Recorded hosts of lawn armyworms include barley, Bermuda grass Cynodon nlemfuensis, green panic, kikuyu, kyllingia Cyperus spp., McCoy grass, nut grass Cyperus spp., oats, orchard grass Dactylis glomerata, pangola, para grass, paspalum, sedge, wheat and most lawn grasses. Life cycle: Lawn armyworm eggs hatch in 2–10 days depending on temperature, and the seven to eight stages of caterpillar growth are completed after 2–5 weeks and adults emerge from pupae after a further 1–2 weeks. Adults live 1–2 weeks and the species may complete two to three generations during summer and spring. Female armyworm moths mate the first night after emergence and 2 days later begin to lay clusters of eggs covered with fine scales onto the undersurface of non-host vegetation and objects such as the eaves of houses. The egg stage of the dayfeeding armyworm lasts 3–5 days and the larval stage as little as 2 weeks in summer. The 363
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newly hatched larvae fall to the surfaces below or are blown some distance on the end of fine silk strands. Larvae are initially found in loose aggregations but as feeding continues, larvae may become more dispersed. Fully grown larvae grow to 45 mm in length and have a pattern of distinctive green, dark green and yellow longitudinal stripes running the full length of the body. Uncrowded larvae show a considerable range of colour variation and do not move much. Crowded larvae are gregarious, have a distinctive blue–black colouration and are active movers. The larvae consume about 90% of their total food intake in the last third of the larval stage (length 10 to 35 mm). Prior to this, the larvae and their damage is easily missed without careful inspection. The movement of larvae across a paddock, together with the habit of feeding during daylight hours led to the name dayfeeding armyworm. When fully grown, larvae burrow into the soil where they pupate in an earthen chamber. The pupal stage takes several days.
leaves of particularly low-growing weeds. Eggs hatch in 3–7 days in warm weather and several weeks in cold weather. Caterpillars take 3–12 weeks to become fully grown. During warm weather, the pupal period is only 2–4 weeks, but in cold weather it can be more than 8 weeks. The larger larvae live in a silk-lined tunnel in the soil to which they may drag cut plants.
Risk Period: Autumn and winter.
Action level: Treatment with insecticides is warranted when there is a rapidly increasing area or proportion of pasture damage in newly planted stands.
Damage: Leaves up to 45 cm from ground level are stripped.
Damage: The caterpillars erode the plant surface when small, and chew out pieces when larger and may sever the stems of young seedlings at or near ground level, thereby causing collapse of the plant. Pasture damage may be patchy and destruction of available food may force the cutworms to migrate to adjacent areas. Monitoring: Inspect pasture 5 days after planting or as grass seedlings emerge. If symptoms are detected, search for cutworm larvae, which are active at night. Laying a wet bag on the soil overnight may attract the larvae and make detection easier.
Monitoring: Carefully check pastures weekly for small larvae from December to March. Damage by large larvae may be apparent during autumn and winter. Signs of damage are eatenout margins of leaves due to feeding of the older larvae and also by the faecal pellets around the base of the plant. The larvae often feed on the leaf blades leaving the midrib and so leaving the plant with a tattered appearance.
Aphaenogaster pythia Forel and A. longiceps Hymenoptera: Formicidae
Action level and control: Methods are similar to common armyworms.
Pest status: Minor pests, found in high rainfall dairy pasture, frequent in occurrence.
Cutworms Agrotis spp. (Lepidoptera: Noctuidae) N (Main entry in Chapter 2 Cereals.)
Pest status in pastures: Minor, widespread, irregular. Life cycle in pastures: Moths live for 4–6 weeks, although the oversummering moth of the common cutworm, Agrotis infusa (Boisduval), lives for about 6 months. They lay several hundred eggs in the top 10 mm of moist, recently cultivated soil or on the stems and
Chemical control: If distribution of the pests can be plotted accurately, spot-spraying may be costeffective.
Funnel ants
Identification: The two species of funnel ants can be distinguished by the shape of their entrance mound (Fig. 11.19) A. pythia is found in wetter area pastures, while A. longiceps is more frequent in drier areas. Worker ants of both species are yellow–brown in colour, 5 mm long and similar in size and shape to meat ants, Iridomyrmex spp. Risk conditions: Damaging populations can occur throughout the year. Funnel ants are an indicator of overgrazed and rundown pastures. Damage: Funnel ants create an artificial drought by removing the soil from around plant roots. In
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heavily grazed pastures, their building activities may cover the pasture with soil. The large entrances holes built by A. pythia (Fig. 11.19) cause nuisance by mound-building over areas such as tennis courts or around farm gates. Cultural control: Funnel ants are best managed by reducing grazing pressure and fertilising to maintain a dense pasture at least 30 cm high.
L EU C A E N A I N S E C T S
Leucaena psyllid Heteropsylla cubana Crawford Hemiptera: Psyllidae Distribution: Originally from tropical America, it is now distributed throughout the Pacific Islands and in Australia, where it occurs wherever leucaena is grown. Pest status: Major, restricted to leucaena, regular. Identification: The adults are aphid-like, 2 mm in length, winged and light green to yellow in colour (Fig. 13.6). They jump and then fly from the plant if disturbed. Eggs are laid on and in the unopened leaves and can be just seen with the naked eye. In large numbers they appear as orange–yellow masses. Nymphs are similar to adults in appearance except they are smaller, wingless and remain on the plant if disturbed. They only occur on the parts of the terminal with expanding leaves. May be confused with: Unlikely to be confused with any other insect on leucaena. Host range: Leucaena and leucaena hybrids.
Life cycle: In warm weather, the life cycle can be completed in 10–15 days. Psyllids prefer high relative humidity and temperatures in the 20s (oC). It is believed that the principal populationcontrolling agent for the species is temperature/ humidity. The population declines rapidly in mid-winter and during hot dry periods in late autumn–early summer. Risk period: April–May and/or September– October in inland Qld, year-round in coastal Qld when conditions are favourable. Damage: Adults and nymphs suck the sap of the terminal leaves, buds and flowers. If the population is low, leaves are puckered and fail to fully expand. With large populations, the expanding leaves and buds turn black and fall off leaving bare twigs. Flowering and hence seed-pod formation is prevented. Fully expanded leaves and mature stems and branches are not attacked. Where large numbers of psyllids are present, sooty mould may grow on the sugary psyllid excretions covering the foliage and stems preventing light reaching leaf surfaces. Annual dry matter (DM) yield losses in ‘coastal’ areas of Qld have been estimated at 50% each year and usually occur throughout the year except January to March. In western Qld, damage is less (ca. 10% DM) and usually occurs in April–May and/or September–October. Monitoring: Check five chest-height terminals at 30 positions over the whole area of leucaena. The infestation may be scattered. Record the number of terminals that are unaffected, distorted or have dead (or shed) leaves and buds due to psyllids. Action level: Spray if feed is limiting or if seed production is required and death of expanding leaves is occurring, particularly where more that 10% (15 out of 150) of terminals have dead leaves. It may be worth spraying seedling stands if a severe attack occurs as it is important that new stands are grown to a height of 3 m as quickly as possible. At this height they are able to compete with other plants, including grasses.
Long soft scale Coccus longulus (Douglas) Hemiptera: Coccidae Fig. 13.6. Leucaena psyllid adult (length 2 mm). (DPI&F Qld)
Distribution: This scale is distributed throughout the world’s tropics. 365
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Pest status: In Australia, it is a major, widespread and irregular pest on leucaena. Identification: The adult female long soft scale is an elongated oval in shape, 4–6 mm long, yellow to greyish-brown in colour, with visible eyespots (Fig. 13.7). May be confused with: It resembles other Coccus spp. scales such as soft brown scale (C. hesperidum Linnaeus), but is longer (up to 6 mm). Host range: In addition to Leucaena spp., hosts of the scale include many tropical fruit trees and ornamentals. Life cycle on leucaena: Adult females produce living young (crawlers) over a period of about 4 weeks. These crawlers disperse and settle on the stems, twigs and leaf stalks (petioles). They develop through two nymphal stages before reaching the adult stage. The complete life cycle takes about 2 months in summer. There may be four to six generations per year. Damage: The loss of sap through the sucking activities of the scale and the resultant honeydew and sooty mould may be severe enough to greatly reduce growth of leucaena. Monitoring: Monitor once in September– October. Using a ×10 hand lens, check 100 mm lengths of five randomly selected stems at chest height at 30 locations throughout the crop. Action level: If half (50%) of the stems have more than 30 scale, control is warranted. If the stems are infested a further sample in the following May will help in determining the potential for infestation the following summer.
Leucaena seed bruchid Acanthoscelides macrophthalmus Coleoptera: Bruchidae Distribution: Leucaena bruchid is recorded overseas as damaging a number of Leucaena spp. and was first found in Australia at Townsville in 1996 and is now found wherever leucaena is grown in Australia. Pest status: Minor (except for seed growers), found wherever leucaena is grown in Australia and occurs regularly.
Fig. 13.7. Adult female long soft scales encrusting a leucaena twig (length of longest: 6 mm).
Identification: Adults are short squat beetles, 2–3 mm in length, with large eyes and mottledbrown wing covers that do not extend to and cover the tip of the abdomen and enlarged hind leg femora. They fly readily when disturbed. Life cycle: Eggs (less than 1 mm long) are laid over a seed on the external surface of the pod or directly on an exposed dry seed. The young larva emerges and chews its way into the pod to the seed below. On emergence, the adult bruchid chews a circular hole in the seed and pod allowing it to escape. The life cycle is probably completed in 3 months during summer. Damage: The bruchid damages up to 95% of seeds produced.
Leucaena flower-eating caterpillar Ithome lassula Hodges Lepidoptera: Cosmopterigidae Distribution: The insect was first recorded in North America and found in Townsville in 1980. Pest status: A major pest for leucaena seed growers, it is currently found where leucaena is grown in Qld and occurs frequently. Identification: The adult has a wingspan of 7–10 mm; the forewing is narrow and has a long margin of hair-scales as long as or longer that the width of the wing. The head, thorax and forewing are shining black, sometimes with a few paler scales. Eggs are spindle-shaped, creamcoloured and measure about 0.5 mm in length. The caterpillars are creamy green in colour, up to 5 mm in length and. and have two thick plates on the thorax just behind the brown head.
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Weediness of leucaena contained by insects Following its initial introduction, leucaena became a pest along waterways due to its prolific growth. A number of host-specific insects have appeared in Australia in the late 20th century. They almost completely eliminate seed production and are expected to greatly reduce or eliminate leucaena’s weed potential once current seed soil reserves have germinated.
Life cycle on leucaena: Eggs are inserted between buds during the very early stages of the development of the flower head. They feed
at the base of the florets in the flower heads and mature; the caterpillar spins a case of silk and flower parts. It is capable of dragging this case about but usually remains within the flower head to pupate. As many as nine caterpillars can be found in one flower head. The pupa is about 3 mm in length. At Townsville the insect can be present throughout the year. Moths start laying eggs 2–3 days after emergence. About 10–16 eggs are laid during its lifespan of 5–6 days under laboratory conditions. Eggs hatch after 2–3 days. The caterpillar stage lasts 10–12 days. The pupal period lasts 8–9 days. Damage: Each caterpillar destroys a number of buds, which appear darker compared with undamaged buds. Ninety per cent damage to pods and seeds has been recorded.
Sources of information Anon. (2007). Additional pictures for these pests are available on the following website . Australian Plague Locust Commission website: Baker, G.L. (1993). Locusts and Grasshoppers of the Australian Region. Orthopterists’ Society and New South Wales Department of Agriculture, Rydalmere. Blight, V.C.N. (1966). The blackheaded pasture cockchafer. Division of Science Services, Entomology Branch, New South Wales Department of Agriculture. Broadley, R.H. (1978). The dayfeeding armyworm in north Queensland. Queensland Agricultural Journal 104: 27–30. Broadley, R.H. (1978). The lawn armyworm, a serious rural and urban pest. Queensland Agricultural Journal 104: 232–235. Broadley, R.H. (1979). Armyworms in southern Queensland field crops. Queensland Agricultural Journal 105: 433–443. Broadly, R.H. and Rogers, D.J. (1978). Pests of pangola grass in north Queensland pastures. Queensland Agricultural Journal 104: 320–324. Common, I.F.B. (1947). The yellow-winged locust, Gastrimargus musicus. BAgrSci thesis, University of Queensland. Common, I.F.B. and Beattie, W.M. (1982). An introduced moth, Ithome lassula (Cosmopterigidae) attacking Leucaena in northern Queensland. Journal of the Australian Entomological Society 21: 195–197. Champ, B.R. (1954). Control of grass caterpillars. Queensland Journal of Agricultural Science 11: 163– 164. Champ, B.R. (1955). The grass caterpillar. Queensland Agricultural Journal 80: 22–24. Elder, R.J. (1965). Webworms damage Northern Tablelands pasture. Queensland Agricultural Journal 91: 566–568.
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Elder, R.J. (1969). A flight study of the pasture soldier fly. Queensland Journal of Agriculture and Animal Science 26: 593–597. Elder, R.J. (1969). Flight studies of pasture webworms. Queensland Journal of Agriculture and Animal Science 26: 599–602. Elder, R.J. (1970). Larval taxonomy of Oncopera brachyphylla Turner and its distinction from Oncopera mitocera (Turner). Queensland Journal of Agriculture and Animal Science 27: 123–128. Elder, R.J. (1970). Distinction between adult Oncopera mitocera (Turner) and Oncopera brachyphylla Turner. Queensland Journal of Agriculture and Animal Science 27: 315–320. Elder, R.J. (1970). A rearing technique for Oncopera brachyphylla Turner and Oncopera mitocera (Turner). Queensland Journal of Agriculture and Animal Science 27: 401–404. Elder, R.J. (1971). Distinction between eggs and pupae of Oncopera brachyphylla Turner and those of Oncopera mitocera (Turner). Queensland Journal of Agriculture and Animal Science 28: 23–28. Elder, R.J. (1971). A study of distribution, morphology and biology of Oncopera mitocera (Turner) and Oncopera brachyphylla Turner (Lepidoptera: Hepialidae). MAgrSci thesis, University of Queensland. Elder, R.J. (1974). Oncopera brachyphylla Turner and Oncopera mitocera (Turner) insecticide control trials 1964–6. Queensland Journal of Agriculture and Animal Science 31: 279–284. Elder, R.J. (1976). Insecticide control of three grass feeding Noctuids and Herpetogramma licarsisalis (Walk.). Queensland Journal of Agriculture and Animal Science 33: 125–127. Elder, R.J. Brown, J.D. and Wicks, R. (1979). The biology and laboratory rearing of a new Leptopiine weevil, a pest of legumes in Queensland. Journal of the Australian Entomological Society 18: 81–89. Elder, R.J. (1983). Locusts and grasshoppers. Queensland Agricultural Journal 109: 189–191. Elder, R.J. (1989). Laboratory studies on the life history of Nomadacris guttulosa (Walker) (Orthoptera: Acrididae). Journal of the Australian Entomological Society 28: 247–253. Elder, R.J. and Mayer, D.G. (1990). An improved sampling method for Heteropsylla cubana Crawford (Hemiptera: Psyllidae) on Leucaena leucocephylla. Journal of the Australian Entomological Society 29: 131–137. Elder, R.J. (1991). Laboratory studies of environmental factors affecting sexual maturation in Nomadacris guttulosa (Walker) (Orthoptera: Acrididae). Journal of the Australian Entomological Society 30: 169–181. Elder, R.J. (1991). Effect of constant temperatures on egg development in Nomadacris guttulosa (Walker) and Locusta migratoria (L.) (Orthoptera: Acrididae). Journal of the Australian Entomological Society 30: 243–245. Elder R.J., Brough, E.J. and Beavis, C. (Eds) (1992). Managing Insects and Mites in Field Crops, Forage Crops and Pastures. Queensland Department of Primary Industries, Brisbane. Elder, R.J. (1995). Temporal incidence and sexual maturation of Austracris guttulosa (Walker) (Orthoptera: Acrididae) in Queensland, Australia. Journal of the Australian Entomological Society 34: 309–317. Elder, R.J. (1996). Morphometrics of field populations of Austracris guttulosa (Walker) (Orthoptera: Acrididae) in Australia. Australian Journal of Entomology 35: 345–347. Elder, R.J. and O’Brien, R.G. (1996). Ponded pasture pests and diseases. In: Beef production from ponded pastures. (Pittway, P.A., Wildin, J.H. and McDonald, C.K., eds). Tropical Grassland Society of Australia, occasional publication No. 7. Brisbane. pp. 50–51.
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Elder, R.J. (1997). Bionomics of Austracris guttulosa (Walker) (Orthoptera: Acrididae) during the 1970–5 outbreak in Queensland, Australia. Australian Journal of Entomology 36: 57–67. Elder, R.J, Middleton, C.H. and Bell, K.L. (1998). Heteropsylla cubana Crawford (Psyllidae) and Coccus longulus (Douglas) (Coccidae) infestations on Leucaena species and hybrids in coastal central Queensland. Australian Journal of Entomology 37: 52–56. Hunter, D.M. and Elder, R.J. (1999). Rainfall sequences leading to population increases of Austracris guttulosa (Walker) (Orthoptera: Acrididae) in arid north-eastern Australia. Australian Journal of Entomology 38 (3): 204–218. Madhavan, S.R. and Thakur, M.L. (1990). On some aspects of biology of Ithome lassula Hodges (Cosmopterigidae): A microlepidopteran pest introduced to India with the host Leucaena leucocephala (Lam.) De Wit. Indian Forester 116: 643–647. Moller, R.B. (1965). The soldier fly pest of sugar cane. Bureau of Sugar Experiment Stations, Queensland. Nambra, R. (1956). Descriptions of the immature stages and notes on the biology of Ithome concolorella (Chambers) (Lepidoptera: Cosmopterygidae), a pest of kiawe in the Hawaiian Islands. Proceedings of the Hawaiian Entomological Society 16: 95–100. Quinlan, T.J, Elder, R.J. and Shaw, K.A. (1972). Webworm control in northern dairy pastures. Queensland Agricultural Journal 28: 176–7. Quinlan, T.J, Elder, R.J. and Shaw, K.A. (1975). Pasture dry matter reduction due to Oncopera spp. (Lepidoptera: Hepialidae). Australian Journal of Experimental Agriculture and Animal Husbandry 15: 219–222. Saunders, G.W. (1968). Funnel ants (Aphaenogaster spp., Formicidae) as pasture pests in North Queensland, II, Control. Bulletin of Entomological Research 59: 281–290. Samson, P.R. (1995). Management of soldier flies in sugarcane. Proceedings of the Australian Society of Sugar Cane Technology 17: 75–82. Smith, D., Beattie, G.A.C. and Broadley, R. (Eds) (1997). Citrus pests and their natural enemies. Queensland Department of Primary Industries and Fisheries, Brisbane. Zborowski, P. (1998). Field Guide to the Locusts and Related Grasshoppers of Australia. Australian Plague Locust Commission, GPO Box 858, Canberra, ACT 2601.
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14 PASTURES—SUMMER RAINFALL: WEED BIOCONTROL AGENTS R.E. McFadyen
(Left) Introduced from South America, common prickly pear invaded 100 000 square km of Queensland grazing land similar to that in this 1928 photo. (Right) The same area 18 months after the introduction of cactoblastis.
WEED
ESTABLISHED AGENT (major control agents in bold)
annual ragweed
variegated ragweed beetle parthenium stemgalling moth parthenium stemgalling moth psyllid groundselbush stemborer groundselbush leafeating beetle groundselbush gall fly groundselbush stemboring moth groundselbush leaf-tying moth groundselbush leafmining moth harrisia cactus mealybug lantana mealybug lantana sucking bug lantana lace bug lantana leafmining beetles lantana leafmining fly lantana seed fly catabena moth hypena moth lantana flower caterpillar lantana leaf-folding caterpillar lantana plume moth leaf-tying moth noogoora burr longicorn parthenium stemgalling moth
Bathurst burr giant sensitive plant groundsel bush
harrisia cactus lantana
mesquites noogoora burr
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WEED
ESTABLISHED AGENT (major control agents in bold)
parkinsonia
parkinsonia leaf bug parkinsonia seed beetles variegated ragweed beetle parthenium stemboring weevil parthenium seed-feeding weevil parthenium stem-galling moth parthenium leaf-mining moth prickly acacia seed beetle cochineal scales cactoblastis cactus moth leaf-feeding moth
parthenium weed
prickly acacia prickly pears
rubber vine
Summer rainfall pastures range from high value pastures in high rainfall areas on rich volcanic soils east of the dividing range in Qld and NSW, to low value rangelands in western Qld and NSW, and include grazing lands on good brigalow soils in central Qld. These areas are characterised by variable rainfall, falling mostly in a few heavy rainfall events in the summer months with odd amounts of rain throughout the year. Most weeds of summer pastures were deliberately introduced as ornamentals or in misguided attempts at pasture improvements. In the better soils and higher rainfall areas, weeds are controlled by slashing, occasional cultivation, and the use of herbicides where necessary. On steeply sloping country and in the poorer rangeland pastures, mechanical control is not possible and the use of herbicides is often uneconomic. Consequently, biological control may be the only economically viable method of management. Biological control of weeds in these areas has been well accepted since the success of prickly pear control in the 1930s. Nearly all biocontrol programs on weeds of summer pastures were undertaken by the Queensland government at the Alan Fletcher Research Station in Sherwood, Brisbane, the original site of the prickly pear biocontrol program in 1920 which has been an active biocontrol research station ever since (Walton 2005).
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Australia. In Australia it occurs in coastal Qld and NSW east of the Dividing Range from Gympie to Port Macquarie, with an isolated infestation in the Atherton Tablelands.
Variegated ragweed beetle Zygogramma bicolorata Pallister Coleoptera: Chrysomelidae Distribution: Native to Mexico, this beetle was first introduced to Qld in 1980 and subsequently redistributed to NSW. It is now present throughout the distribution of annual ragweed. Beneficial status: Significant but varies locally and seasonally. Populations are reduced by predation and by entomopathogenic fungi in the soil. Identification: Adults are large (8–10 mm) beetles, white with a black pattern. Larvae are slug-like, white–green in colour, and often hide at leaf bases during the day. (See figures in parthenium weed, page 386.). May be confused with: Unlikely to be confused with any other insect. Host range: Restricted to parthenium and annual ragweed. Life cycle: See parthenium weed.
Annual ragweed Ambrosia artemisiifolia Asteraceae Originally from eastern North America, now present in the Ukraine, southern China, and
Active period: On annual ragweed, adults emerge in spring after rain, later in the south. Adult-feeding and oviposition continues until March, when annual ragweed flowers. Damage: See parthenium weed.
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Integrated control: Herbicides only affect very young larvae.
a
Parthenium stemgalling moth Epiblema strenuana Walker Lepidoptera: Tortricidae Distribution: Native to Mexico and eastern North America as far as Ontario, Canada, this moth was first introduced to Qld in 1982 and subsequently redistributed to NSW. Beneficial status: Major in northern NSW and Qld, minor on the New England Tablelands and other colder areas.
b
Identification: See parthenium weed. May be confused with: Unlikely to be confused with any other ragweed insect. Host range: The closely related genera Ambrosia, Parthenium and Xanthium. Life cycle: See under parthenium weed. Active period: Adult moths emerge in late August or early September. Larval development occurs during October and the spring generation of adults emerges in late October or early November. Generational overlap then occurs, with adults continuously present until the plant begins to flower in March or April, at which time larvae enter pre-pupal diapause and overwinter in the dry stalks. In cold areas, there may only be one summer generation and one overwintering generation, while in warmer areas there may be three or four generations during the summer (McFadyen 1984, 1989). Damage: Small larvae produce galls on stems (Fig. 14.1), reducing the vigour of the plant, and larger larvae mine the stem (Fig. 14.1). Integrated control: Any destruction of the dry stalks destroys the overwintering larvae. Herbicides do not damage larger larvae or pupae.
Bathurst burr
Fig. 14.1. (a) Annual ragweed plant with galls caused by larvae of parthenium stemgalling moth, and (b) stem damage caused by a late-stage larva. (QNRM: M.W. Lee)
introduced insects may sometimes be found on this weed, they do not provide useful control. Parthenium stemgalling moth, which also attacks parthenium weed and noogoora burr, is occasionally found on Bathurst burr.
Giant sensitive plant Mimosa invisa Mimosoidea Distribution: A native of Brazil, this plant now occurs in Africa, South-East Asia, Papua New Guinea, some Pacific Islands and Australia, where it appears to have been accidentally introduced in about 1929. It is now occasionally present as a weed in tropical pastures along the coast of far north Qld.
Psyllid
Xanthium spinosum Asteraceae
Heteropsylla spinulosa Muddiman, Hodkinson & Hollis Hemiptera: Psyllidae
A native of South America, Bathurst burr has now spread to most continents including Australia, where it has been recorded in all states. Although a number of native and
Distribution: Imported from Brazil and released in Qld during 1988. Later taken to Papua New Guinea for biological control of giant sensitive plant. 373
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Distribution: Native to southern Brazil, where its host plant is Baccharis microdonta DC, it was introduced into Qld in 1978. Redistribution into NSW was not successful, and it is restricted to swampy lands in Moreton Bay and adjacent islands. Beneficial status: Very damaging where found, but very restricted distribution. Fig. 14.2. Adult psyllid (to the right of the match head) on giant sensitive plant. (QNRM)
Beneficial status: Major, widespread in Qld, regular.
Identification: Adult beetles are large (13– 18 mm long) and red and black in coloration (Fig. 14.3). Larvae within the stem are yellowcoloured and long and narrow.
Description: Adult psyllids are about 2.5 mm long and green (Fig. 14.2).
May be confused with: Adults are unlikely to be confused with any other insects on this plant; larvae may be confused with the stemboring moth but the mouth parts are distinctive.
Host range: H. spinulosa feeds only on giant sensitive plant.
Host range: Restricted to Baccharis species, of which only B. halimifolia occurs in Australia.
Life cycle: Adults congregate on young foliage where they feed and mate 1–2 days after the moult to adults. Egg-laying begins 1 day after mating. Eggs are about 0.5 × 2 mm and attached by a small stalk to leaflets. The eggs hatch to nymphs, which develop through five nymphal stages before moulting to adults. The total development time is 15–22 days. There may be up to eight generations per year (Willson and Garcia 1992).
Life cycle: There are no distinct generations, and all stages may be found at all seasons. Adult beetles do not feed, and survive for about 3 weeks. Eggs are laid singly into cracks in the bark on large stems and the emerging larvae feed by tunnelling just below the bark, in the cambium layer. Large larvae feed into the wood, and frass is ejected through one or more holes. Pupation takes place in the larval tunnel. The entire life history takes 4–9 months.
Damage: Feeding of adults and nymphs stunts and distorts leaves, prevents flowering and destroys young plants.
Groundsel bush Baccharis halimifolia Asteraceae
Active period: All stages are found throughout the year, although activity is lowest in the winter. Damage: Larval-feeding severely damages affected stems and may kill whole stems and even plants.
Originally from Texas and Florida, USA, now found in Europe (southern France, Ukraine) and Australia. In Australia, groundsel bush occurs in coastal Qld and NSW east of the Dividing Range from Gladstone to Port Macquarie. Prior to 1990, groundsel bush was a very serious weed of pastures in northern NSW and coastal Qld. Land use changes, a major chemical control campaign and biocontrol have greatly reduced the abundance.
Groundsel stemborer Megacyllene mellyi (Chevrolat) Coleoptera: Cerambycidae
Fig. 14.3. Adult groundsel bush stemboring beetles, 15 mm long. (QNRM)
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Integrated control: Burning or slashing destroys the larvae but larger larvae or pupae are unaffected by herbicides.
Groundsel leafeating beetle Trirhabda baccharidis (Weber) Coleoptera: Chrysomelidae Distribution: Native to Florida, USA, the beetle was first introduced into Qld in 1969. Redistribution to NSW failed, and the beetle is restricted to swampy lands in Moreton Bay and the bay islands. Beneficial status: Minor, very restricted distribution. Identification: Larvae are black and slug-like, up to 10 mm long (Fig. 14.4). Adults are black and yellow, shiny and 7–12 mm long (Fig. 14.4). May be confused with: Cannot be confused with any other insect on this plant. Host range: Restricted to Baccharis species. Life cycle: There is one generation per year. Adults emerge in October and commence feeding on the leaves. Eggs are laid in crevices in bark but pass the summer in diapause, hatching in autumn. Young larvae feed on new leaves. Mature larvae pupate in the soil. Adult emergence is triggered by rainfall. Active period: From April to November. Damage: Large populations can leave plants completely defoliated. Regrowth occurs but affected plants produce few flowers or seeds, and seedlings or small plants may be killed. Integrated control: Burning during the larvalfeeding period will destroy larvae, but can be
Fig. 14.4. Adult of the groundsel bush leaf-eating beetle, 10 mm long. (QNRM)
carried out when pupae are present in the soil. Herbicide treatments do not kill the beetle.
Groundselbush gall fly Rhopalomyia californica Felt Diptera: Cecidomyiidae. Distribution: Native to California, USA, where its host plant is Baccharis pilularis DC. First released in Qld in 1982, it has been redistributed to NSW. Beneficial status: Significant, widespread, regular in cooler and shady locations; minor, restricted in open hot sunny locations where it is heavily parasitised by native parasitoids. Identification: Galls are large (1–3 cm diameter), containing orange–red larvae or pupae. Adults are seldom seen but are small (3–5 mm long) with orange–red bodies and transparent wings (Fig. 14.5). May be confused with: Unlikely to be confused with any other insects on groundsel bush. Host range: The gall fly is restricted to Baccharis species, of which groundsel bush is the only species occurring in Australia. Life cycle: The gall fly has many generations per year and all stages occur at all seasons. The adult flies live only a single day, emerging midmorning, mating immediately and then laying the eggs into growing stem tips or axillary buds. The eggs are oval, 1 mm long, and yellow– orange in colour, and several are laid in a single tip, partially inserted into the tissue (Fig. 14.5). On emergence, the larvae bore into the meristem tissue, which reacts by swelling into a roughly
Fig. 14.5. Adult tip gall fly, 5 mm long, laying eggs onto groundsel bush. (QNRM) 375
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a
Fig. 14.6. A gall on a groundsel shoot caused by the tip gall fly. (QNRM) b
spherical gall. Larvae develop in separate tunnels within the gall. When full-grown, larvae cut an emergence ‘window’ leaving only the epidermis layer intact, and pupate within the tunnel. Adults emerge through the ‘window’. The whole life cycle takes about 20 days in summer and proportionally longer in cooler weather. Active period: All stages are active all year, although development is longer in winter and adult emergence is stimulated by sunshine and higher temperatures. Damage: Gall development acts as a metabolic ‘sink’, reducing the supply of metabolic products to other growing tissues (Fig. 14.6). The impact of a single gall may be small but numerous galls reduce stem growth and seed production. Affected seedlings and regrowth after slashing or burning may die, as may heavily galled stems. Integrated control: The tip gall fly is particularly effective in shady or cool locations, and against regrowth developing after slashing or burning, although burning destroys the insect. Herbicide applications do not affect the insects.
Fig. 14.7. (a) Adult groundsel bush stemboring moth (wingspan 35 mm), and (b) eggs laid in leaf axil. (QNRM)
from releases in 1985 and has since been redistributed to NSW. Beneficial status: Major, widespread, regular throughout the distribution of the weed. Identification: Adult moths are large (wingspan 30–40 mm, Fig. 14.7) and may be seen flying at lights at night. They lay white, ovoid eggs in axils (Fig. 14.7). Frass is extruded externally from the larval tunnels and is a good sign of activity (Fig. 14.9). Larvae are thin, yellow–white, up to 25 mm long (Fig. 14.8). Death of 4–5 cm of stem tips is a characteristic symptom. May be confused with: Adults cannot be confused with any other insects; larvae and
Groundselbush stemboring moth Oidoematophorus balanotes (Meyrick) Lepidoptera: Pterophoridae Distribution: Native to the south-eastern USA, it was first released in Qld in 1969 but the releases failed. It was successfully established in Qld
Fig. 14.8. Stem split to show larva, 15 mm long, in channel. (QNRM)
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Fig. 14.9. Frass ejected from larval tunnel is indicative of larval damage. (QNRM)
their frass or tunnels may be confused with Megacyllene larvae but the moth is much more common. Host range: Restricted to plants in the genus Baccharis, of which only B. halimifolia occurs in Australia. Life cycle: The moth does not have separate generations and all stages may be found all year. Adult moths are nocturnal and may feed at flowers or sugar sources but only live 14 to 21 days. Eggs are laid singly on leaves and the larvae feed singly in the stem tips, killing 5–6 cm of stem and causing characteristic damage. Larvae then move down into the main stem and tunnel in the pith, expelling frass through one or more holes. Mature larvae pupate in the tunnel and adults emerge through emergence windows. The complete life cycle takes 6–8 months. Active period: All year, although adult emergence may be more common in the warmer months. Damage: Death of stem tips is common and damaging. Larger larvae do not kill stems but weaken them so that breakage occurs from wind or other causes. Heavily affected plants are stunted and produce few flowers or seeds. Integrated control: Slashing or burning will destroy the larvae in the stems. Herbicide application results in death of small larvae but not larger ones.
Fig. 14.10. Adult groundsel bush leaf-tying moth (body is 10 mm long). (QNRM)
subsequently redistributed into NSW, and is now common in both states. Beneficial status: Minor. Identification: Adults are small (7–10 mm) and pale brown (Fig. 14.10). Larvae feed within a cocoon of leaves tied together, leaving irregular eaten holes in the leaves. May be confused with: Moths are seldom seen. Larvae cannot be confused with any other insect on this plant, and larval-feeding damage is distinctive. Host range: Restricted to Baccharis species. Life cycle: There are no separate generations and all stages are present all year. Moths lay the minute (< 1 mm) pale green eggs singly on the leaf under-side, and the newly-hatched larvae initially mine in the leaf, between the leaf surfaces. Larger larvae tie two or more leaves together and live between, emerging to feed by eating irregular holes in adjacent leaves. Pupation occurs in the larval refuge. The life history takes about 5 weeks. Active period: All year. Damage: Larval-feeding damages leaves, but populations are never sufficiently large to cause significant damage. Integrated control: Not relevant as damage from this moth is not significant.
Groundselbush leaf-tying moth Aristotelia sp. Lepidoptera: Gelechiidae
Groundselbush leafmining moth
Distribution: Native to Florida, USA, this moth was first introduced to Qld in 1969 and
Bucculatrix ivella Busck Lepidoptera: Gracilaridae 377
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Distribution: Native to Florida, USA, the moth was first introduced to Qld in 1985 and subsequently redistributed to NSW. It is now widespread in both states. Beneficial status: Minor. Identification: Adult moths are tiny (4 mm long) and silver–grey in colour. Larval mines are thread-like and irregular. May be confused with: Cannot be confused with any other insect found on this plant. Host range: Restricted to Baccharis species. Life cycle: There are several generations a year, and all stages present at all times. Adults lay minute (< 0.5 mm) eggs singly on the leaf underside. On hatching, the larvae commence mining the leaf. Large larvae leave the mines to feed directly on the leaves, making characteristic square feeding holes. Pupation occurs inside a ribbed white silk cocoon attached to stems or the leaf under-side. The complete life cycle takes 6–8 weeks. Active period: All stages are present throughout the year, although growth is slower in winter. Damage: Larval-feeding damage to the leaves is seldom sufficient to affect plant growth.
Harrisia cactus Eriocerus martinii and E. tortuosus Cactaceae Native to northern Argentina and Paraguay, harrisia cactus is now weedy in South Africa and Australia, where it is widespread in Qld and NSW and also occurs in WA. Prior to successful biocontrol, Harrisia cactus was a major weed of pastures in brigalow soils in north and central Qld, displacing grasses to reach up to 80% cover and reducing pasture productivity to near zero. Major Qld and NSW government programs used chemical herbicides to reduce impact and prevent spread. Since biocontrol was achieved in the late 1970s, these cacti have become minor components of ground vegetation in brigalow soils from north to central Qld, and are occasionally more abundant in southern Qld and northern NSW.
Distribution: Native to Argentina, Paraguay and Bolivia, this species has been introduced into Australia and South Africa. Beneficial status: Major over most of range, less effective in northern NSW and southern Qld where cold winters restrict development. Identification: Adult female grows up to 2 mm long and 4 mm wide with the body covered by a white, woolly covering. Eggs are 0.44 mm long, the first-stage nymph is 0.25–0.27 mm, the second stage is 0.45–0.65 mm and the third stage is 0.75–0.88 mm long. Males have a ‘pupal’ stage, 0.75–0.88 mm long, which hatches to a winged male 0.88–1.0 mm long. All stages are found in dense colonies around buds, including flower buds and young growth (Fig. 14.11). Mealybugfeeding distorts the cactus growth to form dense matted clumps of twisted stems, often resembling a clenched fist at the stem tip or axillary bud. The mealybug colonies live deep within these characteristic growth masses. Host range: Most cacti in the sub-family Cereanae: sword pear, night-flowering cereus, but not the common epiphytic moonlight cactus (Hylocereus). Life cycle: There are up to eight generations per year in the north, as few as three in northern NSW. Some 20 days after fertilisation, the female lays two to four eggs per day for up to 35 days. Eggs hatch to crawlers within 20 minutes. The three female immature stages have a mean duration of 28 days up to the final moult to adult, and the immature males take a mean of 30 days to develop into adults. Adult females live for a mean of 60 days.
Harrisia cactus mealybug Hypogeococcus festerianus (Lizer y Trelles) Hemiptera: Pseudococcidae
Fig. 14.11. A dense colony of cactus mealybugs on harrisia cactus. (QNRM)
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Active period: Summer.
Beneficial status: Minor, restricted, irregular.
Damage: Colonies of the mealybug arrest new growth and prevent flowering (Fig. 14.11). Severe mealybug attack over 3 years will kill established plants.
Identification: Adult female mealybugs are 2–3 mm long, with a white to cream waxy covering secreted by the insect.
Management: The mealybug relies on passive dispersal through the crawlers, which are windblown onto new plants. Manual redistribution into new patches of cactus may be necessary: in early summer, clumps of distorted growth containing live mealybugs are cut off, transported to new sites, and scattered among the cactus plants. Herbicidal control of cactus does not integrate well with the use of the mealybug. Chemical control kills the larger plants, which carry large mealybug colonies, while often missing small plants and seedlings. This slows the mealybug population growth and spread. For rapid results, it is best to spread the mealybug densely, infesting every plant, and only use chemicals on plants in dense shade where low daytime temperatures may slow mealybug growth (McFadyen 1979).
Lantana Lantana camara Verbenaceae Lantana is a native of tropical Central and South America, introduced to most tropical and subtropical countries worldwide, including Australia, as an ornamental. Lantana is a major weed of summer rainfall pastures along coastal NSW and Qld. Biological control of the weed in Australia has included the introduction of at least 29 species of insects, of which 15 are recorded as having established (Day et al. 2003).
Host range: Includes solanaceous crop plants, such as potato and tomato, as well as other plant families, but has not been recorded as a pest of crop plants in Australia. Life cycle: The female produces eggs parthenogenetically, and the eggs hatch to firststage crawlers which disperse to the under-side of new leaves where they insert their mouth parts and feed on the sap of the lantana. The second (nymphal) stage and the third (adult) stages may move between feeding sites. Damage: Feeding on lantana plants by small to medium populations of mealybugs may weaken plants, and high populations cause dieback in local areas.
Lantana sucking bug Aconophora compressa Walker Membracidae Distribution: Introduced to Australia from Mexico, now established at several sites in coastal NSW and Qld. Beneficial status: Minor, restricted, irregular. Identification: Adult bugs are 3–6 mm long, brown in colour, with a pronounced thoracic projection resembling a thorn (Fig. 14.12). They sit along stems where they are very well camouflaged. Unlikely to be mistaken for any other insect. Host range: Lantana and fiddlewood trees. Duranta (Duranta repens) may occasionally also be attacked.
Of those established, none are recorded as controlling lantana, eight offer partial control, six inflict minor damage and it is presently too early to judge one species. In addition, a leaf rust, Prospodium tuberculatum, Uredinales Puccinaceae, from Brazil, is established at a few sites.
Lantana mealybug Phenacoccus parvus Morrison Pseudococcidae Distribution: A native of Brazil, was apparently introduced accidentally to Australia.
Fig. 14.12. Acanophora compressa with eggs. (AFRS) 379
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Life cycle: Eggs are laid in a mass adhering to the stem and guarded by the female. Nymphs feed on stems in groups, moving down the stem as they grow. Adults disperse to find new stems for oviposition.
Qld but has established only in northern Qld. The adult of this bug is 2–3 mm long with transparent, lacy forewings projecting back beyond the body. Where it occurs, it causes leaf damage and defoliation.
Damage: Feeding by large numbers of these sap-sucking insects causes defoliation, dieback of branches and reduced flowering.
Lantana leafmining beetles Chrysomelidae
Lantana lace bug Teleonemia scrupulosa Stål Hemiptera: Tingidae Distribution: Originating from Mexico, this species was introduced to Australia in 1936, and is the most effective of the lantana biocontrol insects yet introduced to Australia. Beneficial status: Major in the north, seasonal. Identification: Adult bugs are 2–3 mm long, light brown in colour with lace-like venation on the forewing (Fig. 14.13). Host range: The species has been recorded on a number of Lantana spp. and also Myoporum spp. Life cycle: These insects have 10–12 generations per year, each generation lasting about 1 month. Colonies of adults and nymphs may be found on the under-side of leaves. Damage: Sap-sucking by these bugs may cause chlorotic lesions, leaf malformation, defoliation and perhaps systemic poisoning by salivary secretions of the bugs. When bug populations are large, seasonal defoliation combined with environmental stress and damage by other agents may kill plants.
Octotoma championi Baly Octotoma scabripennis Guérin-Méneville Uroplata fulvopustulata Baly Uroplata girardi Pic Beneficial status (all species): Minor, restricted Australian distribution, irregular occurrence. O. championi is a native of Central America and southern North America introduced to Australia in 1954. The species is presently established in small areas of north Qld where it causes minor damage to lantana. O. scabripennis is a native of Central America, and was released in Australia in 1966, where it provides partial control of lantana in Qld. Feeding causes defoliation during summer, reduced flowering and loss of vigour, but does not kill the plant. Adult beetles feed on the upper surface of leaves and lay eggs singly beneath the leaf epidermis (Fig. 14.14). Up to four larvae may mine a leaf (Fig. 14.14) but their a
A second tingid lace bug, Leptobyrsa decora Drake, has been released widely in NSW and
b
Fig. 14.13. Adult (right) and nymphs of Teleonemia scrupulosa. (QNRM)
Fig. 14.14. (a) Adult O. scabripennis, and (b) larval leaf mines and damaged bush. (QNRM)
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a
b
Fig. 14.15. (a) Adult U. girardi, (b) larval mines and damage to lantana. (QNRM)
mines do not extend to other leaves. Development from egg to adult takes 34–45 days and the female takes a further 3–4 weeks to produce eggs. There are normally three generations per year. U. fulvopustulata is native to Central America, was introduced to Australia in 1976 where it established in small patches in tropical Qld but does not provide useful control. U. girardi is native to Central and South America. The species was introduced to Australia from Brazil in 1976. In Australia, it is established and partially controls lantana along the coasts of Qld and NSW. Adults feed on the upper leaf surface and scarify areas of the leaf tip (Fig. 14.15), causing leaf curl. Larvae mine the leaves of lantana at the rate of one or two mines per leaf (Fig. 14.15). The life cycle takes 31–52 days and there are normally three generations per season.
Lantana leafmining fly Calycomyza lantanae (Frick)
Fig. 14.16. Adult lantana leafmining fly and larval mines on leaf. (QNRM)
Both of these species are natives of southern North America, Central America and northern South America. C. lantanae was introduced to Australia in 1974 and is now widespread in coastal Qld where it provides partial control of lantana through leaf damage caused by larvae mining leaves. O. lantanae damages fruit but its impact on seed viability is uncertain.
Catabena moth Neogalea sunia (Guenée) Noctuidae This species occurs naturally from southern North America to southern South America. It was established in Australia in the 1950s but is uncommon and does not contribute to the control of lantana.
Hypena moth Hypena laceratalis Walker Noctuidae This species is a native of Africa and appears to be specific to Lantana spp. It has been in Australia since prior to 1965 and has some impact on lantana in Qld, causing seasonal defoliation of lantana together with other insects. The adult moth lays eggs on the lower surface of leaves where the hatched larvae feed for about 12 days. Under favourable conditions, populations of this moth can increase rapidly.
Lantana flower caterpillar Epinota lantana (Busck) Tortricidae
Lantana seed fly Ophiomyia lantanae (Froggatt) Diptera: Agromyzidae
A native of Central America, this species was introduced to Australia in 1914, where it is established along the east coast. Larvae damage shoots and flower heads of lantana but 381
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populations are always small and control negligible.
Lantana leaf-folding caterpillar Salbia haemorrhoiodalis Guenée Pyralidae A native of southern North America and Central America, this species is established in Australia but has little impact. Larvae feed on leaves, which they fold together with silk.
Lantana plume moth Lantanophaga pusillidactyla (Walker) Pterophoridae This species is a native of Central America. Its introduction to Australia appears to have been accidental. It is common over most of the range of lantana, where larvae eat flowers but do not provide any control of the weed.
Mesquites Prosopis spp. Mimosoidea Mesquites are native to southern North America, Central America and northern South America. Introduced to a number of countries as stock food, mesquites are now woody weeds in parts of Asia, Africa, South America and Australia, where they were introduced in the 1930s. In Qld and WA, they are regarded as serious weeds of semi-arid rangelands, and also occur as pastoral weeds in parts of all mainland states and NT.
Life cycle: Female moths lay eggs into cracks and fissures of the plant. First-instar larvae mine within leaves and subsequent instars defoliate the bush by tying leaves together. The psyllid Prosopidopsylla flava (Hemiptera: Psyllidae) is reported as established in southwest Qld but populations are very small and damage to the weed negligible.
Noogoora burr Xanthium occidentale, Asteraceae A native of North America, noogoora burr has now spread to Asia, Africa, Europe and Australia. In Australia, where it was accidentally introduced during the 1860s, it is now present in all mainland states and is a major weed in NSW, Qld and Vic. A number of insects have been introduced as biocontrol agents with variable success. The rust Puccinia xanthii, illegally introduced to Australia in 1974, provides excellent control in central and southern Qld but not in the far west or far north.
Noogoora burr longicorn Nupserha vexator (Pascoe) Coleoptera: Cerambycidae First introduced and released in the early 1960s, this stemborer has become established and widespread in central Qld. Beneficial status: Minor, restricted to central Qld.
Parthenium stemgalling moth Leaf-tying moth Evippe sp. Lepidoptera: Gelechiidae Distribution: Imported from Mexico, this moth is now well established in the Pilbara, northwestern WA. Beneficial status: Major in WA where it is widespread and regular in occurrence. Damage: Defoliates the plant and reduces seed production. Host range: In Australia, Leucaena leucocephala may also host small populations (Klinken et al. 2003).
Epiblema strenuana (Walker) Lepidoptera: Tortricidae Introduced to control parthenium weed, this species also provides partial control of noogoora burr (Fig. 14.17) in parts of its range. See parthenium weed.
Parkinsonia Parkinsonia aculeata Caesalpiniaceae Parkinsonia is a native of southern North America, Central America and northern South America, and introduced to Australia as an
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Distribution: P. germaini was imported from Argentina and released in Qld in 1995 and is now established in NT, Qld and WA where spread was rapid. M. ulkei was imported from the USA and was first released in 1993. It is established at several sites, but did not establish as readily as P. germaini. Beneficial status: P. germaini: minor, widespread, irregular. Previously recorded as destroying a significant proportion of seeds, its effectiveness is now limited by a native parasitoid. M. ulkei: Minor, restricted, irregular. Identification: P. germaini is a small (5–6 mm long) brown beetle. M. ulkei is a small (about 5 mm long) two-tone grey beetle (Fig. 14.18). Fig. 14.17. Stem of noogoora burr split to show channel and a pupa (arrowed) of parthenium stem galling moth. This moth also damages other weeds. (QNRM)
ornamental. It is now a woody weed in semi-arid rangelands of northern NSW, NT, Qld and northern WA. Biological control by seed beetles initially was effective but control is now inadequate and searches for new agents continue.
Parkinsonia leaf bug Rhinacloa callicrates Herring (Hemiptera: Miridae) was released in Qld in 1988 and NT in 1989 from a culture derived from Arizona. The leaf bug has only established in Qld, where it is widespread but does not result in control of the weed. Adults and nymphs feed on the green tips and foliage of the weed, leaving white spots around feeding punctures and black faecal spots on the leaf surface. The adult inserts a cigarshaped egg under the epidermis of the leaf rachis, producing a small blister. After hatching, the nymphs develop through five stages, with the fourth and fifth nymphal stages having wing buds. Adults fly and are attracted to lights. The life cycle is completed in about 3 weeks (Donnelly 2000).
Host range: Restricted to the genus Parkinsonia. Life cycle: The life cycle of P. germaini includes about 9 days in the egg stage, 22 for the four larval stages, and eight for the pupal stage. Females lay eggs singly on the outside surfaces of pods and can lay up to 350 eggs each. Eggs are white and can be seen against the darker pods. Female M.ulkei beetles lay clusters of eggs in cracks and holes in the pods. Larvae of both species tunnel into seeds after hatching, and each larva spends its entire development period in the same seed where usually only one larva survives. The larva pupates in the seed, having eaten all the living contents of the seed rendering it unviable. The new adult emerges through a hole at one end of the seed and a hole in the side of the pod, and these round holes in the pods indicate where beetles have emerged. The life cycles of both species range from about 5 weeks in the hot months of the year to about 12 weeks in the winter. Individual beetles of
Parkinsonia seed beetles Penthobruchus germaini (Pic) and Mimosestes ulkei (Horn)
Fig. 14.18. Adult Mimosestes ulkei (5 mm long). (C. Wilson)
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both species can live for up to 2 months but more typically they live for about 5 weeks. Larvae overwinter in the seeds on the ground, and begin pupating in late winter, with adults emerging in spring (Briano et al. 2002). Damage: Larval-feeding destroys seeds of parkinsonia. P. germaini was initially recorded as destroying up to 95% of seeds, but it is no longer effective in control of the weed.
a
b
Parthenium weed Parthenium hysterophorus Asteraceae Parthenium weed is originally from the Gulf of Mexico (Mexico and southern USA), and is now found in India, Taiwan, east Africa, Madagascar, and Australia. In Australia, parthenium weed is found in north and central Qld, chiefly in rangelands in the 500–1000 mm rainfall zone, but increasingly on the coast and south to the NSW border. A total of nine insect species were introduced during a biocontrol program from 1978 to 1995, of which only one failed to establish, although three are still rare. Two rust diseases (Puccinia melampodii and Puccinia abrupta) were also introduced, and rust damage can be seen in wet situations such as along creeks or after good rain. Control has been very successful, resulting from the combined effects of several agents acting in different seasons and environments (Dhileepan 2001; McFadyen 1992).
Variegated ragweed beetle Zygogramma bicolorata Pallister Coleoptera: Chrysomelidae
Fig. 14.19. (a) Egg and (b) slug-like larva of parthenium leaf-feeding beetle. (QNRM)
May be confused with: Unlikely to be confused with any other insect found on parthenium. Host range: Parthenium and annual ragweed. Life cycle: There are two to three generations during summer if rainfall is adequate. Adults live for at least 12 months, spending the dry season in diapause in the soil, emerging after rainfall. After a period of feeding, the small (1 mm long) yellow eggs are laid in batches of two to 10 on the under-side of leaves. Larvae initially feed on young leaves and then on older leaves, hiding among the leaf folds during the day. Mature larvae burrow into the soil and construct small cells 2–8 cm below the surface in which they pupate. Adult emergence does
Distribution: Native to Mexico, this beetle was introduced into Qld in 1980. It became widely established on parthenium from 1995 and is now present in central Qld from Clermont south. Beneficial status: Major in most seasons where present. Identification: Eggs (Fig. 14.19) are yellow and elongate. Larvae (Fig. 14.19) are greenish-white and slug-like, often hiding during the day. Adults (Fig. 14.20) are large (8–10 mm) ‘ladybird’ beetles, black with white markings.
Fig. 14.20. Parthenium leaf-feeding beetle adult, body 10 mm long. (QNRM)
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not occur until after soaking rains have moistened the soil. The life cycle from egg to adult is 6–8 weeks but may be greatly increased by diapause.
a
Active period: Adults are active during summer so long as green plant is present. During winter (from May on) and in dry seasons, they are in diapause in the soil. Eggs and larvae may be found whenever adults are present. Damage: Feeding by larvae and newly-emerged adults can completely strip leaves from parthenium plants over large areas. Young stem tissue is also eaten, and plants are usually killed.
b
Integrated control: Deep ploughing kills diapausing adults and pupae. Herbicides only affect young larvae.
Parthenium stemboring weevil Listronotus setosipennis (Hustache) Coleoptera: Curculionidae Distribution: Native to southern Brazil, Paraguay and north-west Argentina, this weevil was introduced into Qld in 1983 and is now widely distributed. Beneficial status: Significant in most seasons. Identification: Larvae are white, curved, up to 8 mm long, found within tunnels in stems or just under the bark on roots (Fig. 14.21). Adults are nocturnal, hiding in soil and leaf litter during the day. They are 5–7 mm long, and grey in colour (Fig. 14.21). May be confused with: Larvae may be confused with Epiblema larvae but are whiter and more curved. Host range: Restricted to Parthenium species. Life cycle: There are no separate generations and all stages may be found in any season. Adults spend the dry season buried in soil and leaf litter, and emerge following rain to feed on the leaves, making small circular feeding holes. The small clear eggs are laid singly in the flowers or on the stems, inserted into a feeding hole and then covered with black faecal material forming a patch about 1 mm across. Larvae feed initially in the flower or leaf stalk, then migrate to enter the lower stem through an axillary bud and tunnel within the stem. In
Fig. 14.21. (a) Parthenium stem split to show stemboring larva (arrowed), and (b) adult weevil (body is 7 mm long). (QNRM)
very dry conditions, the larvae migrate into the crown and root, feeding just under the bark. The plant may respond by swelling into a partial gall. Mature larvae leave the plant and pupate in the soil, inside an earth cell. Adult emergence is triggered by rainfall. The whole life cycle takes about 2 months but may be prolonged by months of diapause induced by dry conditions. Active period: Adults are active so long as green plants are found; for example, after rainfall. Damage: Larval-feeding is very damaging, especially when the roots are attacked, and affected plants usually die or are severely stunted and produce few or no seeds. Integrated control: Soil cultivation is damaging to diapausing adults and pupae. Herbicides do not affect larger larvae.
Parthenium seed-feeding weevil Smicronyx lutulentus Dietz Coleoptera: Curculionidae Distribution: Native to Mexico, the weevil was introduced to Qld in 1980. Widespread in central Qld. 385
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affected flower head normally do not develop. Consequently, high populations destroy many of the seeds, and the reduction in the seed bank is significant. Integrated control: Deep ploughing destroys pupae and adults in the soil. Herbicides only affect very small larvae.
Parthenium stem-galling moth Epiblema strenuana Walker Lepidoptera: Tortricidae Fig. 14.22. Parthenium seed-feeding weevil larva (body is 4 mm long) and eaten-out seed. (QNRM)
Beneficial status: Where present, significantly reduces seed production. Identification: Adults are small (3–5 mm) grey weevils with a long rostrum. Larvae are white, 3 mm long and fat (Fig. 14.22). They are found inside single seeds (achenes). May be confused with: Unlikely to be confused with any other insect on parthenium. Host range: Restricted to P. hysterophorus. Life cycle: One or two generations per year, depending on rainfall. Adults are long-lived and pass the dry season in diapause in the soil, emerging after soaking rains to feed on young leaves, making small round holes. When flowering starts, the small translucent eggs are inserted singly into very young flower buds. Larvae feed within a single achene, which swells to accommodate the full-size larva, usually resulting in abortion of the other seeds in the flower. When the mature flower falls to the ground, the larvae enter the soil and form individual cells at 5–7 cm depth, in which pupation occurs. Adult emergence is delayed until after soaking rains.
Distribution: Native to Mexico, the moth was first introduced into Qld in 1982 and is now widespread. Beneficial status: Major impact in most seasons. Identification: The adult moths are nocturnal, small, 8–12 mm long, and dark in colour. Larvae are white–yellow, up to 12 mm long and found within spindle-shaped galls 10–20 mm long, 5–10 mm diameter, in stems or axillary buds. May be confused with: Adults are often found resting on the plants but cannot easily be distinguished from other similar moths. Larvae may be confused with weevil larvae but are straighter and the gall shape is distinctive. Host range: Restricted to the closely related genera Parthenium, Ambrosia and Xanthium. Life cycle: There are up to seven generations per year, and all stages may be found at any season. Adults live up to 3 weeks, and the colourless eggs (1 mm long) are laid singly on the leaf under-side or stems. Newly hatched larvae initially mine in the leaf, then migrate to a stem tip or axillary bud and commence feeding within the meristem. Feeding induces gall development and the larva feeds in a tunnel within the developing gall. Up to seven larvae
Active period: There are two peaks of adult emergence, in November and in March; however, adults are active throughout the growing season so long as flowering plants are present, and larvae are present in flowers throughout. Damage: Larval-feeding on a seed within an achene (Fig. 14.22) seems to act as a metabolic sink, as the other four achenes present in an
Fig. 14.23. Adult parthenium stemgalling moth (body is 10 mm long). (QNRM: M.W. Lee)
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may be found in a single gall but each occupies a separate tunnel. Mature larvae cut an emergence ‘window’ leaving only the epidermis, and pupate below this. Adult emergence is triggered by high temperatures and occurs around noon or shortly before. Shortening day lengths coupled with temperatures below 10°C or with poor food quality will induce diapause in pre-pupal larvae. End of diapause, followed by pupation and adult emergence, is stimulated by temperatures above about 20°C. Active period: Adults are present all year from the end of August till about April. Larvae are found from October till May and diapausing larvae in dry stems from May till August. In the north, diapause may not occur and development continues all year. Damage: The galls are metabolic sinks, diverting plant products from growing tissues. Galls thus reduce stem growth and seed production. Seedling plants may be killed, and heavy attack prior to flowering severely restricts seed production. Affected plants grow slowly and are not competitive in pasture. Integrated control: Destruction of overwintering plants (by fire or mechanical damage) will destroy diapausing pupae and severely reduce the spring population. The use of herbicides does not harm larvae.
Fig. 14.24. Larval mines of parthenium leaf-mining moth. (QNRM)
Host range: Restricted to P. hysterophorus. Life cycle: Adults are nocturnal and live up to 10 days. The minute (< 0.5 mm) eggs are laid singly on the leaf under-side and the newly hatched larvae immediately begin to mine within the leaf. Larger larvae feed on the leaf causing characteristic small holes (Fig. 14.24). Moulting between instars occurs in small temporary silk cocoons constructed against leaf veins. Mature larvae descend the plant to construct a solid silk pupal cocoon. The whole life cycle takes 5–7 weeks in summer. Despite their small size, the adults are strong fliers and disperse very widely. Active period: Adults and larvae can be found throughout the year if green tissue is present, but are most abundant in late summer. The dry season is passed in diapause in the pupal or prepupal stage.
Parthenium leaf-mining moth
Damage: Larval-feeding damages leaves and, if very heavy, will reduce seed production.
Bucculatrix parthenicola Bradley Lepidoptera: Bucculatricidae
Integrated control: Not required.
Distribution: Native to Mexico, this moth was first introduced to Qld in 1984 and is now established throughout the parthenium area.
Prickly acacia
Beneficial status: Minor. Identification: Adults are small (4–6 mm long) and silver–grey in colour. Larval mines are narrow and tortuous. Larger larvae are greenbanded with grey and found on the leaf underside. Pupal cocoons are beige to silver–grey, 3–5 mm long and longitudinally ribbed, attached to stems or leaf under-sides. May be confused with: Unlikely to be confused with any other insect found on parthenium.
Acacia nilotica Mimosoidea A native of Africa and west Asia, this species was introduced to Australia as an ornamental and to feed stock. It is a woody weed of summer rainfall pastures in north and central Qld and NT, and has spread to NSW, SA and WA.
Prickly acacia seed beetle Bruchidius sahlbergi Schilsky Coleoptera: Chrysomelidae 387
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Distribution: Cultures of this beetle were imported from Pakistan and released in Qld in 1982. Beneficial status: Minor, well established in Qld. Identification: Not easily distinguished from other seed-feeding beetles. Life cycle: The adult lays eggs in the crevice of partly dehisced (green) pods. On hatching, the (first-stage) larvae work their way up inside the pod to an unattacked seed and force entry, using the pod wall for leverage (Parsons and Cuthbertson 1992). Larvae develop by eating the seeds in the pod. Host range: In Australia, the only host is prickly acacia. Damage: Larval-feeding reduces the viability of mature seeds. In ungrazed areas, the beetle is recorded as destroying up to 65% of mature seeds but it is of limited value in stocked rangelands because stock eat the green seed pods and pass viable seeds in their dung, aiding the distribution of the plant (Radford et al. 2001). A naturalised seed-eating beetle, Caryedon serratus (Olivier) Coleoptera: Chrysomelidae, is reported as a minor destructor of prickly acacia seed. The tip-feeding moth Cuphodes profluens Meyerick Lepidoptera: Gracillariidae, imported and released at the same time as the seed beetle, has not established.
Prickly pears Opuntia spp. Cactaceae Opuntia aurantiaca, tiger pear, is very common in NSW and southern Qld. O. imbricata, devil’s rope, is common in NSW, Qld, and southern WA and occurs in all other states. O. monacantha, drooping tree pear, is common in parts of Qld and SA and occurs in all other states. O. robusta, wheel cactus, is restricted to parts of NSW, SA and Vic. O. streptacantha, white-spined prickly pear, and O. stricta, common prickly pear, are very common in NSW, Qld and southern WA and occur in all other states. O. tomentosa, velvety tree pear, is very common in NSW and southern Qld. Numerous other Opuntia species are locally naturalised (Hosking et al. 1988). Prickly pears are natives of southern USA, Central and South America. All species were
deliberately introduced to Australia, usually as ornamentals but some for human or stock food. Some species have escaped into pastures and rangelands, where their successful colonisation has in the past rendered large tracts unsuitable for grazing. Many species of prickly pear are now under biological control by one or more introduced insects and now only exist in small isolated patches (Hosking et al. 1988).
Cochineal scales Dactylopius spp. Hemiptera: Dactylopiidae Devil’s rope pear cochineal, D. tomentosus (Lamarck) Monacantha cochineal, D. ceylonicus (Green) Prickly pear cochineals, D. opuntiae (Cockerell) and D. confusus (Cockerell) Tiger pear cochineal, D. austrinus DeLotto Distribution: Cochineal scales are native to South America, Central America and southern USA. Several species were introduced to Australia in the 1920s and 1930s for biological control of Opuntia spp. Cochineal insects have been introduced to southern Africa for biological control of Opuntia spp. and to the Canary Islands as a source of dye. Beneficial status: Major: D. austrinus on tiger pear, D. tomentosus on devil’s rope, D. opuntiae on white-spined prickly pear and on common prickly pear in southern Australia and D. ceylonicus on drooping tree pear. Minor: D. opuntiae on wheel cactus and velvety pear tree, D. confusus on common prickly pear and drooping tree pear. Identification: Cochineal insects appear on cactus hosts as irregular white clusters of wax up to 10 mm long (Fig. 14.25). The wax is secreted as a protective covering by the insect below. Mature females often shelter eggs and crawlers beneath the covering. When squashed between the fingers the resulting fluid is a brilliant scarlet colour, characteristic of Dactylopius spp. and the basis of the cochineal dye industry. May be confused with: The species of cochineal insects cannot be distinguished in the field, and specific identification must be undertaken by
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a
b
Fig. 14.26. Winged male of Dactylopius opuntiae (3 mm long). (SARDI: P.T. Bailey)
Fig. 14.25. (a) Dactylopius opuntiae on Opuntia stricta, and (b) O. tomentosa. (QNRM: J. Mann)
specialists. The host plant is the best indicator of the likely identity of the cochineal species. Host range: Cochineal insects are specific to Opuntia cacti, and most individual species are restricted to a few cactus species. Life cycle: The life cycle of the tiger pear cochineal, which is typical of the other species, consists of three growth stages for females and five for males. The mature fertilised female produces eggs that hatch to crawlers within a few hours. These waxless crawlers, slightly less than 1 mm long, live for an average of 21 days. Female crawlers move to other parts of the plant or are blown by wind to other plants. They insert their mouth parts into the plant, start secreting wax and generally remain fixed there for the remainder of their development. The second stage, 2–3 mm long, is lightly covered in wax and has a mean duration of 13 days. The third-stage female is 4–5 mm long and completely covered in wax. The female may survive if unfertilised up to 80 days, and after fertilisation takes a further 50 days to develop eggs (Moran and Cobby 1979). Males are small, winged, pink–red in colour and inconspicuous. Male crawlers remain under the maternal cover and begin to feed and develop into a second stage. After some days they move
from the female and produce a waxy cocoon in which the third and fourth stages develop. The 3 mm long adult (fifth stage) has two wings (Fig. 14.26) and two long filaments protruding from the back of its abdomen. The non-feeding male flies to a female, burrows through the waxy coating and, after mating, dies. Active period: With overlapping generations, cochineal insects are present on their hosts year round. Damage: Large numbers of cochineal insects can cause severe damage, especially if the plant is under drought stress. Desiccated patches form and whole plants can be killed. Management: Where the distribution of hosts is patchy, manual distribution is used to speed up control. Choice of the correct cochineal is very important; this is ensured by redistributing colonies collected on the appropriate host cactus. Integrated control: The relative importance of C. cactorum and cochineal insects varies with cactus species and location. In cooler areas, cochineal insects may be of greater importance. Chemical control is used against velvety tree pear and other tree pears where biological control is ineffective.
Cactoblastis Cactoblastis cactorum (Berg) Lepidoptera: Pyralidae Distribution: Native to southern South America, it was imported to Australia twice, with the 1924 introduction being successful. It established in 389
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pear and white-spined prickly pear, but is not as damaging.
Fig. 14.27. Egg stick of Cactoblastis cactorum (7 mm long) attached to a cladode of common prickly pear. (QNRM)
Qld, from where it has spread to NSW, SA and Vic., and was subsequently introduced to southern Africa and elsewhere in the world. Beneficial status: Major on common prickly pear in Qld and NSW; minor on white-spined prickly pear, velvety tree pear and tiger pear. Identification: Eggs are characteristic: they are laid as sticks fastened to the cactus cladode which appear superficially to be spines of the plant (Fig. 14.27). Larvae are orange-coloured with four black spots that coalesce to black bands in older larvae (Fig. 14.28). Adults are medium-sized moths 2.5 cm long with grey– brown forewings and white hindwings. May be confused with: Unlikely to be confused with any insects on prickly pears. Host range: Common prickly pear. It will also complete its life cycle on tiger pear, velvety tree
Life cycle: C. cactorum has two generations per year in Qld and northern NSW: a short summer and long winter generation with spring (September–October) and autumn emergences. In southern Australia, the species has only one generation. Adult moths mate in early morning and females begin to lay eggs after 18–28 days in summer, 30–32 days in winter. Moths do not feed and live for up to 18 days, during which time the female lays up to three to four egg sticks each of up to 150 eggs. Adults do not disperse far if the cactus plants are dense, but can move nearly 10 km if host plants are patchy. There are six larval stages, with total development about 50 days in summer and 180 days in winter. When fully grown, larvae spin a cocoon on the host plant or nearby vegetation and pupate. Adults emerge 35–42 days after cocoon formation (Dodd 1940). Active period: Overlapping generations result in larval feeding throughout the year in northern areas. Damage: Larvae, which are gregarious feeders, eat out a cladode and then progress to nearby cladodes of common prickly pear. On whitespined prickly pear and velvety tree pear, larvae destroy small plants but do not develop in the woody cladodes of old plants. On tiger pear, larvae destroy cladodes but do not penetrate underground bulbs, which quickly regenerate.
Cactus moth Tucumania tapiacola Dyar (Lepidoptera: Pyralidae) was introduced from Argentina in 1935 and is widely established in Qld. The adult moth is 15–20 mm long, brown in colour and nocturnal. Eggs are laid singly on the plants and larvae feed singly inside the cactus cladodes. Mature larvae leave the plant to pupate in a silk cocoon in leaf litter on the soil. Identification: The solitary purplish-red larvae are unmistakable. Host range: The cactus moth attacks tiger pear, devil’s rope and harrisia cactus.
Fig. 14.28. A group of fifth-stage larvae of Cactoblastis cactorum feeding on a cladode of common prickly pear. Each larva is about 20 mm long. (QNRM: N.Riding)
Damage: Damage is usually slight and, though widespread, the moth is never abundant.
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Rubber vine
a
b
Cryptostegia grandiflora Asclepiadaceae Originally from Madascar, rubber vine was introduced as an ornamental and planted as a prospective source of rubber. It is now a widespread weed in pastures and woodlands of central and north Qld. Successful control has largely been achieved through the leaf rust Maravalia cryptostegiae introduced in 1995.
Leaf-feeding moth Euclasta whalleyi P & G. Lepidoptera: Pyralidae This moth was introduced from Madagascar in 1988 and is now widespread and locally abundant and damaging. The adult is 20 mm long, light buff in colour, and rests in a characteristic posture with wings furled and body tilted up on the long legs. Eggs are laid in batches on the leaves (Fig. 14.29) and small larvae feed in groups. Larvae are brown and very effectively camouflaged where they rest along the leaf midrib, petiole or stem (Fig. 14.29). Mature larvae pupate in silk cocoons in leaf litter (Mo et al. 2000)
Fig. 14.29. (a) White egg mass on leaf and two young larvae on the midrib of a rubber vine leaf and (b) various-sized larvae of the leaf-feeding moth. (QNRM)
Active period: Summer, with largest populations at the end of summer and early winter. Adult moths are attracted to lights and may become locally abundant. Damage: Feeding by large populations of larvae may completely defoliate large plants.
Sources of information Briano, J.A., Cordo, H.A. and Deloach, C. J. (2002). Biology and field observations of Penthobruchus germaini (Coleoptera: Bruchidae), a biological control agent for Parkinsonia aculeata (Caesalpiniaceae). Biological Control 24: 292–299. Day, M.D, Wiley, C.J., Playford, J. and Zalucki, M.P. (2003). Lantana: current management status and future prospects. ACIAR, Canberra. Dhileepan, K. (2001). Effectiveness of introduced biocontrol insects on the weed Parthenium hysterophorus (Asteraceae) in Australia. Bulletin of Entomological Research 91: 167–176. Dodd, A.P. (1940). The biological campaign against prickly pear. Commonwealth Prickly Pear Board, Brisbane. Donnelly, G.P. (2000). Biology and host specificity of Rhinacloa callicrates Herring (Hemiptera: Miridae) and its introduction and establishment as a biological control agent of Parkinsonia aculeata L. (Caesalpiniaceae) in Australia. Australian Journal of Entomology. 39: 89–94. Hosking, J., McFadyen, R.E. and Murray, N.D. (1988). Distribution and biological control of cactus species in eastern Australia. Plant Protection Quarterly 3: 115–123. Klinken, R.D. van, Fichera, G. and Cordo, H. (2003). Targeting biological control across diverse landscapes: the release, establishment, and early success of two insects on mesquite (Prosopis spp.) in Australian rangelands. Biological Control 26: 8–20. McFadyen, R.E. (1979). The cactus mealybug, Hypogeococcus festerianus (Hemiptera: Pseudococcidae) for the biological control of Eriocereus martinii in Australia. Entomophaga 24: 281–287.
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McFadyen, R.E. (1984). Annual ragweed in Queensland. Proceedings of the 7th Australian Weeds Conference, Perth, pp. 205–209. McFadyen, R.E. (1989). Ragweed, parthenium and noogoora burr control in the post-Epiblema period. Proceedings of the 5th Biennial Noxious Plants Conference, NSW. McFadyen, R.E. (1992). Biological control against parthenium weed in Australia. Crop Protection 11: 400–407. Mo, J., Treviño, M. and Palmer, W.A. (2000). Establishment and distribution of the rubbervine moth Euclasta whalleyi P-G & C (Lepidoptera: Pyralidae) following its release in Australia. Australian Journal of Entomology 39: 344–350. Moran, V.C. and Cobby, B.S. (1979). On the life-history and fecundity of the cochineal insect, Dactylopius austrinus de Lotto (Hemiptera: Dactylopidae), a biological control agent for the cactus Opuntia aurantiaca. Bulletin of Entomological Research 69: 629–636. Parsons, W.T. and Cuthbertson, E.G. (1992). Noxious Weeds of Australia. Inkata Press, Melbourne. Radford, I.J., Nicholas, D.M. and Brown, J.R. (2001). Assessment of the biological control impact of seed predators on the invasive shrub Acacia nilotica (prickly acacia) in Australia. Biological Control 20: 261–268. Walton, C. (2005). Reclaiming Lost Provinces. A century of weed biological control in Queensland. Department of Natural Resources and Mines, Brisbane. Willson, B.W. and Garcia, C.A. (1992). Host specificity and biology of Heteropsylla spinulosa (Hom.: Psyllidae) introduced into Australia and western Samoa for the biological control of Mimosa invisa. Entomophaga 37: 293–299.
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15 PASTURES (INCLUDING LUCERNE)— WINTER RAINFALL GRASS PASTURES AND TURF C. Pavri and C.J. Young
Grass dominant irrigated pasture
GROUP Grass pastures and turf mites
springtails crickets, mole crickets and grasshoppers beetles
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PEST (major pests in bold) cereal rust mite blue oat mites bryobia pasture mite balaustium mite lucerne flea black field cricket mole crickets wingless grasshopper (see Chapter 18) Argentine stem weevil white fringed weevil African black beetle blackheaded pasture cockchafers redheaded pasture cockchafer other pasture scarabs
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GROUP
PEST (major pests in bold)
caterpillars
black cutworm corbies, winter corbies, underground grassgrubs ghost moths oxycanus grassgrub pasture tunnel moth (or philobota) pasture webworms, Hednota spp. weed web moth (see cotton webspinner) armyworms
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Legume pastures Lucerne
Winter rainfall pastures are typically composed of grasses (annual and/or perennial), legumes (annual and/or perennial) and broadleafed plants (often regarded as weeds). In this chapter, grass pastures and legume pastures and lucerne are treated in separate sections; however, grass and lucerne or other legume forage plants are often grown together to maximise grazing potential. Pastures may be irrigated and fertilised to increase quality and yield, and thus animal productivity. Pastures may also be grown for seed and to make hay. The relative composition of the pasture is the result of management practices. The activities of invertebrates can greatly hinder pasture establishment and growth, alter the pasture species mix in deleterious ways, render the pasture less palatable than normal, encourage further damage by vertebrate predators, lead to the invasion of weed species and expose the infested area to soil erosion. Native and introduced grasses provide food for a number of invertebrate pests. Broadleafed plants such as capeweed and brassicaceous weeds may act as reservoirs of a number of crop pests. Grass pastures and cereal crops share a number of insects in common.
Cereal rust mite Abacarus hystrix (Nelapa) Acarina: Eriophyidae Pest status: Minor, widespread and regular inhabitant of winter grass pastures. The cereal rust mite is common on perennial and annual ryegrass, in southern Australian pastures. The cereal rust mite is similar in
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appearance and life history to the wheat curl mite (Chapter 2). It is the vector of ryegrass mosaic virus (RMV), a widely distributed pathological disorder of grasses. While RMV has been shown to decrease pasture yields under some circumstances, its economic impact in Australia is likely to be insignificant. These mites may be preyed upon by predatory mites, Parasitus sp. (Parasitidae) and infected by the fungus Hirsutella sp. In some irrigated dairy pastures, heavy strip grazing at irregular intervals reduces numbers of mites but has not been demonstrated to reduce the incidence of RMV (Frost 1995).
Blue oat mites Acarina: Penthaleidae Penthaleus major (Dugès), P. falcatus (Qin and Halliday) and P. tectus Halliday Pest status on grass pastures: Minor, widespread, irregular. Identification and life cycle: See Chapter 2. Risk period: Autumn to spring, but especially at germination. Damage: Damage is visible as a silvering of the leaves. Seedling mortality and reduced seedling growth of both grasses and legumes can occur in germinating pastures in autumn, reduced production and quality of older green plants can occur during the growing season, and reduced seed yield in legumes can occur in spring. In winter pastures, the majority of damage and production loss is to grasses in the pasture.
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a
b
Fig. 15.1. Bdellodes lapidaria (length 2 mm), a generalist predator of pasture mites. (SARDI)
Monitoring: Start monitoring from seedling emergence, especially when weather conditions meet the criteria for hatching of the oversummering eggs. Newly hatched larvae and young nymphs are very small so close inspection is essential. Action level: When mite emergence coincides with germination, any sign of mite activity or damage to seedlings warrants control. Chemical control: The most effective time to control blue oat mites is within 2–3 weeks from the break of season; however, regular inspection of the pasture for evidence of mite activity is necessary as damage to emerging seedlings could occur before the spray is applied. Spring spraying, as recommended for redlegged earth mite, is not effective on blue oat mite due to their differing diapause strategies. There is also evidence of tolerance to some insecticides by P. falcatus (Robinson and Hoffmann 2001). Cultural control: Heavy grazing of pastures in winter and spring significantly reduces mite populations. Natural enemies: Anystis wallacei was introduced from France for biological control of blue oat mite, redlegged earth mite and lucerne flea (Fig. 15.41). The predatory mites of Balaustium sp. have also been observed preying on blue oat mite. Bdellodes lapidaria (Fig. 15.1), a generalist predatory mite, occurs in pastures at the same time and may feed on blue oat mite. All of these predators have limited impact on blue oat mite populations.
Fig. 15.2. (a) Eggs and first-instar nymphs of mole crickets (PIRSA: P.R. Birks) , and (b) adult mole cricket, 30 mm long (Graphic Science: © Denis Crawford) .
Mole crickets Gryllotalpa spp. Orthoptera: Grylloidea Distribution: Several species of Gryllotalpa are distributed throughout Australia. Pest status: Minor, localised particularly to sandy soils, irregular. Identification: Readily identified by their large, powerful mole-like forelegs, which are used for digging burrows (Fig. 15.2). May be confused with: Native species may be confused with the introduced Changa mole cricket but the native Gryllotalpa spp. have four digging claws on the forelegs, not two as in the Changa mole cricket. In WA, sandgropers (Chapter 2) may be distinguished from mole crickets by their elongated shape and lighter colouration. Host range: Feed on a wide range of plants. Some species are carnivorous on soil arthropods. Life cycle: Single generation per year. Mole crickets live permanently in burrows, which they extend to form temporary feeding galleries just beneath the surface of the soil. At the end of the horizontal galleries is a vertical shaft several centimetres deep where the insect rests and lays its eggs. They mostly feed on plant roots. When 395
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mature, male mole crickets will sit in the entrance to their burrow in a specially designed dome-shaped opening and sing for a mate. The shape of the burrow opening amplifies the sounds of their calls. Female mole crickets lay eggs within their own burrows, remaining close by to protect them until they hatch. The nymphs resemble small adults but lack wings. They develop through several moults, growing wing buds that eventually develop into wings with the final moult. As the soil dries in summer, the insects gradually migrate to moist patches or go deep underground. They are seldom seen on the surface in the daytime, but occasionally come out at night seeking food. Risk period: Usually most active in early autumn and spring. Damage: Rarely cause economic damage to pastures but when in high numbers, they can damage turf and lawns. Monitoring techniques: Dig areas where damage or tunnelling is occurring to look for nymphs and adults. Action level: Small mounds or raised ridges on the soil surface indicate that mole crickets are active in turf. Chemical control: Because of its underground habits, control is difficult and not usually costeffective. If treatment is warranted, baits (such as those used for grasshopper control) or lawn grub insecticides are recommended. Natural enemies: Various species of amphibians, reptiles, birds, mammals, insects and spiders prey on mole crickets.
Argentine stem weevil Listronotus bonariensis (Kuschel) Coleoptera: Curculionidae Distribution: Originated from South America, now present in Australia and New Zealand. In Australia it has established in NSW, SA, Tas., Vic. and WA. Pest status: Minor, restricted, irregular in Australia; however, it is a major pest in New Zealand. Identification: The adult weevil is a hardbodied beetle about 3–3.5 mm long (Fig. 15.3). It
Fig. 15.3. Adult Argentine stem weevil (length 3 mm). (AgResearch Lincoln: M. McNeill)
is grey–brown in colour with three pale longitudinal stripes on its thorax and short, stiff bristles covering the body. The larvae are crescent-shaped, tapering at either end with a creamy white body, small straw-coloured head and, unlike cockchafers, do not have legs. May be confused with: Larvae of sitona weevil, Sitona discoideus. Host range: Grasses, mainly Italian and endophyte-free perennial ryegrasses, annual ryegrasses, cereals and germinating canola. Life cycle in winter rainfall pastures: In Australia, there may be three to four generations per year between August and May, with autumn adults overwintering in plant litter on the soil surface. Adults emerge in spring, mate and lay eggs in the leaf sheath of grass. The larvae tunnel into the stems and later move into the soil to feed on the roots (Power 1984). Risk period: Severe damage can occur to grasses in winter rainfall pastures after the first generation larvae emerge in spring. Damage: Adult feeding damage on the grass leaf near its tip is not normally significant in winter rainfall pastures. Mining of the stem by the larvae causes the greatest damage, the vegetative tillers wilt and become yellow. Flowering tillers whiten and the head may break in the wind. With heavy infestations, turf shows patches of yellow, followed by browning and death (Fig. 15.4). Monitoring: Usually not necessary in Australian pastures. Action level: Populations found in Australian pastures do not normally warrant control.
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Fig. 15.4. Argentine stem weevil damage to turf. (SARDI: K. Henry)
Chemical control: In areas with a history of damage, a preventative spray applied in spring to kill the adults before egg-laying commences, but follow up sprays are often required. Cultural control: Light spring grazing of heavily infested pastures encourages natural re-seeding of grasses. Infested pastures that will be sown to a susceptible crop in the following year should be cultivated at least 3 weeks before sowing. Host plant resistance: Long rotation perennial ryegrasses are more resistant than short rotation varieties. Natural enemies: An entomopathogenic nematode, Heterorhabditis sp., has proved to be effective in controlling stem weevil when applied as a biocide in turf (Ford et al. 2000). The nematode burrows through the intersegmental membranes of the larva and releases bacteria that kill the larva. The use of this nematode for controlling stem weevil in broadacre agriculture is not cost-effective but may be useful for high value turf.
Fig. 15.5. African black beetle (left to right): egg; firstinstar larva (6 mm long); second-instar larva; and thirdinstar larva (30 mm long). (SARDI: K. Henry)
25–30 mm in length with a white to creamy white body and a light brown head (Fig. 15.5). The rear end has a dark grey tinge caused by faecal matter in the hind gut until pre-pupation when the gut is cleared. When resting, the grubs are curled into a C-shape (Goodyer 1983). The adult beetle is a rich chestnut colour when newly emerged, but soon changes to shiny black (Fig. 15.6). The beetles are stout-bodied and 10–15 mm in length. May be confused with: Adults of the native black beetle (Metanastes vulgivagus) appear the same as African black beetle; howeve, the native black beetle is only an irregular and minor pest. Adults of redheaded pasture cockchafer are also similar, but they fly earlier in spring. Larvae of the lesser pasture cockchafer (Australaphodius frenchi) may be confused with this species, but its larvae (3–4 mm long) are never larger than the first instar of African black beetle. The third-
African black beetle Heteronychus arator (Fabricius) Coleoptera: Scarabaeidae Distribution: Native of Africa, now present in Australia and the North Island of New Zealand. In Australia, it occurs mainly in south-western WA and from coastal south-eastern mainland up to south-east Qld. It is not recorded in Tas. Pest status: Minor, widespread, irregular. Identification: Newly hatched larvae are about 5 mm long with six legs, white bodies and pale brown heads. The fully grown larvae are
Fig. 15.6. African black beetle adult, 15 mm long. (Graphic Science: © Dennis Crawford)
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instar larvae of other native cockchafer species may also be confused with the African black beetle, but their larvae are usually more abundant in winter than African black beetle larvae. Host range: In pastures, African black beetle is commonly found attacking grasses, particularly paspalum and ryegrass, as well as carpet grass and couch. They can also feed heavily on phalaris and kikuyu without killing the grass. Ryegrass appears to suffer less damage than most other grass species, while the legume component of pastures is usually ignored. African black beetle also attacks a wide variety of horticultural crops including potatoes, maize, sweet corn, sugarcane, vegetable seedlings, olives and young grape vines, as well as transplants of tomatoes, cabbages and cauliflowers and plantation tree seedlings. Life cycle on winter rainfall pastures: African black beetle has one generation per year but in some seasons the generation can take 2 years. Eggs are laid in the soil in spring and hatch within 2–5 weeks, depending on the temperature. Larvae hatch and pass through three stages before pupating and emerging as adults between late December and early February (Fig. 15.6). The period from egg to adult is about 3–4 months, while the adult lifespan is about 10 months. The adults remain sexually immature until spring when mating and oviposition occurs. While some third-stage larvae may be found during winter, they are late developers that fail to survive (Matthiessen and Ridsdill-Smith 1991). Risk conditions: Most damage to pastures is caused by the third-stage larvae, which occur during summer from December to February. This follows annual grass pasture senescence, which is why African black beetle is not considered a serious pest in winter rainfall pastures. Perennial pastures, however, can be severely damaged. Damage: The soil-dwelling larvae cut off the grass roots below the surface of the soil. Severe infestations cause the pasture to wilt and ‘brown-off’; underfoot the soil feels loose and spongy, and the pasture can be rolled back like a carpet due to the complete severing of the roots from the plant tops (Fig. 15.7). Birds, such as magpies and crows, often break up the pasture
Fig. 15.7. African black beetle damage to grass pasture; root-pruned pasture can be rolled back. (Graphic Science: © Denis Crawford)
as they forage for larvae and can do as much, if not more, damage than the larvae themselves (Goodyer, unpublished). Adult-feeding may also cause significant damage to pasture plants. Monitoring: Inspect pasture during summer– autumn. Soil-sampling: Numbers of larvae can be determined by spade-sampling. Randomly take 10 separate square samples each the width of a spade (20 cm wide) and 15 cm deep in a line diagonally across the paddock. Break up the soil over a plastic sheet and count the number of grubs found. Adult flight activity can be monitored during summer–autumn by the use of light trap catches or observing activity around lights near buildings. Adult ‘roaming’ catches: Monitor the adults during their ‘roaming stage’, a period during spring when adults emerge from overwintering in the soil and roam on the pasture in search of a mate. The beetles are clumsy walkers and accumulate in pitfall traps or sharp-sided plough lines. Check traps fortnightly during the adult ‘roaming’ between September and October. Action level: If monitoring for larvae or adults reveals damaging populations, spraying should be considered, especially in newly sown or establishing pastures. Chemical control: Insecticide application against subterranean larvae during summer is ineffective. Adults may be killed before they lay eggs during the spring ‘roaming’ stage. But monitoring for the presence of adults and timing
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of the spray is critical: if the spray is applied too early (e.g. August), the adults may not have become sufficiently active to be contacted and will not be killed; and if the spray is too late (e.g. after mid-October), the adults may have already laid their eggs (Fisher and Learmonth, undated). Cultural control: Delay autumn-sowing until May wherever possible as the new-generation beetles will have slowed their rate of feeding with the colder weather. Pastures sown into areas with a history of beetle damage should contain a legume and a tolerant grass in addition to the susceptible grasses.
Blackheaded pasture cockchafers Coleoptera: Scarabaeidae Acrossidius (= Aphodius) tasmaniae (Hope) and A. pseudotasmaniae (Green) Distribution: Both species are native to Australia. A. tasmaniae is recorded from southern Australian pastures in rainfall areas as low as 375 mm, and also occurs along the east coast of Australia. A. pseudotasmaniae occurs in Tas. and is recorded as an introduced pest of New Zealand pastures. Pest status: A major pest of southern pastures, minor along the east coast of Australia, restricted within their range, irregular in occurrence. Identification: Newly hatched larvae are 1– 1.5 mm long and have large dark brown heads which, during the second and third larval stages, darken to almost black (Fig. 15.8).
Fig. 15.8. Third-instar larvae of blackheaded pasture cockchafer (body length 20 mm). The larva on right has laid down fat (white colour) and is close to pupating. (SARDI: P.G. Allen)
Fig. 15.9. Two species of blackheaded pasture cockchafer: (left) Acrossidius tasmaniae (12 mm) and (right) A. pseudotasmaniae, (10 mm) which has hairier wing covers. (DPI&W Tas)
Pre-pupae are 1.3–1.8 cm long with creamy bodies and brown–black heads (Fig. 15.8). The pre-pupae contract to form pupae, 1–1.2 cm long. Adult beetles are 8–12 mm long, dark brown, reddish-brown or almost black in colour (Fig. 15.9). May be confused with: Larvae are similar to other scarabs, but have a dark head colour (Fig. 15.8), and the surface-feeding habit of the larvae of both species are distinctive. Blackheaded pasture cockchafers produce soil casts at the opening of larval tunnels. This distinguishes them from tunnels of underground grass grubs, which are silk-lined, and from corbies, which have silk runways leading to tunnel entrances. Larvae of the two species of blackheaded pasture cockchafer are difficult to distinguish. Adult males of A. psuedotasmaniae have conspicuous rows of hairs on the wing covers, which are lacking in A. tasmaniae (Given 1950). Host range: Older larvae prefer legumes but will eat annual grasses, weeds and dung. Life cycle: One year. Adult beetles prefer bare patches to lay eggs and, if the summer is dry, are concentrated in damp patches in the pasture. During wet summers, egg-laying is more widespread. Eggs are laid 5–15 cm deep in the soil and hatch in 2–4 weeks. Larvae hatch during February to March and make their way to the soil surface but do not move far along the surface. Larval survival is dependent on rain after hatching, and mortality (up to 50%) increases markedly during dry autumns. With sufficient moisture, young larvae feed on surface organic matter and construct vertical tunnels to 20 mm depth. After 2–3 weeks they moult to their second stage, which is usually of 2–3 weeks duration but may be prolonged up to 3 months. 399
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very sandy or very heavy soils and lightly grazed perennial pastures. Favoured egg-laying sites include stock camps, tops of rises, trees, posts and fences. However, if the weather during late summer–autumn is cold and windy, adults usually remain near to their larval tunnels, mate on the soil surface and re-infest the same area. Mated females lay 20–40 eggs in soil 3–6 days after emergence and may lay a second batch of 10–20 eggs after a feed of dung. Adults live for about 2 weeks (Carne 1956). Risk period: From mid-autumn to mid-spring (April to late October), when third-stage larvae are feeding.
Fig. 15.10. Blackheaded pasture cockchafer larval tunnels are constructed vertically beneath a feeding area. The larva (arrowed) drags excised vegetation to the bottom of its tunnel. (DPI&W Tas)
About 2 weeks after moulting, the second-stage larvae change from feeding on dead organic matter to feeding on live plants growing around their tunnels. Second- and early third-stage larvae move from their tunnels to feed at nights following sufficient rainfall. Larvae feed for 2–3 nights when enough food is taken into their tunnels to last for 7–10 days. Once they have eaten out an area close to their tunnels, they move up to 4 m and construct new tunnels. Fully fed third-stage larvae live in tunnels 8 mm in diameter and to 20 cm depth (Fig. 15.10). The third larval stage lasts 3–5 months. Excessive rain during autumn, winter or spring increases larval mortality by a combination of drowning, larval competition and fungal disease. In late winter or early spring, larvae descend to the bottom of their tunnels and enter a nonfeeding pre-pupal stage of 1–4 months’ duration. Pre-pupae construct cells at the bottom of their tunnels and develop into a pupal stage of up to 1 month duration. The adults are present in December and wait in their tunnels until rain during January–March softens the ground and they ascend to the surface. Adults emerge at dusk and may disperse up to 3 km if the weather is warm and humid, with frequent thunderstorms. Adults swarm around dung, bare patches and are attracted to lights. Females prefer to lay eggs in sandy loam, loam or light clay soils and avoid
Damage: Heavily infested annual pastures may be thinned or eaten out (Fig. 15.11), reducing the quantity of feed during winter and spring followed by weed invasion. Early damage is localised but spreads outwards as the season progresses. Lawns, parks and golf courses may be damaged (Goodyer 1984). Monitoring: Check areas that have been infested in previous years or annual pastures on which organic matter has accumulated for 3–4 years. Use spade samples to separate and count larvae. Action Level: For pastures, a population density of young larvae greater than about 300 per square metre may result in severe damage. Chemical control: Is possible, as larvae are surface-feeders. Treatment of affected areas should begin when larvae begin active feeding in April–May, or where parks are watered, control
Fig. 15.11. Blackheaded pasture cockchafer damage to grassy pasture. (SARDI: P.G. Allen)
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may commence earlier. Apply when larvae are actively feeding, usually following rain. Cultural control: Planting perennial pastures and avoidance of overgrazing during summer markedly reduces damage. Natural enemies: Naturally occurring diseases including Cordyceps aphodii (Fig. 15.22) (Coles 1980), have been recorded in natural populations of blackheaded pasture cockchafers but their controlling impact has not been recorded. A species of flower wasp (Hymenoptera: Tiphiidae) has been recorded parasitising up to 50% of late larval stages of blackheaded pasture cockchafers (Fig. 15.20). Adults of the wasp feed on nectar from some trees and shrubs. The rate of parasitism is related to availability and distance from these plants (Carne 1956).
Redheaded pasture cockchafer Adoryphorus couloni (Burmeister) Coleoptera: Scarabaeidae Distribution: Native to Australia. Occurs in NSW, Qld, south eastern SA, western districts of Vic., Tas., and the South Island of New Zealand. Favours sandy loam soils. Pest status: Major, restricted, irregular. Identification: The larvae have six legs, a reddish-brown head and whitish-grey to cream body (Fig. 15.12). The larvae curl into a C-shape when disturbed. The adult is a reddish-brown to black beetle, shiny in appearance and 10–15 mm in length (Fig. 15.13). May be confused with: Many other scarab larvae, including pasture scarab (Sericesthis nigrolineata), small pasture scarab (Sericesthis nigra), pasture beetle (Scitala sericans) and
Fig. 15.12. Comparison of sizes: fully grown larvae of the blackheaded pasture cockchafer (left), shiny pasture scarab (centre), and the redheaded pasture cockchafer (length 25 mm) (right). (SARDI: K. Henry)
Fig. 15.13. Redheaded pasture cockchafer adult (length 15 mm). ‘Redheaded’ refers to the larval head colour. (DPI&W Tas: C. Young)
African black beetle (Heteronychus arator). Fully grown larvae of the redheaded cockchafer are considerably larger than Sericesthis and Scitala larvae but smaller than African black beetle larvae, which can also be distinguished from these native species by their yellow to pale brown heads. Larvae of the hairy scarab (Saulostomus villosus) are difficult to distinguish from redheaded cockchafer. The raster may be used as a guide to separate some common species of scarab larvae (Fig. 15.14). The pattern of hairs (raster) on the ventral surface of the last abdominal segment of scarab larvae may be used in separating common pest species but is not diagnostic in separating pest species from non-pests. Host range: Annual and perennial grasses, subterranean clovers. The adults do not feed. Life cycle on winter rainfall pastures: The redheaded pasture cockchafer has a 2-year life cycle. The adult beetle emerges from the soil between August and early October, flies to trees and then lays eggs singly in the soil under the pasture. The eggs hatch after about 2–4 weeks and the larvae moult twice before becoming nearly fully grown by early autumn the following year. These large final-stage larvae cause the most damage as they feed through autumn and winter. In late spring–early summer the larvae become a creamy colour, move downwards into the soil and then pupate in December–January. The adults emerge from the pupae at about the end of January and remain in the soil until early the next spring when they come to the surface, fly for a brief period, then return to the pasture where they burrow into the 401
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redheaded pasture cockchafer, Adoryphorus coulonii
African black beetle, Heteronychus arator
hairy pasture scarab, Saulostomus villosus
pasture beetle, Scitala sericans
pasture scarab, Sericesthis nigrolineata
Sericesthes micans
Fig. 15.14. Rasters of some winter rainfall pest scarab larvae. Rasters are a useful aid in separating scarab species but by themselves may not be diagnostic. (NSWDPI: A. Westcott)
soil, mate and lay eggs for the beginning of the next generation (Heap 1998). Risk period: Due to the 2-year life cycle, damage is likely every other year. In winter rainfall pastures, the majority of damage will be from germination through to late autumn–early winter when the large larvae are active and before they become semi-dormant over winter.
Action level: Four larvae per spade square is equivalent to 100 per square metre. If large numbers (> 200 per square metre) are present, management adjustments may be required in the affected paddock. Chemical control: Surface-applied insecticides are not effective as they do not penetrate beyond the surface layer of the soil.
Damage: Damage from larval feeding mainly occurs in pastures containing annual and perennial grasses and subterranean clover that are 3 or more years old (Fig. 15.15). Heavy infestations can sever roots so that the pasture can be rolled back like a carpet, as is done by foraging forest ravens. Monitoring: Inspect pastures older than 3 years in autumn–early winter. Numbers of larvae can be determined by spade-sampling. Randomly take 10 separate square samples each the width and depth of a spade (20 cm wide) (Fig. 15.16) in a line diagonally across the paddock. Break the soil over a plastic sheet and count the number of larvae.
Fig. 15.15. Bare patches caused by feeding of redheaded pasture cockchafer on roots of perennial grasses. (SARDI: P.G. Allen)
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irregular occurrence; however, in relatively small patches and in some years they may cause noticeable pasture damage. A number of these scarabs have 1-year life cycles but some species have life cycles that may extend to 2 or more years. Adult beetles fly for short periods during summer (November–February). Many species feed on the foliage of eucalypts, which they sometimes defoliate. Adults of the pruinose scarab, for example, feed on leaves of a variety of Eucalyptus species, particularly candlebark, Blakely’s redgum and manna gum. Christmas beetles (Anoplognathus spp.) and spring beetles (Diphucephala spp.) feed on eucalyptus foliage. Fig. 15.16. The density of redheaded pasture cockchafer larvae in this spade sample is equivalent to about 750 per square metre. (SARDI: P.G. Allen)
Cultural control: Reducing pasture residue in paddocks reduces larval densities, also removing excess pasture residue by grazing pastures heavily during spring and summer, heavily grazing historically susceptible pastures during the growing season, especially in years of expected damage, grazing pastures through the summer to remove cover prior to autumn and cutting pastures for hay in spring. Host plant resistance: Perennial grasses are more tolerant to damage than annual pasture plants. Lucerne appears not to be attacked by redheaded cockchafer.
Females lay eggs in the soil, usually under pasture. Larvae have six legs with a head colour of yellow, orange or various shades of brown, depending on species, with a soft, greyish-white to creamish-grey body, often called curl grubs. Larvae live in the soil without coming to the surface, feeding on roots of native and introduced grasses and legume species. They pass through three instars, of which the thirdinstar larvae are the most voracious and hence the most damaging. When they are fully grown, third-instar larvae cease eating for a period (the ‘pre-pupal’ stage) and then develop into pupae, which can often be found in pupal cells when dug from the ground. Variations in the life cycles between species is mainly due to differences in duration of the third larval and pupal stages.
Natural enemies: A strain of Metarhizium anisopliae, an indigenous fungus, has been formulated into granules and is available commercially. Good control has been recorded when the granules are sown with the seed of new pastures. General predators such as birds have been observed eating larvae.
The main factor influencing outbreaks of pasture scarabs is weather. Dry springs kill many newly emerged beetles and suppresses egg-laying by survivors. Hot, dry weather during summer and autumn kills many eggs and newly hatched larvae by desiccation and heat stress. On the other hand, an unusually wet autumn and winter may result in drowning or disease (See Natural enemies, below) (Goodyer 1985).
Pasture scarabs that are minor pests, with restricted distribution and irregular in occurrence
Minor pest scarabs include the next six species described below.
Several scarab beetles whose native habitat is Australian grasslands have adapted to eat introduced pasture species. Land clearing and improved pastures have made these scarabs more common. They are minor pests in most pastures, with a patchy distribution and
Dusky pasture scarab Sericesthis nigrolineata Boisduval The adult of this species is 11–13 mm long and dark reddish-brown in colour. It is found in 403
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coastal pastures and tablelands of ACT, NSW and Qld, Murray–Darling basin, SA, Tas and Vic. It favours friable, well-drained soils. Eggs hatch after about 1 month and the larvae pass through three stages. Total larval development can take from 10–20 months. Some individuals complete their life cycle in 1 year, but a proportion of the population takes 2 years. Damage mainly occurs in pastures that are 3 or more years old and that include annual and perennial grasses and subterranean clover (Davidson et al. 1972).
widespread in southern and eastern Australia in NSW, SA, Tas. and Vic. It has a life cycle of 1 year. Oviposition is subterranean and occurs mainly in December. The larvae pass through three instars between January and September– early October before pupating. The pre-pupal– pupal stage lasts about 5–6 weeks and is present in the soil between late September and November. The adult develops from the pupa and remains in the pupal cell for several weeks before emerging (Hardy 1976b).
Hairy scarab
Small pasture scarab
Saulostomus villosus Waterhouse
Sericesthis nigra (Lea)
The adult beetle is 12–13 mm long with a dark brown pronotum and yellow–brown to dark brown elytra, 10 mm in length. It is found in NSW, SA, Tas. and Vic. on sandy to clay loam soil in regions of warm-temperate and humid or Mediterranean climates with average annual rainfall between 600–1200 mm. The hairy scarab takes 2 years to complete its life cycle (Hardy 1976a).
This species is found in the south-east coastal pastures of NSW, Tas. and Vic.
Pasture scarab Sericesthis micans Blackburn This species is found in the Murray–Darling basin of NSW, SA, and the southern gulfs of SA and in Vic.
Pruinose scarab Sericesthis geminata Boisduval The adult is 11–16 mm long and has a head and pronotum that are reddish-brown with a purplish-green iridescence, particularly on the pronotum. The elytra are a brownish-yellow. It is found from Townsville, along coastal NSW, across southern Vic. and the south-east of SA. It has a 1-year life cycle (Carne and Chinnick 1957).
In addition, Goodyer (1977) lists the blacksoil scarab, Othnonius batesii Oliff (2-year life cycle)and rhopaea canegrub, Rhopaea magnicornis Blackburn (2-year life cycle) as pasture pests in northern NSW. McQuillan (1985a) lists a number of additional species of scarabs as occurring in Tasmanian native and introduced pastures but does not indicate their pest status. These include Pimelopus nothus Burmeister, Cheiroplatys latipes (GuérinMéneville), Anoplognathus suturalis Boisduval, Phyllotocus macleayi Fisher, P. bimaculatus Erichson, P. nigripennis lea, P. rufipennis (Boisduval), Colpochila obesa Boisduval, Liparetrus sp., Telura sp., T. vitticollis Erichson, Automolius depressus (Blanchard), Diphucephala colaspidoides (Gyllenhall), and Heteronyx tasmanicus Blackburn. Risk conditions: Pastures are at risk during late autumn and early winter when third-stage larvae are feeding actively. Germinating pastures, or established pastures under conditions of drought or overgrazing, are likely to show signs of scarab damage. For scarab species with a 2-year life cycle, there will be a risk of damage every second year.
Scitala sericans Erichson
Damage: Dense populations of any of these pasture scarabs cause the pasture to wilt and ‘brown-off’. Underfoot the soil feels loose and spongy, and the pasture can be rolled back like a carpet due to the root system being severed from the plant top.
The adult beetle is 10–14 mm long, semi-shiny and dark brown in colour. This species is
Monitoring and action level: As for redheaded pasture cockchafer.
Shiny pasture scarab (pasture beetle)
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a
Fig. 15.17. Scarab predators: carabid beetle larva in side view (25 mm long), showing predatory mouth parts and legs. (NSWDPI: A. Westcott) b
Fig. 15.18. Scarab predators: legless robber fly larva (Diptera: Asilidae) in top view (40 mm long). Asilid larvae live in the soil and are capable of seeking and killing scarab larvae. (NSWDPI: A. Westcott)
Chemical control: Chemical control of scarab larvae in pastures is not cost-effective.
Fig. 15.20. (a) Third-instar scarab lava being consumed by a larva of a hairy flower wasp (NSWDPI: C. Haywood) , and (b) the adult of which feeds on flowers on nearby shrubs (K. Power) .
Cultural control: Reducing pasture residue in paddocks during summer exposes larvae to desiccation and predation. Natural enemies: Scarab larvae are preyed upon by a variety of birds, the most important being the straw necked ibis, the Australian magpie, mammals (especially bandicoots) and insects. These include carabid beetle (Coleoptera: Carabidae) larvae (Fig. 15.17), robber fly larvae (Diptera: Asilidae) (Figs 15.18 and 15.19), and click beetle (Elateridae) larvae (Fig. 15.19). Scarab larvae are parasitised by wasps, including hairy flower wasps (Hymenoptera: Scoliidae) (Fig. 15.20), flower wasps (Hymenoptera: a
Fig. 15.21. The scarab larva on the left has milky disease, evident in the white abdomen and the white drop of haemolymph (insect blood) on the third leg, which has been cut. The larva on the right is healthy, with a drop of translucent haemolymph on a cut third leg. (CSIRO Entomology: R. Milner)
Tiphiidae) and parasitic flies (Diptera: Tachinidae). Land clearing has removed trees and shrubs which are used as shelter and nectar sources by these natural enemies (Goodyer 1985). Larvae may also be infected by a number of diseases, including milky disease, caused by the bacterium Paenibacillus popilliae (Fig. 15.21) (Milner 1981) and the fungus Cordyceps (Fig. 15.22) (Coles 1980), and viruses (Fig. 15.23).
b
Fig. 15.19. Evidence of predation: (a) scarab larva eaten by asilid (Diptera: Asilidae) larvae, and (b) scarab larvae killed by elaterid larvae (Coleoptera: Elateridae). (NSWDPI: L. Turton)
Fig. 15.22. Healthy blackheaded pasture cockchafer larva (left) and a larva killed by the fungal pathogen Cordyceps aphodii (right), recognisable by outgrowths from the head. (R. Coles) 405
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Fig. 15.24. A mature corbie larva (50 mm long). (DPI&W Tas: C. Young)
Fig. 15.23. Virus-infected larva (left) compared with a healthy larva (right). (DPI&W Tas: L. Hill)
Black cutworm
years, depending on species. Their numbers appear to have increased after land clearing for pastures, with the introduction of exotic pasture species for ‘improved’ pastures, with the use of fertilisers and, in some cases, irrigation.
Agrotis ipsilon (Hufnagel) Lepidoptera: Noctuidae
Corbie, Oncopera intricata Walker
N (Main entry in Chapter 2 Cereals.)
Pest status: Corbie: major in Tas., widespread in pastures, irregular. Winter corbie: major in southern NSW and wetter parts of Tas., widespread in pastures, irregular.
Pest status (turf only): Major, a pest of turf in northern NSW and Qld, regular. Identification: See Chapter 2. Adult moths lay eggs singly on the tips of grass blades. When the eggs hatch, larvae move to the base of grasses. Larvae damage creeping bentgrass, tall fescue and perennial ryegrass. Regular mowing removes most eggs but clippings need be removed from the area. Planting resistant grasses such as Kentucky bluegrass helps avoid damage. Insecticide application on grassy areas surrounding managed turf may be cost-effective (Potter and Williamson 1997).
Winter corbie, Oncopera rufobrunnea Tindale
Identification: Eggs are less than 1 mm, ovoid, white when first laid and turn black after 24 hours. Newly hatched larvae are about 3 mm long and reach 6 cm in length by late spring (Fig. 15.24). These bluish-grey larvae have three pairs of thoracic legs and four pairs of abdominal prolegs. The corbie moth (Fig. 15.26) female is about 20 mm long with a wingspan of nearly 40 mm and the male is slightly smaller. The forewing is fawn with grey markings, while the hindwing is uniformly light brown. The
Corbies and grassgrubs Lepidoptera: Hepialidae N (See also Chapter 13 Pastures—summer rainfall:
flatheaded pasture webworm, roundheaded pasture webworm, Ebor grassgrub, Tindale’s grassgrub.)
Corbies and grassgrubs are all Australian natives and are characteried by their larvae building vertical tunnels in pastures. Flatheaded and roundheaded pasture webworms and the Ebor grassgrub have been recorded damaging summer rainfall pastures (Chapter 15). In winter rainfall pastures, the underground grassgrub, corbie, winter corbie, oxycanus grassgrub and the lesser ghost moths have been recorded in winter pastures. They have life cycles of 1 or 2
Fig. 15.25. A winter corbie larva in its silk-lined tunnel. (DPI&W Tas: C. Young)
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Fig. 15.26. An adult winter corbie moth (17 mm). (DPI&W Tas: L. Hill)
winter corbie moth is about the same size but the wing markings are less extensive. Host range: Recorded as primarily grass-eaters, but may eat other pasture species such as clovers although they avoid flat weeds. Life cycle: These two species have a 1-year life cycle similar to that of the underground grassgrub. Winter corbie moths fly during evenings between late November and midJanuary, while corbie moths fly somewhat later, during evenings between late January and midMarch. Eggs are laid, usually in rank cover, in up to three batches totalling about 2000 for winter corbies and 500 for corbies. Eggs of the winter corbie hatch after about 3 weeks and those of the corbie hatch about 8 weeks after laying. About 6 weeks after hatching, larvae have dispersed from where they were laid as eggs. They construct vertical silk-lined tunnels (Fig. 15.25) and feed on the surface at night, biting off foliage at ground level and dragging it into their tunnels. During the day they shelter in their tunnels and eat stored herbage. Damage by winter corbie is usually not evident until mid July and by corbies until August (Martyn 1958). Larvae usually pass through eight larval instars, the last five to six of which are typically spent in the same tunnel. The fully grown caterpillar finishes feeding then seals its tunnel with a silken cap. Pre-pupae and pupae of winter corbies may be found in their tunnels in November, and corbies in mid-January. The life cycle of the winter corbie includes eggs and young larvae being in the field during summer, where they may be exposed to desiccation, except in local moist areas and during years of favourable summer rainfall. Overhead irrigation, including that used on turf, may favour survival of eggs and larvae.
In contrast, eggs and young larvae of corbies occur in the field during autumn, and their survival into winter is favoured by dry conditions and friable, well-drained soils. Wet winters increase the mortality of corbie larvae compared with dry winters. A similar mortality is induced by flood irrigation. Excess pasture, including dead grass, during summer and autumn appears to provide shelter and increased larval survival. Risk conditions: Winter corbies damage Tas. pastures during winter and spring. Wet summers and abundant pasture growth appear to increase the likelihood of winter corbie damage during the following winter. Corbie damage is noticeable in August and severe damage develops from mid-September until late November. Drier than average winters increase the likelihood of corbie damage during the following spring. Outbreaks of the two species appear to be independent of each other. Damage: Damage in spring can be rapid, with extensive areas being denuded after several weeks. Denuded areas are invaded by weeds that may persist for a number of seasons (Fig. 15.27). In southern NSW, the bare areas created by winter corbies provide favourable oviposition sites for blackheaded pasture cockchafer in the following season. In Tas., winter corbie feeding can greatly reduce the grass component in dairy pastures increasing the risk of bloat in cattle (McQuillan and Ireson 1987). Monitoring: Closely inspect pastures in April (winter corbie) or June (corbie) and look especially where there are tufts of dead grass for evidence of feeding on the new shoots. Heavily
Fig. 15.27. Corbie damage the previous year reduces the grass component of the pasture and allows invasion by weeds. (DPI&W Tas)
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infected paddocks have a biscuit-brown appearance, rather than light green.
not recorded as a pest. F. polyspila is recorded from SA.
One method of estimating larval densities includes removing the top 2.4 cm of soil in a 30-cm square, and placing a metal plate over the cleared square. After 24 hours, remove the plate and count the number of holes that have been opened by resident larvae (Miller and Martyn 1952). This should be repeated in a number of sites to obtain a representative sample. A more rapid alternative method is to use a spade to remove a square sod of pasture which is broken over a bag and the larvae counted. Counts from about 30 sites are converted to numbers per square metre.
Pest status: F. polyspila: minor in SA, restricted, irregular.
Action level: Larval densities of 30 larvae per square metre may cause significant pasture damage (McQuillan 1985b), but in some areas of Tas. larval grassgrub densities (all species) as low as 24 per square metre in May and August caused damage, while densities of about 75 per square metre severely damaged pasture (Hardy 1974). Cultural control: Heavy grazing of pastures during summer reduces dead herbage and exposes corbie eggs and larvae to desiccation (Reid and Hardy 1978). Biological control: A number of pathogens have been identified causing disease among Tas. corbies, including the fungus Cordyceps oncoperae, Pandora gammae, a fungus infecting fourth- and fifth-instar larvae of winter corbies, protozoa, Vavraia sp., Mattesia sp., and the nematode Steinernema feltiae, recorded as infecting fifth- to seventh-instar larvae of corbies (Wright 1993). Chemical control: May be cost-effective but timing is important. Sprays applied in August against corbies and May against winter corbies are reported as giving satisfactory control. Young larvae may be spot-sprayed when they are still feeding on the surface. Herbage should be grazed short prior to application.
Host range: F. polyspila is recorded as eating veldt grass (Erharta sp.) and lucerne. Life cycle: F. simulans has a 1-year life cycle (Hardy 1973a) but the life cycle of F. polyspila has not been studied. F. polyspila occurs in dry areas (300 mm) of SA (Fig. 15.19). Adults have been observed flying in May. Larval tunnels up to 20 cm deep have been observed in November and mature larvae have been observed during January. Damage: Areas up to 8 ha of pasture have been denuded of grass and damage to lucerne has been reported.
Oxycanus grassgrub Oxycanus antipoda (Herrick-Schäffer) (= O. fuscomaculatus Walker) Lepidoptera: Hepialidae Pest status: Major along the north-west coast of Tas. and King and Flinders Islands, where it is a restricted and irregular pest (McQuillan and Ireson, 1987). Minor, restricted and irregular in south-east Australia and south-west of WA. Identification: New tunnels (6 cm deep) with a single entrance appear in the field in December, and by mid-January the tunnels are about 12 cm deep with silk lining one side of the tunnel and with a single entrance covered by a small mound of soil and debris held together by silk. By March, the tunnels are about 15 cm deep with two entrances, and by August the tunnels are 25 cm deep, 1 cm diameter and with two silk-lined entrances 7–10 cm apart (Fig. 15.28). The newly
Ghost moths Fraus simulans Walker and Fraus polyspila Meyrick Lepidoptera: Hepialidae Distribution: Both are Australian natives. Fraus simulans occurs in NSW, SA, Vic., and Tas. but is
Fig. 15.28. Removal of topsoil to show tunnel holes of Fraus polyspila near a 5 cent coin. (PIRSA: P.R. Birks)
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hatched larva is 3 mm long with a brown head and grey body which, by the following January, has turned to fawn on the 30 mm long larva. In May, the 1-year-old larva has grown to 70 mm long and 7 mm wide with a greenish-brown colour. The pupa is 30–40 mm long and 7–10 mm diameter. Adult females have a wingspan of about 73 mm and the smaller males 61 mm. The forewings are brown and distinctively patterned in the males, but less so in females (Hardy 1974). May be confused with: Larvae of corbies and winter corbies, but these are shorter than oxycanus larvae. Oxycanus grassgrub is distinguished by having a tunnel with two entrances. The tunnel is not vertical, the amount of silk lining the tunnels is less than in corbie tunnels and larvae do not have silk-lined runways leading to their feeding areas. Life cycle: Oxycanus grassgrub has a generation time of 2 years. Mating flights are recorded during late April–early May. Mating is nocturnal, during which low-flying males perform a zigzag mating flight then mate with females on grass stems. Eggs are laid during April–May, often following rain. The eggs probably do not hatch until September–October. The first two larval instars live on the surface in silk-lined horizontal runways constructed among dead grass. Between December and March these young larvae may be found in the field in vertical tunnels, together with the fully grown larvae of the previous year’s generation. By March, the tunnels are 15 cm deep with two entrances, and by the following August the tunnels are about 25 cm deep, 1 cm diameter with the two entrances about 7 cm apart. The larvae pupate between February and March with a pupal stage of 8–12 weeks (Hardy 1974). Host range: Larvae are selective grass-feeders but will eat clovers when larvae are dense. Risk period: December–February. Outbreaks do not appear to be related to those of corbies and winter corbies. Damage: Larvae graze new grass growth after autumn rains reducing pasture available to stock in autumn, winter and spring; however, during spring and early summer clover pastures grow in infested areas more prolifically than uninfested areas, the result of selective feeding (Hardy 1974).
Monitoring: See corbie and winter corbie. Action level: See corbie and winter corbie.
Underground grassgrub Oncopera fasciculata (Walker) Pest status: Minor in SA and Vic., restricted, irregular. Identification: The presence of grassgrubs in a pasture is usually indicated by their vertical, silk-lined tunnels in the soil capped with webbed plant debris. From the entrance of the tunnel, runways along the surface, partly lined with silk, lead to feeding areas. Eggs are ovoid, 0.7 × 0.5 mm. Newly hatched larvae are 2.6 mm long, the head and first thoracic segment are brown, the rest of the body is translucent. The fully grown larva is 50–60 mm long with three pair of thoracic (true) legs, four pairs of abdominal prolegs and a terminal pair of anal prolegs. Adult moths have a brown patterned forewing and unpatterned hindwing (Fig. 15.29). May be confused with: Blackheaded pasture cockchafer whose surface-feeding produces bare patches but produces surface mounds without silk. Host range: Annual grasses and clovers, but will also feed on perennial grasses and other pasture species. Life cycle of the underground grass grub in SA and Vic.: There is a single generation each year. Females lay soon after emerging from their pupal case, and may lay 1000–1500 eggs in their lifetime. Eggs are laid in groups on the ground. They hatch after 3–5 weeks, mostly during October and early November. For about 2 weeks the young larvae live in silken shelters among surface debris, after which they make individual vertical tunnels in the soil in which they spend the remainder of their larval and pupal life. By December, larvae are about 13 mm long with a tunnel of 20–30 mm deep, by March the 30 mm long larvae have extended their tunnels up to 90 mm deep and by late June–early July the 45–60 mm long mature larvae live in a tunnel up to 175 mm deep. Pre-pupae may be found in tunnels during July, changing into pupae after 2 weeks from which adults emerge after a further 409
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a
b
Fig. 15.30. Underground grassgrub damage to pasture. (PIRSA: P.G. Allen)
Damage: The first sign of damage is thinning of new growth. Heavily infested pastures may be eaten out (Fig. 15.30), followed by weed invasion. Larval densities of 65 per square metre during the risk period eat new pasture as it appears, over 100 per square metre eat both new feed and old dry pasture (Madge 1954). Action levels: Between 10 in Vic. (Dunn and Miles, undated) or about 40 per square metre in southern NSW (Goodyer, unpublished) are suggested.
Fig. 15.29. (a) Underground grassgrub adult male in side view (15 mm long), and (b, upper) male, wingspan 45 mm, (lower) female, wingspan 50 mm. (SARDI: P.T. Bailey)
2–3 weeks. Adults shelter inactive on the ground under foliage during the day and make low, dispersal flights at night soon after emergence during early spring September–October). They then make distinctive pre-mating flights with audible humming at dusk, before mating on foliage. Adults do not feed, and probably die after a short period of egg-laying (Madge 1954). Risk period: April–July, when grassgrubs are actively feeding and pasture is scarce. Monitoring: In late autumn or winter, take 30 or more sod samples (30 × 30 cm) across the pasture with a spade to a depth of 25 cm, carefully break the sod on a bag, count the larvae and convert the tally to numbers per square metre.
Cultural control: Excess herbage during summer, especially if there has been summer rain, favours egg-laying and build up of larval numbers. Heavy grazing in late spring or summer exposes young stages to desiccation and trampling by stock. Chemical control: May be cost-effective if pastures are sprayed in autumn, winter or early spring.
Pasture tunnel moths Philobota spp., including P. productella (Walker) Lepidoptera: Oecophoridae Distribution: Native to Australia. Recorded in SA and Vic. Pest status: Minor, restricted to pastures in high rainfall (greater than 500 mm) areas, irregular. Identification: The larva is a grey caterpillar with a black head growing to 25 mm long and
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2 mm thick (Fig. 15.31). Moths are elongate, 20 mm long, and creamy white in colour.
a
May be confused with: Other insects constructing vertical tunnels in pasture. Pasture tunnel moths have entrances to tunnels raised about 10 mm above the ground, and constructed of fragments of vegetation, sometimes associated with areas of blackheaded pasture cockchafer. Host range: Annual and perennial grasses, and clovers. Life cycle in pastures: Adults have been recorded at lights during November–December, but they may be active for longer in the summer. Eggs appear to be laid by late summer. Larvae construct silk-lined tunnels (Fig. 15.31) of about 3 mm diameter to a depth of some 75 mm. They emerge from the tunnels at night to feed on surrounding pasture.
b
Risk period: Between autumn rains and December. Damage: Dense colonies have been recorded as causing local defoliation of pastures. Densities of over 70 per square metre appear to be damaging (Allen, unpublished).
Fig. 15.31. (a) Larval pasture tunnel moth (20 mm long) removed from its tunnel (Graphic Science: © Denis Crawford) , and (b) aboveground entrances are constructed of fragments of vegetation (SARDI: G.J. Baker) .
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LEGUME PASTURES C. Pavri
Legume dominant pasture
GROUP
PEST (major pests in bold)
mites
balaustium mite blue oat mites bryobia pasture mite redlegged earth mite lucerne flea black field cricket wingless grasshoppers and other grasshoppers (see Chapter 18) cowpea aphid bluegreen aphid pea aphid spotted alfalfa aphid sitona weevil small lucerne weevil vegetable weevil (see Chapter 5) blackheaded pasture cockchafer redheaded pasture cockchafer armyworms (see Chapter 2) clover casebearer corbies, grassgrubs pasture day moth
springtails crickets and grasshoppers aphids
weevils
scarab beetles caterpillars
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Legume pastures may serve a dual purpose in winter rainfall cropping systems: they are fodder for grazing stock and, when integrated into a cropping rotation, they provide nitrogen for the following crops (ley farming). Pest control strategy in the legume ley-farming system is to support legume pasture establishment and self-regeneration by controlling pest damage, increasing feed availability during winter by preventing pestfeeding and maximising seed production by reducing pest damage during pod set and seed fill. Medic (Medicago spp.) pastures are grown on alkaline soils in 250–450 mm rainfall zones in SA, Vic. and WA, while clovers (Trifolium spp.) are grown on acidic soils in 350–450 mm rainfall zones of NSW, SA, Vic. and WA. Pest risk to annual medic pastures is reduced if no legumes are grown in the paddock during the previous season, if autumn rains come early, if more than 500 plants per square metre germinate during autumn and they grow vigorously to allow regular grazing. Conversely, pastures are liable for pest damage if there is carryover of redlegged earth mites (RLEM) and lucerne flea (LF) eggs, autumn rains are late, plant density is less than 200 plants per square metre and herbage production is slow. Aphids, especially bluegreen aphids, may reduce production of herbage and seeds. Nitrogen fixation may be reduced by sitona weevil larvae feeding on medic nodules. During spring, the need for a seed bank for regeneration in later years needs to be estimated. The more years to elapse before the next anticipated ley pasture phase, the greater the quantity of hard seed needed in the current spring. Indicators of poor seed set in spring include low plant density and failure to control RLEM, LF and aphids during autumn and winter (Taverner 1995). Medics and clovers grown for commercial seed production are particularly susceptible to insect damage.
Fig. 15.32. Adult balaustium mite (length 2 mm). (DAFWA: S. Micic)
Identification: Adult balaustium mites have a brown–red body and bright red legs. The body is covered with short bristly hairs (Fig. 15.32). The adult mite grows to almost twice the size of redlegged earth mite. The nymphs are smaller and bright red in colour. May be confused with: Redlegged earth mite (Halotydeus destructor) and bryobia mite (Bryobia praetiosa). Host range: Feeds on broadleafed weeds, pasture legumes, grasses, cereal crops, lupins and canola and is also predatory on other mites. Life cycle on winter rainfall pastures: Oversummering eggs hatch after rain, but do not have a cold temperature requirement like redlegged earth mite. Provided green material is present, balaustium mites will survive if they hatch with summer rainfall. Newly hatched larvae have six legs and are orange in colour; nymphs and adults have eight legs and
Balaustium mite Balaustium medicagoense Meyer and Ryke Acarina: Erythraeidae Distribution: Originally from South Africa, now present throughout the Mediterranean climate areas of southern Australia. Pest status: Minor, restricted, irregular.
Fig. 15.33. Balaustium mite feeding areas on lucerne leaves. (SARDI)
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gradually darken as they mature to a greyish, brown–red colour. Development from egg to adult takes about 5–6 weeks. Several generations can occur each year (Halliday 2001). Risk period: Autumn to spring, but especially when plants are at the seedling stage. Damage: Balaustium feeding bleaches leaves, resulting in wilting and even death of the plant, especially when plants are water-stressed. Monitoring: Balaustium mites are easiest to find on plants in the mid to late afternoon when the weather is warm. Even quite high populations of balaustium mite present in pastures are unlikely to cause significant damage, whereas in germinating crops damage may result. Chemical control: No chemicals are currently registered for control of balaustium mite, and some chemicals used for the control of other mites have been found to be ineffective, even at high rates. Cultural control: Control of summer weeds in and around pastures and cropping paddocks will prevent the build up of mite populations. Natural enemies: Not known.
Bryobia pasture mite Bryobia praetiosa Koch Acarina: Tetranychidae Distribution: Widely distributed in North and South America, Europe, Asia, Africa and Australia. Pest status: Minor, widespread, irregular. Identification: Adult mites are 0.75 mm long. The body is oval-shaped, flattened dorsally and rusty brown, pale orange or olive in colour, and the eight legs are pale red–orange. The front legs are very long and held out in front of the body, often being mistaken for antennae (Fig. 15.34). The young are smaller and bright red with pale legs (Fig. 15.34). Compared with redlegged earth mites, bryobia mites are sluggish and slow moving on plants. May be confused with: Redlegged earth mite, but with closer inspection the body colour is obviously not black like redlegged earth mite. The long front legs held out in front of the body are also very different to the more uniform leg
Fig. 15.34. Eggs and adult bryobia mite (top left), sixlegged nymph (0.4 mm) (bottom left), and adult bryobia mite (right) (length 0.75 mm). (SARDI: P.T. Bailey)
length of redlegged earth mite. Bryobia nymphs are bright red and may be confused with the nymphs of Balaustium medicagoense that are also bright red. Host range: There are a wide range of herbaceous hosts including pasture legumes, grasses and weeds, as well as many crop species, especially canola and lupins. Life cycle on winter rainfall pastures: Adult bryobia are most active in late spring, summer and autumn. Most mites overwinter as eggs, but all life stages can be present in protected areas. Overwintering eggs hatch as conditions dry and warm in spring. Newly hatched larvae have six legs and are bright red, turning dark grey–green in a few days. These then pass through two eight-legged nymphal stages before moulting to the adult stage. Each stage lasts between 2–6 days and the life cycle from egg hatch to adult is usually completed in 1 month. There are several generations a year. Risk conditions: Early autumn. Pastures germinating in paddocks with summer or early autumn weeds and warm dry conditions after germination are most at risk. In years with cool, wet autumns, bryobia mite is unlikely to cause damage at germination. Damage: On succulent growth of herbaceous plants, the damage may initially resemble leafminer damage. The mite creates long trails composed of whitish grey spots on the upper leaf surface of the food plant. On grasses, bryobia feeding results in leaf-silvering similar to the damage from redlegged earth mite. Bryobia mite rarely causes economic damage to pastures. Monitoring: Monitor together with other pasture mites and lucerne flea.
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Chemical control: Many of the insecticides commonly used for mite control are not registered for control of bryobia mite. Insecticide rates commonly used for redlegged earth mite, blue oat mite and lucerne flea control are not effective against bryobia mite and advice should be sought on effective rates. Cultural control: Summer weeds in and around pastures should be controlled. Heavy grazing of pastures in spring and cropping paddocks will prevent the build up of mite populations.
Redlegged earth mite Halotydeus destructor (Tucker) Acarina: Penthaleidae Distribution: Originated in South Africa, now found in Australia and New Zealand. In Australia it is widely distributed in winter rainfall dominant regions of southern Australia, where winter (May–October) rainfall is over 205 mm, summer (December–March) rainfall is less than 225 mm and the mean monthly maximum temperature of the hottest month is less than 33°C. Pest status: Major, widespread, regular. Identification: The adult mite is about 1 mm long, with velvety black body and eight red legs (Fig. 15.35). May be confused with: Redlegged earth mite is similar in appearance to blue oat mite; however,
Fig. 15.35. Adult redlegged earth mite (length 1 mm) (upper right), and blue oat mite (lower left) showing the red oval area on its back. (CSIRO Entomology: J. Green)
Fig. 15.36. Aggregation of redlegged earth mites feeding on subterranean clover cotyledon. (CSIRO Entomology: K. Gaull)
blue oat mite can be distinguished by a small oval red area in the middle of the back (Fig. 15.35). Damage caused by blue oat mites is the same as that of redlegged earth mites, but blue oat mites feed mostly on grasses and cereals. Damage to leaves of legumes may be confused with that of lucerne flea, but redlegged earth mite damage appears as a silvering of the leaf surface (Figs 15.37 and 15.38), while lucerne flea damage appears as thin, transparent windows (Fig. 15.42). Hosts: The redlegged earth mite damages all field crops and pastures, especially at seedling stage. It is a major pest of legume pastures and canola. Life cycle on winter rainfall pastures: The redlegged earth mite is active in the cool, wet months from May to November. They hatch in autumn at the break of season from oversummering eggs that have been in a state of arrested development (diapause) since the end of the previous spring. Hatching is triggered by a
Fig. 15.37. Redlegged earth mite feeding damage on subclover seedlings. (CSIRO Entomology: C. Pavri)
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optimum time for monitoring is on fine, still days. Approach very quietly as mites will disperse quickly if disturbed. If no mites are seen on the plants, look carefully at the soil surface to detect movement and then identify if it is redlegged earth mite. A hand lens may be used to detect newly hatched larvae and young nymphs.
Fig. 15.38. Redlegged earth mite feeding damage to mature subclover pasture. (CSIRO Entomology: C. Pavri)
significant rainfall event combined with a period of 7–10 days where the mean daily maximum temperature is below 21°C. Eggs hatch into sixlegged larvae, and then develop through three nymphal stages into adults. Nymphs and adults have eight legs. During winter, the redlegged earth mite passes through three generations on average, each lasting about 8 weeks. When conditions are favourable their numbers can increase rapidly, with peaks in autumn and/or spring. The first two generations of adults lay winter eggs, usually on the under-surface of the leaf of food plants. These hatch normally after 10 days. In early spring, oviposition ceases and the mites produce oversummering eggs that are retained within the body of the female mite. In late spring, when temperatures increase, rainfall diminishes and the pasture senesces, the mites die and the adult female bodies, packed with eggs, lie on the soil surface throughout the summer (Ridsdill-Smith 1997). Risk period: Autumn–spring, but especially at germination. Damage: Damage is visible as a silvering of the upper surface (Figs 15.37 and 15.38) of the leaves and results in seedling mortality and reduced growth of legume seedlings during autumn in germinating pastures. There is also reduced production and quality of older green plants during the growing season and reduced seed yield of legumes in spring. In winter pastures, the majority of damage and production loss occurs to the subterranean clover or annual medic component of the pasture. Monitoring: Monitor pastures regularly from the time of first emergence of seedlings, especially when weather conditions meet the criteria for hatching of the oversummering eggs. The
Action level: Any sign of mite activity or damage at germination warrants control. At other times of the season, feeding damage to more than 20% of the leaf area may warrant control. Chemical control: Treating seed with systemic insecticide before sowing pastures protects seedlings from attack. Chemical sprays do not kill mite eggs so it is important to time sprays when most mites have emerged. Spraying should be timed for autumn or spring. In autumn, chemicals should be applied after the break of season, and after all oversummering eggs have hatched but before adult mites start laying eggs. Post-emergent sprays in autumn will provide a benefit to seedlings at that time; however, significant damage may already have occurred by the time the mites are seen. Mites are also likely to build up during the winter, so that by spring damaging populations may be indicated by monitoring. Spraying in spring after the adults have stopped laying winter eggs and before they start to produce oversummering eggs is more effective than spraying in autumn. Spraying at this time benefits production and seed yield and, if the mites are killed before they produce oversummering eggs, the generation of the following autumn is prevented from developing. The optimum date for spraying on particular properties in spring may be estimated by using TIMERITE® (Ridsdill-Smith and Pavri 2004), a CSIRO–Australian Wool Innovation information package on redlegged earth mite. Cultural control: Redlegged earth mite populations thrive in tall dense pastures, which provide the optimum microhabitat for survival. Heavy grazing in winter and spring reduces mite populations. Capeweed and other broadleafed weeds provide large rosette leaves under which the mites can shelter. Control of the broadleafed weeds in summer can reduce mite populations in autumn. Grazing pasture short in spring reduces mite numbers.
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Fig. 15.39. Anystis wallacei feeding on redlegged earth mite (right). (DAFWA)
Host plant resistance: New varieties of annual pasture legumes are currently available that are less susceptible to mite feeding. Natural enemies: A predatory mite, Anystis wallacei Otto (Anystidae) (Fig. 15.39), was imported from France to Australia in 1965 for biological control of the redlegged earth mite, and has established at some sites where it has caused significant mortality of redlegged earth mites. Its effectiveness as a predator is limited by its slow dispersal.
Lucerne flea Sminthurus viridis (Linnaeus) Collembola: Sminthuridae Distribution: Originated in Europe, now found worldwide. In Australia, it is an established pest in southern agricultural areas of SA, Tas., Vic. and WA, and is particularly abundant in areas with a Mediterranean climate (warm, dry summer and cool, wet winter). Recently, it has been recorded as a pest in NSW.
Fig. 15.40. Adult lucerne fleas (3 mm long). The ‘spring’ can be seen protruding from the back of the body. (SARDI)
present in pastures, although they are usually less numerous (Ireson 1993). Host range: Both nymphs and adults are pests of broadleaf pasture plants such as clovers, medics, lucerne, serradella and capeweed. Although not a favoured food source, lucerne flea will damage ryegrasses. Life cycle on winter rainfall pastures: Lucerne flea has six generations a year between autumn and late spring. The number of generations and length of each generation is temperaturedependent and the length of each generation may vary from 3–5 weeks. The first generation develops from oversummering eggs after the onset of favourable moisture and temperature
Pest status: Major, widespread, regular. Identification: Adults are 2–3 mm long with a globular-shaped abdomen, wingless and with light green or yellow markings and darker, irregular pigmented areas over the body (Fig. 15.40). Newly hatched nymphs are 0.5 mm long and pale yellow; young nymphs resemble the adult, but are smaller. When disturbed on the pasture they will ‘spring’ into the air. May be confused with: Feeding damage on pasture legumes can be confused with redlegged earth mite damage. Other species of lucerne flea, introduced and native, may also be
Fig. 15.41. Lucerne flea with eggs (0.3 mm diameter) covered in excreted soil. (TIAR: J. E. Ireson)
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a
establishment. Activity may be detected by walking through the pasture and observing the lucerne flea springing from plants as they are disturbed or by beating into an ice-cream container. Action level: As a guide, an average of 20 small holes per trifoliate legume leaf may require treatment. In pastures, it is cost-effective to spray only severely damaged areas.
b
Fig. 15.42. (a) Medic leaf showing window created by lucerne flea, and (b) clover badly damaged by lucerne flea. (SARDI)
conditions, usually around April. The nymphs moult several times before reaching the adult stage which takes about four weeks. The last generation of females lays oversummering (diapause) eggs on the soil surface that do not hatch until favourable conditions return the following autumn. In mid to late spring, lucerne fleas die with the onset of warm dry weather, leaving oversummering eggs on the soil surface. To protect the eggs from desiccation and heat, the female lucerne flea covers the eggs with excreted clay soil and fluid (Fig. 15.41). Lucerne fleas are more abundant in regions with heavier soils (McDonald 1995).
Chemical control: If lucerne flea regularly causes damage, seed-dressing insecticides should be applied when sowing or reseeding pastures. If damage warrants control, spray pastures about 3 weeks after the lucerne flea eggs have hatched in autumn. At this time, all of the oversummering eggs should have hatched and the first generation females will not have started laying eggs. If lucerne fleas are still observed after the initial spray, a follow-up spray 4 weeks later may be required. Springspraying can reduce the number of lucerne fleas the following autumn by preventing the last generation of females from laying oversummering eggs. For effective control, it is important to heavily graze the pasture before spraying and to ensure that complete spray coverage is achieved in the affected area of pasture (Jennings 2002). Natural enemies: In south-eastern Vic., Tas., and southern WA, an introduced predator, the spiny snout mite, Neomolgus capillatus (Kramer) Acarina: Bdellidae, significantly reduces lucerne flea numbers in pastures (Ireson and Webb 1996) (Fig. 15.43.) In WA, the pasture snout mite, Bdellodes lapidaria Acarina: Bdellidae (Fig. 15.1), is also an effective predator.
Risk period: Autumn–spring, but especially at pasture germination. Damage: Lucerne flea feeds on the soft tissues of the pasture plants, stripping the epidermis and feeding on the soft cells below, leaving a whitish film resembling small ‘windows’ between veins. When severely damaged, the leaf becomes skeletonised (Fig. 15.42). Damage to young leaves may cause the leaf to tear as it grows and this may be mistaken for insect chewing damage. Monitoring: Inspect regularly for plant damage, which can be patchy especially during pasture
Fig. 15.43. Adult Neomolgus capillatus (length 2 mm) (right) with lucerne flea (left). (DAFWA)
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Black field cricket Teleogryllus commodus (Walker) Orthoptera: Gryllidae Distribution: Throughout Australia. Pest status: Major in winter rainfall pastures, restricted to cracking clay soils (Fig. 15.45), regular. Identification: Adults are 25 mm long, brown– black in colour with two pairs of wings, long thin antennae, three pairs of legs and a pair of tapering lateral processes (cerci) at the end of the abdomen (Fig. 15.44). Their hind legs are spined and strongly developed for jumping. When not in use, the fan-shaped hindwings are folded along the back beneath the short hardened forewings. Females have a long spear-shaped ovipositor. The late-stage nymphs are uniformly brown–black, have wing pads and the females have ovipositors. May be confused with: Black field crickets are similar to Teleogryllus oceanicus (Walker) and are distinguished by recording their songs. Also encountered may be inland field crickets, Lepidogryllus parvulus (Walker) (15–20 mm long), with a light brown back and cream underside, and pygmy crickets, Pternemobius regulus, which are smaller (adults 3–5 mm long) with relatively large hind legs. The little evening cricket, Yarrita pakaria Otte and Alexander, is also an occasional pest of pasture in south-east Australia. Host range: Omnivorous, recorded as damaging pastures (ryegrass, subterranean and white clover, lucerne) cereals, field crops and horticultural crops. Life cycle on pastures: Two or three generations between spring and autumn. They overwinter as eggs in cold areas and as eggs and late-stage
Fig. 15.44. Adult black field crickets: male (left) and female (right) (25 mm long). (Graphic Science: D. Crawford)
nymphs of the autumn generation in warmer areas. Adults mate at 2–3 weeks of age. Egglaying is stimulated by rainfall and warm conditions during spring, summer and autumn. The banana-shaped eggs (2.5–3 mm long, white or yellow) are laid singly through the long ovipositor of the female, 10–40 mm deep in soil and sometimes on the surface of bare soil. Females lay eggs in late afternoon and evenings, and during cool, cloudy days. Nymphs emerging from the eggs pass through 8–12 (usually 9–10) stages before becoming adults 2–3 months later. Young nymphs are white, 2.5–3 mm long and lack wings or ovipositors. Older nymphs turn brown–black with a white stripe across the back and later develop wing buds, and the females develop ovipositors. Adult females may swarm during warm, humid evenings 1–2 days following rain. Females may disperse up to 10 km in search of oviposition sites. Population levels are largely determined by weather. Dry winters desiccate overwintering eggs and nymphs, while cold wet weather in autumn, winter or spring also causes heavy mortality. Adults and nymphs emerge from soil following showers during autumn, spring and summer, exposing them to predation by birds. Risk conditions: Damaging outbreaks may occur when mild winters, which favour survival of eggs and nymphs, are followed by warm dry summers, which crack the soil and provide shelter to nymphs and adults (Fig. 15.45). Pastures are particularly vulnerable to damage during late summer and autumn. Damage: Adults and especially late-stage nymphs feed on leaves and stems of seedlings and young plants, and on flowers and immature seeds. Seed-feeding retards pasture regeneration and allows invasion by weeds.
Fig. 15.45. Black field crickets hide in cracks during the day and emerge to feed on pastures during the night. (PIRSA: P.R. Birks)
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Monitoring: Pastures with a prior history of damage should be monitored during midsummer (December–January) to estimate field populations. Newly established crops and pastures should be monitored in cricket-prone areas.
Pea aphid (PA), Acyrthosiphon pisum (Harris)
A shelter bag method can be used to give a relative index of numbers. Place at least 10 hessian or cloth bags over cracks in the ground at a minimum of 20 metres apart in a paddock. Several times a day, count the numbers of crickets sheltering beneath the bag (Dunn and Miles, undated).
Original range included Mediterranean Basin and West Asia.
Action level: An average of five to seven crickets per bag, or one or more bags with more than 20 crickets may indicate the need for control (Dunn and Miles, undated). Control: Baiting with insecticide-treated grain (whole or coarsely ground wheat, barley or oats) effectively controls black field crickets. Baiting is most successful when there are few alternative food sources available to crickets; that is, when pasture is short and dry.
Aphids
Original range included northern Europe, now worldwide.
Spotted alfalfa aphid (SAA), Therioaphis trifolii Monell
Distribution: All these aphids have been accidentally introduced to Australia, where they are recorded as lucerne pests in all states. Pest status on pasture legumes: BGA: major. CPA: major. PA: minor. SAA: minor. All are widespread, and irregular in damaging numbers. In eastern Australia, aphids appear to have declined in pest status since the introduction to Australia of BGA and SAA in the 1970s when they caused widespread damage. This decline may be attributable to the development of aphid-resistant pasture cultivars, the build up of native and introduced natural enemies and perhaps the seasonal and agronomic conditions. In parts of WA, BGA and CPA remain major pests of pastures. Identification: See Table 15.1.
Hemiptera: Aphididae Bluegreen aphid (BGA), Acyrthosiphon kondoi Shinji Originally probably from Asia, now widely distributed.
May be confused with: BGA is distinguished from PA by size (BGA adults are smaller) and colour; however, immature PA is hard to distinguish from BGA.
Cowpea aphid (CPA), Aphis craccivora Koch
Host ranges: BGA: introduced legumes, including the Genera Medicago, Trifolium, Melilotus and Lens.
Originally probably from north Europe, now worldwide.
CPA: a wide range of introduced plant families, including legumes.
Table 15.1. Distinguishing features of some aphids commonly found on legumes
Length of adult Colour Adult markings
Activity period
Bluegreen aphid (Fig. 15.47a)
Cowpea aphid (Fig. 8.4)
Pea aphid (Fig. 15.47b)
Spotted alfalfa aphid (Fig. 15.47c)
2–3 mm Light bluish green or dark greenish blue No dark markings
2.0 mm Black
2.4–4.4 mm Pale green, yellow– green or pink Black knees. Dark joints on antennae
Most numerous in spring but also active in autumn and winter
Numerous in both autumn and spring
1.5–2.0 mm Yellow to yellowish green Six or more transverse dark bands on the back Most numerous in autumn but also active in spring and summer
No markings
Most numerous in spring and autumn but also active in summer
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a
b
c
wingless or winged. All stages may be present in aphid colonies. Colony development is dependent on temperature; it is retarded by cold temperatures and by hot summer temperatures. Winged adults disperse to new hosts. Risk period: Seedling plants and those during the first year of establishment are most at risk of aphid damage. The risk period is autumn, spring and cool wet summers when aphid damage is most likely. Early planted pastures are at risk from SAA and CPA, which are favoured by warm autumns, particularly following an early break. Damage to legume pastures: BGA damages terminal buds, upper leaves and stems, and reduces plant production. BGA feeding may reduce flowering in seed crops. Symptoms of BGA damage are stunted growth, shortened internodes and distorted leaves turning yellow. BGA is the most damaging of the pasture aphids because of its ability to increase in numbers during winter when pasture growth is slow. CPA numbers may build up during autumn and spring and may contribute to poor productivity of medic pastures in dry years. They are reported as major pests in parts of WA, perhaps attributable to widespread planting of pulse crops.
Fig. 15.46. (a) Pea aphid (3.5 mm long) is the largest but least frequently occurring aphid likely to be found on legumes (DAFWA) . (b) Wingless blue green aphids (2.6 mm long) have a bluish tinge and are frequent on legumes (DAFWA) . (c) The smallest, spotted alfalfa aphid (2.0 mm long), is distinguished by bands of dark spots (SARDI) . (A fourth aphid common on legumes, cowpea aphid, is easily distinguished by its black colour (Fig. 8.4).)
PA: introduced legumes, including Medicago, Trifolium, Lathyrus, Lens, Pisum, Vicia and Glycine.
PA is usually the least abundant of aphids on legume pastures and often occurs in association with BGA. Symptoms include dying of the tops of plants. SAA is now a minor pest, with occasional damage reported in eastern Australia and few reports of damage from WA. Aphids are capable of transmitting a number of known viral diseases within legume pastures and also between pastures and nearby legume crops.
SAA: two biotypes are present in Australia. One (sometimes referred to as form maculata) has a strong preference for lucerne. Another biotype of this aphid prefers clovers as hosts. The two biotypes are morphologically indistinguishable (Milne 1998).
Monitoring: Frequent monitoring (once or twice weekly) of seedlings and establishing plants is necessary to detect rapid increases of aphid populations. Stem samples give useful estimates of aphid density. Plant-beating into a white bucket is also used. Aphid distribution, especially CPA, may be patchy in a paddock.
Life cycle: Females of these species produce live young which grow through wingless nymph stages. Adult males and females may be
Action level: When damage is apparent, intervention may be necessary. Moderating factors include degree of aphid tolerance of the 421
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cultivar, availability of moisture and the incidence of predators and parasitoids. Chemical control: Probably cost-effective only for seed crops. Plant resistance: Varieties of clovers and medics tolerant to BGA and SAA are available but there is little tolerance to CPA in presently available varieties. Conservation of natural enemies: Broadspectrum insecticides should be avoided.
Sitona weevil Sitona discoideus Gyllenhal Coleoptera: Curculionidae Distribution: Native to the Mediterranean region, now present in Europe, North Africa, South Africa, Australia and New Zealand. Pest status: Major in SA, minor in NSW, Tas. and Vic., restricted, irregular. Identification: The larvae are small, brownheaded, white-bodied, legless grubs up to 5 mm long living in the soil near plant roots (Fig. 15.47). Adult weevils are 3–5 mm long, varying from grey to dark grey–brown with three pale, longitudinal stripes on the thorax. The snout is short and broad with antennae attached to the front (Fig. 15.48). May be confused with: Small larvae of sitona may be difficult to distinguish from vegetable weevil (Listroderes difficilis), and the small lucerne weevil (Atrichonotus taeniatulus). The closely related clover root weevil, Sitona lepidus, is present in the North Island of New Zealand but is not recorded in Australia.
Fig. 15.48. Sitona weevil pupa (length 5 mm). (DAFWA: S. Learmonth)
Host range: Lucerne, annual medics, subterranean clover. Life cycle on winter rainfall pastures: Adult weevils are active from spring to autumn. Eggs are laid in autumn in the soil around the base of plants and hatch with opening rains, when the emerging larvae burrow into the soil. Larvae feed through winter and early spring, then pupate in the soil and emerge as adults in late spring–early summer (Howes 1990). Risk period: As both adults and larvae feed on the host plant, damage can occur at all times during the growing season. Adults, especially if active at the break of season when germination occurs, will feed on the leaves and epidermis of young plants and can kill the plant. Larvae feed on root hairs and root nodules in the soil and can cause more serious damage than the adults by reducing nitrogen fixation and retarding plant growth. Damage: Adult-feeding produces U-shaped notches on the leaves, and areas of stem stripped. Larval-feeding on the root hairs and nodules may retard growth of the plant, which may appear pale and lacking vigour.
Fig. 15.47. Sitona weevil larva (length 5 mm). (DAFWA:
Fig. 15.49. Sitona weevil adult (length 3–5 mm) and the parasitic wasp Microctonus aethiopoides. The wasp lays its egg into the body cavity of the adult weevil. (AgResearch
S. Learmonth)
Lincoln: M. McNeill)
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Monitoring: Inspect pastures for signs of seedling damage. Action level: Only if adults are in large numbers and killing seedlings in autumn should chemical control be required. Chemical control: Adults can be controlled with insecticides, and if in large numbers causing extensive leaf loss, it may be worthwhile. Natural enemies: The sitona weevil parasitoid Microctonus aethiopoides (Loan) (Fig. 15.49) was introduced into Australia as a biological control agent from the Mediterranean region. M. aethiopoides lays a single egg within the weevil’s body cavity causing sterilisation within a few days and, after full parasitoid larval development, eventual death of the weevil. It has limited impact on sitona damage.
Clover casebearer (or clover seed moth) Coleophora alcyonipennella Kollar (= C. frischella (Linnaeus)) Lepidoptera: Coleophoridae Distribution: A native of Europe, now recorded from North America, New Zealand and Australia (1925), where it is recorded from NSW, SA, Tas. and Vic. Pest status: Major in clover seed crops, restricted, irregular. Identification: The adult is a small (7 mm long) dark moth (Fig. 15.50). Host range: White clover, Trifolium repens, and strawberry clover, T. fragiferum. Life cycle on clover: In seasons when clover flowers are plentiful, there may be two (spring and summer) generations and a partial third per year, but during poor seasons only one and a
Fig. 15.50. Adult clover casebearer (length 7 mm long) on clover flower. (PIRSA: P.R. Birks)
partial second generation may develop. In summer, the life cycle may be completed in 9 weeks. Adults mate at dawn and dusk on clover flowers, and females start to lay eggs 1–2 days later and lay an average of 67 eggs during a mean 12-day lifetime. Eggs are laid on flowers that are close to anthesis and hatch after about 14 days. Larvae feed on developing seeds (Fig. 15.51). Larval development up to the end of the third instar is completed in less than 2 weeks and the duration of the fourth stage is 20–30 days. The fourth-instar larva shelters itself in a cigar-shaped case. Adults emerge from pupae after 35 days in spring (November) and 10–20 days in summer. At the end of summer, the larva in its case drops to the soil surface, enters an overwintering diapause, after which it pupates to emerge as an adult during the following spring or early summer. The life cycle may be extended to 10–12 months if the overwintering larva enters diapause. About 12% of the larvae do not pupate in the first spring but remain in diapause for a further 12 months (Hardy 1983). Risk period: During flowering, in spring to early summer (mid-November to mid-January). Damage: A larva may destroy an average of 24 seeds (Fig. 15.51), the contents of about six seedpods, during its development. Average seed losses of between 3% for crops in the first year of flowering and 46% in second and subsequent years (9 and 114 kg ha–1) have been recorded (Hardy 1973b). Monitoring: The adults do not travel more than a few hundred metres and spring adult densities are the result of the amount of seed available to the larvae in the immediate area during the previous season.
Fig. 15.51. Clover flower head sectioned to show larval damage (two arrows) to developing seeds. (PIRSA: P.R. Birks)
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Chemical control: Chemicals cannot be used because they kill pollinating bees. Cultural control: Damage reduction is best achieved through management of the local moth populations to ensure seed production is not attempted in the same area 2 years in a row and by separating the new seed production area from paddocks where there was significant seed set in the previous year by at least 500 metres. Heavy grazing, especially in spring and early summer, to ensure heads are removed prior to seed maturity is important in reducing populations. Natural enemies: Pristomerus sp. (Hymenoptera: Ichneumonidae) has been recorded as parasitising less than 1% of larvae.
Pasture day moth Apina callisto (Angas) Lepidoptera: Noctuidae Distribution: An Australian native, recorded from south-eastern Australia including Tas., and south-western WA. Pest Status: Minor, widespread, irregular.
Fig. 15.52. Larva of the pasture day moth (50 mm long). (DAFWA)
Fig. 15.53. Adult pasture day moth. (DAFWA)
Identification: Larvae grow to 50 mm, are dark brown with orange–yellow markings and have two prominent yellow spots near the end of the body (Fig. 15.52) (Emery et al. 2005). Adults are distinctively coloured (Fig. 15.53). Host range: Saltbush and some herbaceous plants including Calendula spp. capeweed, clovers, Erodium spp., plantago, Malva spp. and Rumex spp. (Common 1993). Life cycle: One year life cycle. Adults fly during autumn, flying close to the ground. Unusual among noctuid moths, the adults fly during the day. Larvae feed during winter and spring. At maturity, the larva excavates a vertical tunnel in the soil and pupates in a cell at the bottom (Common 1993). Damage: Larvae cause little damage eating the leaves of broadleafed plants. They may damage young crops if their host plants are killed by herbicides (Emery et al. 2005) and they have been recorded defoliating saltbush, Atriplex nummularia.
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LUCERNE P.T. Bailey and G. Goodyer
(a) Dryland lucerne, used for seed and grazing
(b) Irrigated lucerne setting seed
seeding establishment growth flowering podding seed maturation/drying
Jun
May
Apr
Mar
Feb
Jan
Dec
Nov
Oct
Sep
Aug
Jul
Jun
May
Apr
Mar
Feb
Jan
seed harvest
First season of a perennial lucerne crop. Stages of the crop are influenced by lucerne type (winter active or winter dormant) whether grown as dryland or irrigated, and on soil, rainfall and climate. Lucerne may be seeded in spring or autumn or sometimes during winter. Emergence is usually 5–10 days after sowing. During growth, the crop may be grazed or cut for hay. Timing of cutting, grazing and irrigation are important in management of some lucerne pests. Lucerne may also be allowed to set seed for commercial seed production.
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GROUP
PEST (major pests in bold)
snails mites
pointed snails balaustium mite blue oat mite (see Grass pastures and turf) redlegged earth mite (see Legume pastures) two-spotted mite lucerne flea wingless grasshopper (see Chapter 18) bluegreen aphid pea aphid spotted alfalfa aphid lucerne leafhopper onion thrips, plague thrips, tomato thrips small lucerne weevil sitona weevil whitefringed weevils lucerne crownborer etiella moth lucerne leafroller native budworm lucerne seed wasp
springtail grasshoppers aphids
leafhoppers thrips beetles
caterpillars
seed wasp
Small pointed snail N (Main entry in Chapter 2 Cereals.)
Pest status on lucerne: Recorded as a major pest of lucerne in Tas., including Flinders and King Islands, regular occurrence.
Redlegged earth mite and blue oat mite
PAGE 426 413 426 426 426 427 498 428 428 428 429 429 430 431 431 432 432 434 435 435
Chemical control: Chemicals are the main control method. Use one or more of: seeds treated with a systemic insecticide; seeds sown into bare ground surface-sprayed with insecticide; foliar sprays. Cultural control: Clean-fallowing, and eliminating weeds from the perimeter of the field at least 1 month before planting may reduce initial numbers.
N (See Grass pastures and turf, and Legume pastures.)
Two-spotted mite
Pest status on lucerne: Major, restricted to temperate growing areas, regular.
Tetranychus urticae Koch Acarina: Tetranychidae
Risk period: Autumn, winter and spring.
N (Main entry in Chapter 3 Cotton.)
Damage: Most damage is caused to seedlings and young plants by feeding on cotyledons, leaving them white, and young leaves. Autumnseeded plants may be stunted and die, and winter production of the survivors may be reduced.
Pest status on lucerne: Minor, widespread, irregular.
Monitoring: Regular (weekly) inspections of germinating seedlings will help detect rapidly increasing numbers. Action level: When mites, or mite-feeding is detected on seedlings.
Risk conditions: Mite activity can be expected following a period of continuous hot weather and frequent application of broad-spectrum pesticides to the lucerne crop. Damage to lucerne: Mite-feeding causes white spots on leaves, usually starting from the base of the plant. In late summer, mites may be found on leaves on the top of the plant. Moderate
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numbers of mites (up to 30 mobile mites per leaflet) towards the end of summer do not appear to reduce seed production in irrigated lucerne, as the crop is usually chemically defoliated prior to harvest. Extreme chemical mismanagement earlier in the growing season may result in the entire crop becoming white from mite-feeding, progressing to premature defoliation and lack of seed production. Monitoring: Visual-sampling of basal stems will detect early mite populations. If mites become numerous on the mid to upper parts of lucerne plants, a flat, white tray can be used to beat both mites and their predators into. The floor of the tray can be ruled into convenient squares and the numbers of mites and predators can be counted in a sub-sample of squares.
Fig. 15.54. Two-spotted mite predators in lucerne crops: mite-eating ladybird adult (3 mm long) and larva (4 mm long). (SARDI: P.T. Bailey)
If visual inspection indicates mite numbers in excess of 10–20 per leaflet in the mid or upper stems of lucerne plants during December– February, use a beating tray to estimate the predator:prey ratio. Action level: If mobile mite numbers exceed the range of 10–50 mites per predator, a rapid increase in mites may be expected. Chemical control: Use of miticides on lucerne is not cost-effective. Conservation of natural enemies: Damaging numbers of two-spotted mites on lucerne are symptomatic of over-use of broad spectrum insecticides against other lucerne pests. The frequency of spraying broad-spectrum pesticides should be reduced to allow build-up of predators. Effective predators that are killed by all but low levels of insecticide residues include the tubular black thrips, Haplothrips victoriensis Bagnall (Thysanaptera: Phlaeothripidae), which are effective predators of mite eggs (Fig. 15.55) and nymphs at low mite densities. Refuges for tubular black thrips include weeds such as Solanum spp. In the absence of mites, tubular black thrips can survive on pollen. The miteeating ladybird Stethorus nigripes Kapur (Coleoptera: Coccinellidae) (Fig. 15.54) increases its numbers in response to higher densities of mites. A predator mite, Phytoseiulus persimilis Athias-Henriot (Acarina: Phytoseiidae) (Fig. 15.56) survives insecticide applications and usually appears naturally during late summer (Bailey and Caon 1986).
Fig. 15.55. Two-spotted mite predators in lucerne crops: red immature of tubular black thrips (2 mm long) eating an egg of two-spotted mite. The red immature stage of this thrips moult to a black adult. (SARDI: G. Caon)
Fig. 15.56. Two-spotted mite predators in lucerne crops: a predator mite, Phytoseiulus persimilis (orangecoloured, 0.4 mm long), near an adult and eggs of two-spotted mite on lucerne. (G. Caon)
Lucerne flea Sminthurus viridis Collembola: Sminthuridae Pest status on lucerne: Minor, restricted to temperate lucerne areas, regular. Life cycle: See Legume pastures. 427
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Damage: Most damage is caused to seedlings and young plants, especially those grown on acidic soils. Early signs include small membranous windows on leaves, which may look speckled and greyish. Severely damaged leaves may be reduced to lower and upper epidermis and main veins. Risk period: Autumn, winter and spring. Chemical control: Insecticide-treated seed, followed by one or more foliar sprays. Cultural control: Clean-fallowing, and eliminating weeds from the perimeter of the field at least 1 month before planting may reduce initial numbers. Application of lime on acid soils before sowing may slow the build-up of lucerne fleas.
Lucerne aphids Hemiptera: Aphididae Bluegreen aphid (BGA), pea aphid (PA), spotted alfalfa aphid (SAA) N (Main entry in this chapter, p. 420.)
Pest status on lucerne: Major, widespread, damaging numbers are irregular in occurrence. Distribution: Recorded as lucerne pests in all states. Identification: See Legume pastures. Risk conditions: Seedling plants and those during the first year of establishment are most at risk of aphid damage. The risk conditions include autumn, spring and cool wet summers when aphid damage is most likely. Aphid populations may rapidly increase during the period following grazing or cutting for hay. Damage to lucerne: Feeding by large numbers of SAA (Fig. 15.57) on susceptible lucerne varieties may kill establishing lucerne plants, cause defoliation of older plants and deplete nutrient reserves of surviving plants. Honeydew secreted by the aphids may interfere with cutting, drying and baling. Honeydew, together with the sooty moulds that grow on it, reduce the forage quality of lucerne. Symptoms of SAA damage are white or yellow leaf veins, defoliation from the base and wilting in moisture stressed plants.
Fig. 15.57. SAA on a susceptible lucerne leaf. (SARDI: P.G. Allen)
BGA damages terminal buds, upper leaves and stems and reduce plant production for grazing and haymaking. BGA-feeding may reduce flowering in seed crops. Symptoms of BGA damage are stunted growth, shortened internodes, and distorted leaves turning yellow. PA is usually the least abundant of the lucerne aphids and often occurs in association with BGA. Symptoms include dying of the tops of plants. Monitoring: Frequent monitoring (once or twice weekly) of seedlings and establishing plants is necessary to detect rapid increases of aphid populations. Stem samples give useful estimates of aphid density. Beating the plant into a white container is also used. Action level: In seedlings, at least two aphids per plant; in establishing plants, at least five aphids per plant; in established lucerne for haymaking, greater than 20 aphids per plant affects quality. In irrigated and high rainfall (greater than 500 mm) crops used for grazing, taking action at aphid densities of 20–40 per stem may be cost-effective, or if symptomatic wilting is evident. Densities of greater than 40– 60 aphids per stem on dryland lucerne may result in death of established plants. Plant resistance: Lucerne lines resistant to SAA and BGA are the main management strategy against lucerne aphids. Conservation of natural enemies: Important natural enemies include wasp parasitoids (Trioxys complanatus Quilis vs SAA and Aphidius ervi (Haliday) (Hymenoptera: Braconidae) vs BGA). General predators, particularly brown
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Life cycle on lucerne: There may be four generations per year. Adult leafhoppers lay eggs singly in slits, mostly in petioles. Eggs hatch in a minimum of 3 weeks. The wingless nymphs feed on the undersides of leaves (Berg and Boyd 1984). Numbers build up in spring to peak during summer. Risk period: Summer and autumn.
Fig. 15.58. A colony of spotted alfalfa aphids on the underside of a lucerne leaf. Many aphids are infected with fungus disease. (SARDI)
lacewings, Micromus spp. Neuroptera: Hemorobiidae), ladybirds (Coccinellidae), spiders and disease (Fig. 15.58) have variable effectiveness in reducing lucerne aphid populations, depending on season and locality. Natural enemies may be conserved by reduced application frequency of pesticides. Cultural control: Maintenance of plant vigour by weed control, fertilising and water management complements the use of resistant varieties and predator conservation. ‘Severe’ grazing is reported effective in managing aphids in dryland lucerne. Chemical control: Chemical control may be cost-effective in some crops during some years. Lucerne aphids have developed resistance to a range of insecticides.
Damage to lucerne: Both species suck the plant sap, but the lucerne leafhopper injects a toxin into the plant, making the leaves yellow and stunted. Feeding by vegetable leafhoppers produces white stippling on the leaves that is rarely damaging; however, some leafhoppers that occur on lucerne may transmit a number of lucerne diseases. These leafhoppers include Austroagallia torrida Evans, Batracomorphus angustatus and the common brown leafhopper Orosius argentatus (Evans) (Pilkington et al. 2004). Monitoring: Sweep net. Action levels: As a guide, 20 or more lucerne leafhoppers per sweep of a 35 cm diameter sweep net. Vegetable leafhoppers in excess of 100 per sweep may require treatment. Chemical control: Spraying is the main control of leafhoppers. Movements of leafhoppers carrying disease from adjacent pastures may be reduced by perimeter spraying (Pilkington et al. 2004) Cultural control: Early cutting of a hay crop or grazing.
Lucerne leafhopper
Thrips
Hemiptera: Cicadellidae
Thysanoptera: Thripidae
Austroasca alfalfae (Evans) and Vegetable Leafhopper, Austroasca viridigrisea (Paoli)
Onion thrips, Thrips tabaci
Distribution: Natives of Australia. Pests of lucerne in NSW, Qld and Vic.
Tomato thrips, Frankliniella schultzei (Trybom)
Pest status: Major, damage restricted to NSW and Qld, irregular. Identification: Both leafhopper adults are wedge-shaped, about 3 mm long, yellowish or yellowish green (Fig. 7.10). Nymphs are similar looking but smaller. They are mainly distinguished by damage symptoms. May be confused with: Both are similar.
Plague thrips, T. imaginis Bagnall Distribution: Recorded in all states of Australia except for tomato thrips, which has not been recorded from Tas. Pest status on lucerne: Major, widespread, regular. Identification: Adult thrips are elongate, slender, 1–1.5 mm long and yellowish-brown, brown or blackish-brown. 429
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May be confused with: Adult tubular black thrips (Fig. 15.55) which are beneficial insects in lucerne. Host range: Includes most crop and pasture plants, and weeds. Life cycle: Adults lay single eggs into plant tissue. Risk period: From flower bud formation until flowers commence to open. Damage: Flower abortion. Feeding on the minute flower buds while the entire flowering stem (raceme) is inside the bud sheath causes buds to die so that when the sheath opens, the buds are greyish-white or ‘blasted’. If the flowering stem grows out of the bud sheath, the dead buds soon drop off, leaving a bare stem or stripped raceme. If feeding occurs a little later, on the petals and tiny pods in the opened flowers, or on the developing pods, the flower petals have white areas with small black dots of thrips excreta. Damaged flowers or very young pods may fall off, leaving a partly or completely bare flowering stem. Older damaged pods may become distorted and hold a mixture of shrivelled brown seeds, malformed seeds and normal seeds (Goodyer, unpublished). This type of damage is sometimes mistakenly attributed to crop mirids that also occur in the crop during flowering. Monitoring: Early detection is important, as thrips numbers can build up rapidly. Regularly inspect crops. Shake handfuls of stems into a white plastic container. Look for thripsdamaged leaves, which have silvery-white blotches. Action level: If thrips occur in large numbers before floral buds appear. Chemical control: Chemical control may be cost-effective in seed crops.
Small lucerne weevil Atrichonotus taeniatulus (Berg) Coleoptera: Curculionidae Distribution: Native to South America, also found in USA, New Zealand and Australia. In Australia it has been recorded in NSW and WA.
Fig. 15.59. Small lucerne weevil adult (length 10 mm). (DAFWA: S. Learmonth)
Pest status: In WA, major, restricted, regular. In NSW, minor, restricted, irregular. Identification: Adult weevils are up to 10 mm long and grey in colour with some brownish mottling (Fig. 15.59). Larvae are creamy white, legless, up to 8 mm long with small pointed brown jaws. Host range: Lucerne, subterranean clover, canola and some tap-rooted weeds. Life cycle on winter rainfall pastures: Adults emerge from the soil from late summer to autumn. Males do not occur; females reproduce parthenogenetically. Weevils are flightless. Eggs are laid at the base of plants and hatch in winter when larvae burrow into the soil and feed on the roots through spring and summer until about mid-January, when they move away from the roots to pupate. The pupae remain in the soil for several weeks before emerging as adults from mid-February to March (Howes 1990). Risk period: At germination in autumn and in summer when the larvae in the soil are nearly full-grown. Damage: Larval-feeding on lucerne tap-roots in late spring may cause death of lucerne plants in summer. The damage area may increase in successive years if the weevils are not controlled. Adult-feeding on cotyledons of germinating subterranean clover in pastures can kill the young plants.
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Monitoring: Monitor after dark for adults that emerge during late summer and autumn. Damage can be detected in roots dug during spring and early summer. Action level: If adult numbers are high in February–March or at time of germination, chemical control may be cost-effective. Chemical control: Control should be aimed at adults in time to prevent egg-laying and damage by the next generation larvae to germinating pasture in autumn. Adults can be controlled with two insecticide applications: the first week after the first weevils emerge in February–March and the second 2–3 weeks later. Cultural control: Rotation of pastures or lucerne with cereal crops can reduce numbers, but a break of at least 12 months is required before lucerne is re-sown into the paddock. For rotations to be effective, all lucerne and other tap-rooted plants such as dock and sorrel must be killed during the crop phase. If some paddocks are free of small lucerne weevils, they should be quarantined to prevent transport of weevils in hay or farm machinery from infested areas. One weevil is all that is required to start an infestation in a previously clean paddock. Natural enemies: No known natural enemies.
Sitona weevil Sitona discoideus Gyllenhal Coleoptera: Curculionidae N (Main entry in this chapter, p. 422.)
necessary to protect young plants. Egg-laying can be reduced by controlling sitona in autumn.
Whitefringed weevils Coleoptera: Curculionidae Naupactus leucoloma (Boheman) and N. peregrinus (Buchanan) Distribution: Originally from South America, now recorded from Europe, North America, southern Africa, New Zealand and Australia. In Australia, N. leucoloma was first recorded in NSW in 1933 and is now recorded in all states. N. peregrinus is recorded from one locality in NSW (Zimmerman 1994) but may be more widely distributed. Pest status on lucerne: Minor, widespread, irregular. Identification: Mature larvae are white, legless grubs that grow to about 15 mm in length. Unlike many soil-dwelling weevil larvae, the white head is tucked back into the body, with only the black mandibles protruding (Fig. 15.60) (Stevens 1998). Adults of N. leucoloma are large weevils, 9–13 mm long, dark grey to dark brown with two white bands along each side of the head and thorax and a whitish band along the margin of each wing cover. The body is covered with fine hairs (Fig. 15.61). The two species are similar and separation is difficult. N. peregrinus is slightly smaller with a smaller snout (Zimmerman 1994). May be confused with: Other white legless larvae on roots include sitona weevil, but whitefringed weevil larvae are larger (about 13 mm long and 6 mm wide when fully grown).
Pest status on lucerne: Minor, restricted, regular. Damage to lucerne: Larval-feeding on nodules and tap roots reduces plant vigour. Adult feeding may kill seedlings and young plants. Feeding by large autumn populations of adults may affect hay production. Monitoring: Scalloping of leaf edges and chewed stems are signs of sitona activity. Adults shelter under stones or debris during the day, and feed from late afternoon to early morning. Regular inspection of establishing plants will detect rapid damage. Chemical control: Insecticide application may be cost-effective. Spring-spraying may be
Fig. 15.60. Whitefringed weevil larvae (length 10– 15 mm). The head is retracted and only the mouth parts protrude from the body. (CSIRO Entomology: P. Yeoh)
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Cultural control: Avoid planting lucerne directly in, or near, land on which whitefringed weevil has been detected in the previous year. Where numerous larvae have been detected, ploughing in September or October may kill mature larvae and pupae. Planting weed-free, legume-free cereal/grass cover crops will clear the land of whitefringed weevils.
Fig. 15.61. Adult whitefringed weevil (N. leucoloma, length 13 mm) showing the characteristic white stripe along the side of the wing cover. (SARDI)
Life cycle: See Chapter 4 Maize.
Natural enemies: Significant mortality of whitefringed weevil by insect-parasitic nematodes has been recorded in Vic. lucerne crops (Sexton and Williams 1981).
Lucerne crownborers
Hosts: Both species are recorded from lucerne and, in addition, N. leucoloma is reported from a wide rage of field and horticultural crops.
Coleoptera: Cerambycidae
Damage: Root-feeding by larvae on establishing lucerne plants during their first year may damage or kill plants, necessitating replanting. Damage to established plants (Fig. 15.62), in conjunction with other stresses, may reduce vigour of plants. Adult leaf-feeding on mature plants is usually not important.
N (Main entry in Chapter 7 Pulses—summer.)
Monitoring: Pre-sowing examination of the roots and soil (to 8 cm depth) of surrounding tap-rooted plants and legumes (especially those wilting or dying) in the autumn or spring prior to sowing will indicate the presence of larvae. In established crops, dying patches may be a sign of larval damage.
Corrhenes stigmatica (Pascoe) and Zygrita diva Thompson Distribution: Both are Australian natives. Pest status: Minor pests of lucerne in northern Australia. Z. diva occurs frequently and C. stigmata infrequently in Qld lucerne (Elder et al. 1992). They are more serious pests of summer pulses. Description and life history: See Chapter 7. Damage: By lucerne crownborers is recorded in old lucerne stands in Qld, where Z. diva is the more common species. Adult beetles lay eggs in the crown of lucerne plants and the larva bores into the stem, causing yellowing of the plant and stem breakage at ground level. Planting new lucerne in or near previously damaged lucerne or pulse crops such as soybeans should be avoided. Once lucerne plants are infested, larvae cannot be cost-effectively controlled with chemicals.
Etiella moth (or lucerne seed web moth) Etiella behri (Zeller) Lepidoptera: Pyralidae Distribution: East and South-East Asia, Pacific Islands and Australia. Recorded on lucerne in NSW, Qld, SA, Tas., Vic., and WA. Fig. 15.62. Lucerne root showing area of larval-feeding (SARDI)
Pest status: Major, mainly a pest in southern mainland lucerne areas, irregular.
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Fig. 15.63. Etiella development stages. From left to right: second, third, and fifth (20 mm long) larval stages, prepupa, pupa and adult. (UnivAd: A. Austin)
Identification: The mature larva has a pale golden–brown head with dark-brown mouth parts, but in young larvae the head is black. Behind the head is a black shield, but the rest of the body is green with a pinkish tinge which becomes more pronounced with age. There are several golden brown stripes along the body which are more prominent in younger larvae (Fig. 15.63). The adult is a greyish moth, 10– 15 mm long. The head is snout-like. The leading edge of the forewing has a white stripe along its full length, and at the base of the forewing there is a transverse orange band. At rest, the moths have an ‘alert’ appearance, with the front end of the body raised (Fig. 15.64). May be confused with: Larvae or their damage are unlikely to be confused with any other insect in lucerne. Hosts: Larvae of the moth have been found on a wide range of native (Acacia spp., bitter peas, bush peas, mallee bush peas) and introduced legume plants. Introduced hosts include medics, clovers, lucerne, field peas, lupins, vetch, lentils and soy beans.
Fig. 15.64. Adult moth (15 mm long) on lucerne seed pods. (A. Austin)
Life cycle on lucerne: There are probably three to four generations each year in spring, summer and autumn. After December, the generations tend to overlap, and all stages may be present in a crop. In southern Australia, adults are first seen in late September with a second peak in late November–early December. A third peak often occurs in late December to early January. The main larval attack on lucerne seed crops occurs in mid-January. In autumn, mature larvae drop to the ground and spin cocoons in the soil and do not develop into pupae until spring. Adult females can lay about 200 eggs, which may hatch within 24 hours in hot weather, but can take up to 2 weeks to hatch in cool weather; 4–7 days is the normal hatching time during summer. On lucerne plants, eggs are mostly laid on the fully formed pods while they are still green and succulent. Eggs are rarely laid on immature pods and have not been recorded from other parts of the plant. On mature pods, eggs may be found between the pod and calyx, or between the coils of the pod. Eggs are ovalshaped, 0.6 m long, colourless when first laid but soon turn pale yellow or orange when ready to hatch (Fig. 7.100). The black head of the unhatched larva can be seen through the egg. The egg hatches into a small larva (Fig. 15.63) which constructs a small funnel-like silk tube around itself, with one end attached to the surface of the pod. The larva then chews into the pod and feeds on the seed, and usually does not leave until most seeds have been eaten. The silk tube and the entry hole quickly disappear. After eating its first pod, the larva enters further pods, often leaving some seed uneaten. The fourth and fifth larval stages are too large (16–20 mm long) to enter pods. Instead, the larva meshes several pods together in a silk tent, which protects it and allows it to eat into the surrounding pods. The pest is often detected in lucerne crops at this stage, by which time some damage has already occurred. During summer larvae take between 2 and 4 weeks to develop into pupae. The pupal stage lasts 2–3 weeks. Adults may live from 1–3 weeks so that the life cycle from egg to egg is complete in about 5–6 weeks (Austin et al. 1993). Risk period: Late summer, when pods mature prior to seed harvest. 433
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Damage: Seed lucerne crops are at risk of damage from this moth. Seed damage is sporadic over years, varying from light to severe. In seed lucerne crops, larvae eat the seeds in the seed pods. A single larva may enter a number of lucerne pods. During the early stages of an attack, there are few signs of damage. Damage to forage and hay crops is usually not significant. Monitoring: The presence of lucerne seed web moths at lights indicates the need for intensive monitoring. Monitoring in seed lucerne crops should aim at detecting the presence of eggs or young larvae in the pods. This can be done by examination of mature seed pods especially when moths are abundant and, in any case, during mid-January. Examine 50–100 racemes. Detection of webbing by mature larvae is too late to prevent damage. Action level: If about five per cent of pods contain small larvae, a spray may be warranted. Chemical control: The aim of spraying is to kill adults before they lay eggs. Chemical control of the damaging stages of lucerne seed web moth is difficult because they are protected by the pod or webbing. Insecticides with fumigant action may kill some larvae but control may not always be satisfactory. It is likely that sprays applied against other pests of seed lucerne crops may coincidently control adult lucerne seed web moths in the crop. Cultural control: The main strategy for control in seed lucerne is to ensure that the final cut or grazing period occurs in mid-December, rather than earlier in December to delay the period of seed set until after the moth flights in late December to early February. Although lucerne seed web moth may live on native legumes, its numbers on these plants are very low. The source of most lucerne seed web moths in crops is probably nearby legume crops or volunteer plants, especially lucerne plants on roadsides or irrigation banks which are not cut or grazed. Removal of volunteer plants near susceptible crops may help control, although it is not know if this practice is effective. Natural enemies: A number of wasp and fly parasitoids and also a disease have been recorded from larvae and pupae. In outbreak years, these do not appear to affect more than 10% of etiella in a lucerne crop.
Lucerne leafroller Merophyas divulsana (Walker) Lepidoptera: Tortricidae Distribution: Australia and New Zealand. In Australia, it is recorded as a lucerne pest in NSW, Qld, Vic. and WA. Pest status: Major, particularly in NSW and Qld, regular. Identification: Young larvae are pale yellow while older larvae are yellowish-green to green, with a dark head. Larvae grow to 15 mm long and, when disturbed, they wriggle backwards and drop from the leaf by a thread. A larva webs together lucerne leaves into a roll in which it lives and pupates. Adult female moths are light brown in colour; the smaller males have dark markings on their wings (Fig. 15.65). May be confused with: May be confused with lightbrown apple moth (Fig. 3.10) where lucerne is interplanted with horticultural crops. Hosts: Native herbs in the Families Polygonaceae, Chenopodiaceae and Asteraceae. Introduced hosts include clover, capeweed, dandelion and sunflower, also summer pulses (Chapter 7). Life cycle on lucerne: There are a number of generations each year (seven in the Hunter Valley at about 5-week intervals during summer (Bishop et al. 1993)). Risk periods: Following cutting or grazing, when the plants are about half regrown. Damage: Feeding on terminal leaves and flowering stems results in stunted growth, reduced quality of hay and reduced seed yield, resulting from webbing of flowers. Damage is accentuated in moisture stressed dryland lucerne.
Fig. 15.65. Adult lucerne leafrollers (8 mm long): female (left) and male (right). (DPI&F Qld: D. Ironside)
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Monitoring: Estimate number of stems on which regrowth terminals are rolled. Action level: Above about 25% stem damage to 2-week old regrowth. Natural enemies: The fungal pathogen Zoophthora radicans (Erynia radicans) and a nuclear polyhedrosis virus, plus the parasitoids Apanteles tasmanica and Voriella uniseta, are reported as effective in delaying or preventing economic damage (Bishop and McKenzie 1991). Cultural control: Cut or graze as indicated by the action level.
Native budworm Helicoverpa punctigera (Wallengren) Lepidoptera: Noctuidae Pest status: Minor, widespread, irregular. Risk periods: During flowering of seed lucerne crops in summer. Spring populations may affect hay production. Damage: Larval feeding on flowers and pods reduces seed production. Young larvae feed in flower buds for 2–3 days before moving to flowers and developing pods. Monitoring: Sampling flower buds and flowers for eggs and small larvae (1–5 mm long) allows early control. Chemical control: One or two well-timed insecticide applications should prevent significant seed loss. Cultural control: Early cutting or grazing of hay crops may forestall loss of production.
Lucerne seed wasp Bruchophagus roddi (Gussakovski) Hymenoptera: Eurymelidae Distribution: Europe and west Asia, North and South America, Australia. In Australia, it has been reported as a lucerne pest in NSW, Qld, SA, Vic. and WA. Pest status: Major, damage more severe in southern growing areas than in the north, irregular.
Identification: White, legless larva in a lucerne seed. Adults are about 2 mm long with black bodies and brown, or yellowish-brown legs (Fig. 15.66). May be confused with: Seed damage (Fig. 15.66) is unlikely to be confused with any other lucerne seed-eater. Adults in sweep nets may be confused with a number of small insects visiting the lucerne crop. Host range: Lucerne. Life cycle: Adult wasps emerge in spring and lay eggs through the young, soft pod into the soft seed. The female wasp cannot penetrate the pod and seed after these have hardened. Once injected into the seed, the egg hatches to a larva, which eats out the seed contents and pupates in the hollow seed. A single adult emerges from the seed by chewing a small, circular hole with jagged margins. There are a number of generations of wasp which build up between spring and summer. These generations overlap as the season progresses, so that during summer all stages of the wasp can be found in the field at any one time. In autumn, the prepupae go into a winter diapause in the seed until the following spring. Emerging adults may sometimes be seen as a haze around harvested lucerne seed held in storage. Risk period: Summer, especially late summer for crops grown for seed. Maturing pods (greater than one-quarter to one-half formed) are not susceptible to damage. Damage: Damage is mainly to seed lucerne crops. Damaged seed does not germinate. The level of damage varies between years and localities. Damaged seeds are dull brown, dark brown or almost black, and can easily be crushed between the fingers. Monitoring: Adult wasps can be detected in crops by sweep-netting. Immature stages can be detected by pulling apart pods and examining seeds with a hand lens. Action level: Any immature lucerne seed wasp detected in the crop destined to be harvested for seed is likely to result in economic loss. Preventative control is necessary in areas with a history of damage by lucerne seed wasp. Chemical control: Only effective against adults. Since emergence is continuous during summer, 435
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pests may incidentally reduce numbers of lucerne seed wasp adults.
a
Cultural control: This is the most important control method. • Early seed harvest reduces the chances of damage from wasp populations, which peak in summer. Uniformity of crop flowering reduces the number of late-set pods. • Reservoirs of the wasp may be reduced by removal of volunteer lucerne from roadsides headlands and irrigation banks, cutting adjacent non-seed crops for hay, and removal of hay before the seed matures. Seeding of uncut lucerne may be prevented by heavy stocking.
b
• Winter carry-over of the wasp may be reduced by harvesting before any pods begin to shatter and by cleaning harvested seed and burning all seed cleanings or seed fumigation. Cultivation of the crop during autumn or winter so that seed is buried at least 5 cm may also reduce overwintering populations. Fig. 15.66. (a) Damage to lucerne seed by seed wasp larvae and emerged adults, and (b) lucerne seed in storage. Emerged adult wasps are on the wall of the container. (SARDI: P.T. Bailey)
Plant resistance: Seed damage by lucerne seed wasp to lucerne lines with tightly coiled pods is reported as being less than those with more open pods
repeated spraying may induce secondary pests and may not be cost-effective. Sprays for other
Natural enemies: None recorded in Australia. Some parasitoids have been recorded in Europe and North America
Sources of information Austin, A.D., White, T.C.R., Maelzer, D.A. and Taylor, D.G. (1993). Biology of Etiella behirii Zeller (Lepidoptera: Pyralidae): a pest of seed lucerne in South Australia. Transactions of the Royal Society of South Australia 117: 67–76. Bailey, P. and Caon, G. (1986). Predation on twospotted mite, Tetranychus urticae Koch (Acarina: Tetranychidae) by Haplothrips victoriensis Bagnall (Thysanaptera: Phlaeothripidae) and Stethorus nigripes Kapur (Coleoptera: Coccinellidae) on seed lucerne crops in South Australia. Australian Journal of Zoology 34: 515–525. Berg, G. and Boyd, M. (1984). Insects in Lucerne. The Grassland Society of Victoria. 28pp. Bishop, A.L. and McKenzie, H.J. (1991). Key mortality factors of Merophyas divulsana (Walker) (Lepidoptera: Tortricidae) larvae in the Hunter Valley. General and Applied Entomology 23: 59–64. Bishop, A.L., McKenzie, H.J. and Whittle C.P. (1993). Use of synthetic pheromone traps to time control of the lucerne leafroller, Merophyas divulsana. Entomologia Experimentalis et Applicata 67: 41–46. Carne, P. (1956). An ecological study of the pasture scarab Aphodius howitti Hope. Australian Journal of Zoology 4: 259–314.
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Carne, P.B and Chinnick, L.J. (1957). The pruinose scarab Sericesthis pruinosa Dalman and its control in turf. Australian Journal of Agricultural Research 8: 604–616. Coles, R.B. (1980). The biology of Cordyceps aphodii (Sphaeriales: Clavicipitaceae). In: Proceedings of the 2nd Australasian Conference on Grassland Invertebrate Ecology. Palmerston North, New Zealand, 22–26 May 1978. (Crosby, T.K. and Pottinger, R.P., eds). Wellington, New Zealand. Government Printer, pp. 207–212. Common, I.F.B. (1990). Moths of Australia. Melbourne University Press, Melbourne. Davidson, R.L., Wiseman, J.R. and Wolfe, V.J. (1972). Environmental stress in the pasture scarab Sericesthis nigrolineata Boisd. I. Mortality in larvae caused by high temperature. Journal of Applied Ecology 9: 783–797. Dunn, C. and Miles, M. (2000) Insectopedia. An electronic insect pest management manual for the south-eastern Australian grain and pasture pests. Department of Natural Resources and Environment. Agriculture Victoria. Emery, R., Mangano, P. and Michael, P. (2005). Crop Insects: The Ute Guide. Western Grain Belt edition. Department of Agriculture Western Australia and Grains Research & Development Corporation. Elder, R.J., Brough, E.J. and Beavis, C.H.S. (1992). Managing Insects & Mites in field crops, forage crops and pastures. Department of Primary Industries, Queensland. Fisher, D. and Learmonth, S. African Black Beetle in Vineyards. Department of Agriculture, Western Australia, Bulletin 4500. Ford, P., Olszewski, I. and Nickson, D. (2000). Argentine stem weevils. Golf & Sports Turf Australia. October 2000. Frost, W.E. (1995). The ecology of cereal rust mite Abacarus hystrix (Napela) in irrigated perennial dairy pastures in South Australia. PhD Thesis, University of Adelaide. 158pp. Given, B.B. (1950). Notes on the Aphodiinae of Australia (Coleoptera: Scarabaeidae). The Aphodius tasmaniae, howitti, yorkensis, andersoni complex. The Proceedings of the Linnean Society of New South Wales 75: 153–157. Goodyer, G.J. (1977). Root-feeding beetle larvae—pests of crops and pastures. Agricultural Gazette of NSW 88: 42–43. Goodyer, G.J. (1983). Underground grass-grubs. Agfacts. Department of Agriculture, NSW. Goodyer, G.J. (1984). Black-headed pasture cockchafer. Agfacts, Department of Agriculture, NSW. Goodyer, G.J. (1985). Scarab grubs in Northern Tableland pastures. Agfacts. Department of Agriculture, NSW. Halliday, R.B. (2001). Systematics and biology of the Australian species of Balaustium von Heyden (Acari: Erythraeidae). Australian Journal of Entomology 40: 326–330. Hardy, R.J. (1973a). The biology of Fraus simulans Walker (Lepidoptera: Hepialidae). Journal of the Australian Entomological Society 12: 113–120. Hardy, R.J. (1973b). A survey of the level of injury to crops of white clover seed by the clover seed moth Coleophora frischella (L.), in Tasmania. Tasmanian Journal of Agriculture 44: 214–216. Hardy, R.J. (1974). The biology and pest status of Oxycanus fuscomaculatus Walker (Lepidoptera: Hepialidae) in Tasmania. Journal of the Australian Entomological Society 13: 317–328. Hardy, R.J. (1976a). Observations on the pasture beetle, Saulostomus villosus Waterhouse (Scarabaeidae: Rutelinae). Journal of the Australian Entomological Society 15: 281–284.
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Hardy, R.J. (1976b). The biology and behaviour of the pasture beetle Scitala sericans Erichson (Scarabaeidae: Melolonthinae). Journal of the Australian Entomological Society 15: 433–440. Hardy, R.J. (1983). The biology and behaviour of the clover seed moth, Coleophora frischella (Lepidoptera: Coleophoridae) in Tasmania. Journal of the Australian Entomological Society 22: 7–14. Heap, J. (1998). Redheaded pasture cockchafer. Fact Sheet 430/662 PIRSA. Howes, K.M.W. (Ed). (1990). Sitona weevil (Sitona discoideus). Insect and Allied pests of extensive farming. Department of Agriculture, Western Australia. Bulletin No. 4185. Ireson, J.E. 1993. Activity and pest status of surface-active Collembola in Tasmanian field crops and pastures. Journal of the Australian Entomological Society 32: 155–167. Ireson, J.E and Webb, W.R. (1996). Redistribution and establishment of Neomolgus capillatus (Kramer) (Acarina: Bdellidae) for the biological control of Sminthurus viridis (L.) (Collembola: Sminthuridae) in Tasmania. Australian Journal of Entomology 35: 243–246. Jennings, G. (2002). Knowledge, timing, key to lucerne flea control. GRDC Advice, November 2002. McDonald, G. (1995). Lucerne Flea. Department of Primary Industries, Victoria, NRE Information Series AG0415. McQuillan, P.B. (1985a). The identification of root-feeding cockchafer larvae (Coleoptera: Scarabadaeidae) found in pastures in Tasmania. Australian Journal of Zoology 33: 509–46. McQuillan, P.B. (1985b). An assessment of pasture damage caused by hepialid (Oncopera intricata) larvae in Tasmania. In: Proceedings of the 4th Australasian Conference on Grassland Invertebrate Ecology. Lincoln College, University College of Agriculture, Canterbury, New Zealand, 13–17 May, 1985. (Chapman, R.B., ed.). pp. 7–16. McQuillan, P.B. and Ireson, J.E. (1987). Tasmanian Pasture Pests – Identification and Control. Department of Agriculture, Tasmania. Madge, P.E. (1954). A field study on the biology of the underground grass caterpillar, Oncopera fasciculata (Walker) (Lepidoptera: Hepialidae) in South Australia. Australian Journal of Zoology 2: 193–204. Martyn, E.J. (1958). Corbie and winter corbie. Tasmanian Journal of Agriculture 29: 125–131. Matthiessen, J.N. and Ridsdill-Smith, T.J. (1991). Populations of African black beetle, Heteronychus arator (Coleoptera: Scarabaeidae) in a Mediterranean climate region of Australia. Bulletin of Entomological Research 81: 85–91. Miller, L.W. and Martyn, E.J. (1952). A sampling technique for underground grass grubs. Journal of the Australian Institute of Agricultural Science 18: 110–111. Milne, W.M. (1998). Comparative performance of two biotypes of Therioaphis trifolii (Monell) (Hemiptera: Aphididae) on clovers (Trifolium) and medics (Medicago). Australian Journal of Entomology 37: 350–355. Milner, R. (1981). A novel milky disease organism from Australian Scarabaeids: field occurrence, isolation and infectivity. Journal of Invertebrate Pathology 37: 304–309. Pilkington, L.J., Gurr, G.M., Fletcher, M.J., Elliott, E., Nikandrow, A. and Nicol, H.I. (2004). Reducing the immigration of suspected leafhopper vectors and severity of Australian lucerne yellows disease. Australian Journal of Experimental Agriculture 44: 983–992. Potter, D.A. and Williamson, R.C. (1997). Behaviour and cultural control of black cutworms in golfcourse turf. Soil invertebrates in 1997. Proceedings of the 3rd Brisbane Workshop on Soil Invertebrates. 1997. (Allsopp, P.G., Rogers, D.J. and Robertson, L.N., eds). pp. 60–62.
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Power, R.J.B. (1984). Argentine stem weevil life cycle. Hortfact. The Horticultural and Food Research Institute of New Zealand. Reid, R.N.D. and Hardy, R.J. (1978). Corbie damage can be reduced by grazing management. Tasmanian Journal of Agriculture 49: 23–24. Ridsdill-Smith, T.J. (1997). Biology and control of Halotydeus destructor (Tucker) (Acarina: Penthaleidae): a review. Experimental and Applied Acarology 21: 195–224. Ridsdill-Smith, T.J. and Pavri, C.C. (2004). TIMERITE® an information package for the control of redlegged earth mites. Australian Wool Innovation Limited. Robinson, M.T. and Hoffmann, A.A. (2001). The pest status and distribution of three cryptic blue oat mite species (Penthaleus spp.) and redlegged earth mite (Halotydeus destructor) in southeastern Australia. Experimental and Applied Acarology 25: 699–716. Sexton, S.B. and Williams, P. (1981). A natural occurrence of parasitism of Graphognathus leucoloma (Boheman) by the nematode Heterorhabditis sp. Journal of the Australian Entomological Society 20: 253–255. Stevens, D. (1998). White fringed weevil life cycle. Hortfact. The Horticultural and Food Research Institute of New Zealand. Taverner, P. (1995). How to win the war against medic pests. In: Pasture Plus: The Complete Guide to Pastures. (Casey, M., ed.). Kondinin Group, WA. pp. 345–346. Wright, P.J. (1993). New records of entomopathogens infecting Oncopera spp. (Lepidoptera: Hepialidae) larvae in Tasmania. Journal of Invertebrate Pathology 61: 315–316. Zimmerman, E.C. (1994). Australian Weevils. Volume II. CSIRO and Entomological Society of America.
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16 PASTURES—WINTER RAINFALL: WEED BIOCONTROL AGENTS D.T. Briese, J. Ireson, J-L. Sagliocco, M.J. Smyth, A.E. Swirepik and P.B. Yeoh
(a) Skeleton weed in a fallow field following a cereal crop
(b) Pasture invaded by Paterson’s curse
WEED
ESTABLISHED AGENT (major control agents in bold)
docks
dock aphid dock moth gorse spider mite gorse seed weevil horehound clearwing moth horehound plume moth nodding thistle receptacle weevil nodding thistle rosette weevil nodding thistle seed fly Paterson’s curse crown weevil Paterson’s curse root weevil Paterson’s curse flea beetle Paterson’s curse pollen beetle Paterson’s curse leafminer ragwort flea beetle
gorse horehound nodding thistle
Paterson’s curse, Viper’s bugloss and Italian bugloss
ragwort
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WEED
ESTABLISHED AGENT (major control agents in bold) ragwort plume moth
skeleton weed Scotch thistle, Illyrian thistle and stemless thistle
PAGE 457
ragwort stem & crownboring moth
458
chondrilla gall mite
460
chondrilla gall midge
461
onopordum capitulum weevil
462
onopordum crown weevil
463
onopordum stemboring weevil
463
onopordum rosette moth
464
spear thistle
spear thistle gall fly
465
St John’s wort
St John’s wort stunt mite
466
St John’s wort aphid
467
St John’s wort leaf beetle
467
St John’s wort midge
468
Research on biological control agents against weeds in winter rainfall pastures has been conducted since the 1920s. CSIRO Division of Entomology, Canberra, and the Keith Turnbull Research Institute, Victoria, have been the main introduction centres. Agents are selected to attack critical parts of the weed’s life cycle and reduce its vigour, density and economic cost to the pasture industry. Some introduced agents are not released from quarantine because they are found to be not target-specific. Of those released, some may fail to establish or, if they establish, may not exert any control over the target weed. In other cases, they may be successful on their own, but often form part of a weed management strategy involving chemical herbicides, tactical grazing and other methods. This chapter includes weeds of winter pastures on which imported biological control agents exert at least some control. Also included is skeleton weed, the main impact of which is in cereal crops, but which is also a weed in the pasture fallow. Weeds of conservation significance are not included.
Docks Rumex spp. Polygonaceae Some Rumex species are native to Australia, but the most significant weeds, curled dock, Rumex crispus, broadleaf dock, R. obtusifolius and fiddle dock, R. pulcher, originated in west Asia and
southern Europe and have now spread to most continents. In southern Australian pastures, they are perennial weeds with deep tap-roots that compete with pasture plants and reduce herbage available for grazing. Plants germinate from seed in autumn and spring, during which time the plants form rosettes and develop a tap-root. Aerial growth and seed heads form between late spring and early summer, while later in the summer the top of the plant dies, leaving a dormant root during summer which re-shoots the following autumn.
Dock aphid Brachycaudus rumexicolens (Patch) Hemiptera: Aphididae Distribution: The origin is uncertain but it was first recorded in North America and is now distributed across North America, Europe, Asia, Africa and Australia. In Australia, it appears to have been accidentally introduced some time in the early 1980s. By 1993, it had been reported from all states of Australia except Qld. In WA, it is present throughout most of the agriculture areas and is predicted (via climatic modelling) to spread in southeastern Australia (Adelaide–Rockhampton– Hobart region) (Scott and Yeoh 1999). Beneficial status: Minor, widespread, irregular, depending on season.
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adults. If, however, the plant condition deteriorates or overcrowding occurs, the adults will produce offspring that will become winged adults that fly off to find another host plant. In the temperature range of 20–28°C, the population can double every 2–3 days. They are not able to grow when temperatures are too cold (< 6.4°C) or too hot (> 32°C) (Scott and Yeoh 1999). For many locations, summer temperatures are too high for the dock aphid to survive and it is assumed that re-invasion occurs each autumn from colonies located in cooler areas.
Fig. 16.1. Green and red wingless dock aphids distorting a dock leaf by their feeding. (CSIRO Entomology: P.B. Yeoh)
Identification: The dock aphid has two forms of adult; winged and wingless. Both have body lengths of 1.2–1.7 mm. Young nymphs are wingless and pale yellowish-green (Fig. 16.1). Individuals destined to become wingless adults remain this colour throughout their lives, whereas those that will become winged adults turn progressively reddish. The winged adults have a black head and thorax. Their abdomen is pale yellowish-green with a single large black patch on its upper surface and a row of large black dots running down each side. May be confused with: The dock aphid is the one most likely to be found on dock plants, and it is the only aphid having green and red wingless individuals that have short stubby siphunculi (pair of tubes at rear of abdomen) that are barely visible, whereas other aphid species have distinct tubes that are many times longer than they are wide. The common name may also cause confusion as it is shared with other aphids such as Aphis fabae Scopoli (black black bean or dock aphid) and Dysaphis radicola (Modvilko) (apple dock aphid). Host range: Polygonaceae. Life cycle: In Australia, all dock aphids are believed to be females that give birth to live offspring without the need to mate. Winged adult females fly onto a host plant and produce young that undergo four moults before becoming wingless adults. Under optimum conditions, these wingless adults produce offspring that also grow to become wingless
Damage: The dock aphid feeds preferentially on the new unfurled leaves, the growing tips of the stems and the flower buds of dock plants. Highly distorted and curled leaves, stunted stems and damaged seed result. Integrated control: The impact of the dock aphid upon the dock plant depends on how early in the season the plant is attacked. Early colonisation is encouraged by suitable alternative host plants (i.e. native docks, Muehlenbeckia and other plants from the Polygonaceae family) along river banks, under heavy shade and in other cooler areas so that local populations of the dock aphid can persist over summer where they would not otherwise survive. These areas may act as insecticide-free refugia for the dock aphid.
Dock moth Chamaesphecia doryliformis Ochsenheimer Lepidoptera: Sesiidae Distribution: The dock moth was imported to Australia from Morocco and since 1989 has been distributed as part of a biological control program to southern pastoral areas in NSW, SA, Vic. and WA. Beneficial status: Moderate, widespread and regular in WA, restricted in NSW and Vic. Identification: Moths appear around dock flowers during spring and summer. Moths have stripes across their body and clear wings which they fold wasp-like along their body. Female moths (15 mm long) are larger than males (12 mm long). Female moths have distinctive orange, black and white bands across the abdomen, while males have black and white bands across a yellow abdomen. The larva has 4 43
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a
soil level and continues to feed and enlarge the channel during summer and autumn. Larvae cease feeding during winter and spin a cocoon around themselves. Larvae pupate in the channel and the adult emerges several weeks later. Damage: Larval-feeding during plant dormancy kills many plants, and summer stresses plants. This results in the death of many and those that survive are prevented from regenerating by larval damage (Fig. 16.2).
Gorse Ulex europaeus Fabaceae: Genisteae b
Gorse is a native of central and Western Europe including the British Isles, where it occurs in native heathland and on disturbed or neglected farmland and forests. Gorse now occurs in most temperate areas of the world and is regarded as a serious weed in New Zealand, Hawaii, Chile, the western USA and Australia. Gorse occurs in all Australian states but is particularly common in south-eastern Australia, principally in Tas. and Vic. where it has invaded pastoral land, significantly reducing pasture and animal productivity. Two introduced biological control agents are now widely established on gorse.
Gorse spider mite Tetranychus lintearius Dufour Acarina: Tetranychidae Fig. 16.2. (a) Dock moth: root of a dock plant, split to show the larva (25 mm long). Larval-feeding on the taproot debilitates and kills the plant (SARDI: K. Henry) . (b) Female adult dock moth (DAFWA) .
legs and is cream with a brown head, and grows to 25 mm long. The larva channels down the root, the only external sign of which is a fine powder at the base of the stem (Fig. 16.2). May be confused with: No other larvae channel into Rumex roots. Host Range: Restricted to several Rumex species (Sagliocco and Coupland 1995). Life cycle: The dock moth has a 1-year life cycle, most of which is spent as a larva. Adults live for only a few days. Females lay eggs on the flowering stems. The hatched larva moves down the stem and channels into the root just below
Distribution: The mite, a native of Europe, was first released in Tas. and Vic. in 1998 and has since been released in southern NSW. It is now found in all gorse-infested regions of Tas. and is still spreading in Vic. and NSW. Beneficial status: Moderate, widespread but irregular in Tas. and unknown in the other states. Impact in many areas has been restricted by predation. Identification: Gorse spider mites range in size from about 0.4–0.6 mm in body length (smaller than a pin head). Immature mites have green abdomens with mouth parts and legs ranging from yellow to orange. On maturing the body reddens. The gorse spider mite forms discrete colonies (Fig. 16.3) which live in a tent-like, white web and move around the host plant ‘en masse’, feeding and web-spinning as they go (Fig. 16.4).
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temperature. The sex ratio can vary considerably and can be one male for every 20 females. Climatic conditions, particularly rainfall, have a direct impact on the establishment and population growth of the mites. On warm days they can be seen moving actively through their webbing, while on cold or wet days they protect themselves by clustering at the centre of the web, often on the leeward side of the gorse stems (Stone 1986).
Fig. 16.3. Gorse spider mite colony. (Landcare Research New Zealand Ltd)
May be confused with: Unlikely to be confused with other invertebrate species occurring on gorse due to the formation of colonies and the fine, white webbing they produce. Host range: The gorse spider mite has never been recorded from any plant other than Ulex species (Ireson et al. 2003). Life cycle: The life cycle is temperaturedependent and takes about 46 days at 15°C, about 32 days at 20°C and 20 days at 23°C. Because it is able to pass through several generations in a relatively short time at warmer temperatures, it can quickly build up in large numbers. The mites pass through six immature stages before the adult stage is reached. Adults may feed from 1–2 weeks before the female starts egg-laying. Eggs, which are brownish when mature, are scattered through the silk webbing of the colony and hatch in 1–2 weeks under warm conditions. In summer, each female lays about 40–50 eggs at the rate of one per day to a maximum of four per day, depending on
Active period: Mite colonies become visible from mid to late spring as temperatures start to increase, and are most active during the warmer summer months. Activity starts to decline as temperatures start to decrease from autumn and is at its lowest through the cooler winter months. Damage: Gorse spider mites have sucking mouth parts that pierce the plant cell wall and extract the cell contents. This ultimately results in the foliage having a bleached or bronzed appearance. Mite-feeding can significantly reduce the green weight of a gorse bush and can reduce flowering and retard growth. Integrated control: Long-term control of gorse will be reliant on the development of integrated management strategies, in which a guild of complementary biological control agents will be useful components in combination with other methods, including grazing, mulching and herbicides. Additional gorse biological control agents of European origin that are currently under investigation include the gorse thrips, Sericothrips staphylinus Haliday (Thysanoptera: Thripidae). Gorse thrips was released in Tas. and Vic. in 2001 and has now been re-distributed to SA. Gorse thrips feeds on all green gorse foliage, including seedlings, and can reduce gorse vigour and seedling survival; however, the agent spreads slowly, so any impact in the short term will need enhancement through local redistribution programs.
Gorse seed weevil Exapion ulicis (Forster) Coleoptera: Benthidae
Fig. 16.4. Gorse spider mite webbing on gorse. (TIAR)
Distribution: A native of western Europe and first released in Tas. in 1939. It is now common and widespread in Tas. and Vic., but its distribution in other states is unknown. 445
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three moults or instars and feed on the seed. After feeding for 6–8 weeks, the larvae pupate. The pupal stage lasts about 4 weeks, the adults being liberated when the pods burst. Active period: Adults occur on gorse bushes throughout the year, although peak activity occurs in spring and early summer. Eggs are laid from mid-August through spring and larvae can usually be found in pods from late September to early January. Fig. 16.5. The gorse seed weevil adult. The body is about 2.5 mm long. (Landcare Research New Zealand Ltd)
Beneficial status: Minor, widespread, regular in Tas. and Vic. Identification: Adults are small (1.5–2.5 mm) and grey with a characteristic long, curved snout (rostrum) (Fig. 16.5). They can be easily collected from individual branches by shaking them over a piece of cardboard. The yellow eggs are found singularly or in batches inside the green pods. Larvae are white, 2.0–2.5 mm in length with brown heads (Fig. 16.6). Pupae range in colour from white to grey, depending on development. May be confused with: Unlikely to be confused with other insects occurring on gorse. Host range: In Australia, gorse seed weevil feeds only on gorse. Life cycle: One generation of the weevil is produced each year. During late winter and spring, adults mate, bore a hole into the gorse pod and start laying their eggs. These hatch in about 4 weeks. Hatched larvae pass through
Fig. 16.6. Opened gorse pods showing larvae of the gorse seed weevil and damaged seed (left). (Landcare Research New Zealand Ltd)
Damage: Adults feed on young shoots and flowers, causing foliage bronzing and conspicuous holes in the flowers; however, the major damage results from larvae feeding on the seeds (Fig. 16.6). Over 90% of the seed produced during spring and summer may be destroyed. As larvae are not present during the second period of seed production in autumn–winter, a large proportion of the annual seed crop escapes attack, which limits the weevils’ effectiveness as a biological control agent. Integrated control: The integration of biological and traditional control strategies for gorse is currently under investigation. The impact of the gorse seed weevil should be greater in combination with the other biological control agents being investigated, particularly if another agent that attacks the seed produced in autumn– winter is found.
Horehound Marrubium vulgare Lamiaceae Horehound is native to temperate Eurasia, Europe, West Asia and the Mediterranean region including North Africa. From its native range, it has spread to North and South America, Australia, New Zealand, Africa and Asia. Horehound prefers alkaline soils, is drought tolerant, and is widely distributed throughout south-eastern Australia, occurring in areas receiving greater than 200 mm rainfall per annum. Horehound is primarily a weed of pastures and crops in southern Australia where it thrives on poor soil and in waste places, invading poor pastures that provide little competition. Horehound contains a bitter alkaloid, which makes it unpalatable for grazing livestock. Horehound burrs contaminate wool, reducing the value of the fleece. As well as
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being an agricultural weed of pastures, horehound has become an important environmental weed, invading disturbed native vegetation. Two biological control agents are currently established on horehound in Australia.
shortly after. Mating may last 1 hour or more, after which each female lays an average of 90 eggs. The black, small eggs (under 1 mm long) are deposited at the base of stems or within the flower calyxes. Newly hatched larvae move to the base of stems and feed on the outer cambium before feeding inside the root.
Horehound clearwing moth
Active period: Larvae feed inside horehound roots from December until October–November of the following year, when they pupate. Adults are active from late November until late December but only live for a few days.
Chamaesphecia mysiniformis (Boisduval) Lepidoptera: Sesiidae Distribution: Spain, Portugal, southern France and Morocco. First released in SA and Vic. in 1999 from one population from northern Spain. It has established in western Vic., but establishment in SA is not yet confirmed. Beneficial status: Currently minor, due to its actual restricted distribution. It has a severely damaging effect on mature plants. Identification: Adults are relatively small, the males being smaller than females (wingspan 11–24 mm). Colouration ranges from silvery to light or dark brown (Fig. 16.7), with transparent wings, especially the hindwings. The larva is ivory white with a light brown head. May be confused with: Other native species of the Sesiidae family, although none has been observed feeding on horehound flower nectar. Host range: Horehound and several other genera of Lamiaciae (Stachys, Ballota, Sideritis), none of which are important in Australia. Life cycle: The horehound clearwing moth has one generation per year. From late November through to December, adults emerge in the morning from attacked plants at the base of roots. Females resting on plants attract males with sexual pheromones and mating occurs
Damage: The larva of C. mysiniformis bores a tunnel inside the horehound root, which is gradually filled up with plant material as a fine brown sawdust. When reaching the bottom part of the root, the larva turns and continues feeding upwards, eventually reaching the base of the plant at the end of the larval stage and preparing an exit hole. Feeding damage generally leads to the death of the plant in the following year and the larval tunnel is later invaded by other insects. Integrated control: There is currently no information available on the integration of chemical control of horehound and the clearwing moth.
Horehound plume moth Wheeleria spilodactylus (Curtis) Lepidoptera: Pterophoridae Distribution: Western and Continental Europe, the British Isles, the Mediterranean region, Asia Minor and Northern Africa. First released in Vic. in 1993, it has been widely re-distributed within ACT, NSW, SA, Tas., Vic., and WA. Its range is still spreading. Beneficial status: Major, widespread, regular in all states and territories in south-eastern Australia; minor, restricted, irregular in WA.
Fig. 16.7. Horehound clearwing moth, 12 mm long. (VDPI)
Identification: Adult moths are 10 mm in length with a wingspan of 20 mm. They are pale cream with fine dark marks on the wings. The forewing is divided into two lobes and the hindwing into three feather-like plumes. The moth has a characteristic resting posture with the body and fully outstretched wings forming a T-shape (Fig. 16.8). The eggs, laid singly on the 4 47
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may limit the spread and density of horehound infestations and enable the establishment or reintroduction of more desirable plant species.
Fig. 16.8. Horehound plume moth male, 10 mm long. (VDPI)
undersides of young leaves, are flattened, oval, bright green and 0.5 mm long. The caterpillars are hairy and green with a darker head. They mimic the colour and texture of the horehound leaf. The pupae are well-camouflaged and green to brown. They are generally found on the upper surfaces of lower leaves. May be confused with: This species is unlikely to be confused with any other insect occurring on horehound. Host range: The horehound plume moth is restricted to horehound. Life cycle: The plume moth usually has three generations per year. A female moth lays approximately 100 eggs over a 2-week period, the majority in the first week. Eggs hatch about 1 week after being laid. Larvae begin feeding in the developing shoot tip. When they have grown larger they move out and feed on the leaves, working their way down the shoot, progressively defoliating the stem, until ready to pupate. The first generation of adults emerge in the spring. The second and third generations emerge during the summer and autumn. Each generation takes 1–2 months to complete its life cycle, depending on environmental conditions. Larvae of the autumn generation overwinter in the leaf buds, ready to complete their life cycle when the weather warms up in spring. Active period: Adults are active from October until April–May. Larvae feed in the developing and growing shoot tips over winter. Damage: Larval-feeding severely damages the growing tips of the plant. This weakens the plant and reduces the number of flowers and seeds produced. Large numbers of plume moths
Integrated control: The horehound plume moth appears able to survive autumn herbicide application, provided that live shoots remain on the horehound continuously until spring. Plume moth larvae may further suppress horehound that has been damaged by herbicide treatment and the following generations of moths in spring are likely to at least delay recovery of the horehound population. Where previous experience suggests that herbicide treatment will leave more than 10% of horehound plants alive and green through winter, plume moths can survive at an effective density for the following spring. Survival of horehound will be higher if some plants are sheltered from spraying by stumps, rocks or tall grass or if application is made in less than ideal conditions. If 90% or more of horehound plants are expected to be killed by herbicide, it may be necessary to leave some patches of mature horehound plants unsprayed as reserves for plume moths where they are active. Spraying strips in alternate years may achieve the same effect (Weiss et al. 2000).
Nodding thistle Carduus nutans Asteraceae: Cardueae Nodding thistle was first noticed as naturalised in Australia in 1950 after it had accidentally been introduced as a seed contaminant from New Zealand. The distribution of nodding thistle is currently limited to the high tablelands areas of NSW (populations have been recorded and eradicated from Vic. and WA). It is native to Europe, North Africa, Asia Minor and Asia. It has naturalised and is considered a weed in North and South America, and New Zealand. Three biological control agents are currently established on nodding thistle (Woodburn and Briese 1996).
Nodding thistle receptacle weevil Rhinocyllus conicus (Frolich) Coleoptera: Curculionidae Distribution: A native of the Mediterranean basin and central Europe. First released in NSW
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May be confused with: Unlikely to be confused with any other insects occurring on nodding thistle. Host range: The nodding thistle receptacle weevil is restricted to thistles of the genus Carduus and Cirsium, with different biotypes of the weevil utilising the different genera as hosts.
Fig. 16.9. Adult receptacle weevil on an unopened capitulum of nodding thistle. The body is 10 mm long. (CSIRO Entomology)
in 1988, it has been re-distributed to Vic., and WA, where it was released on slender thistle (Carduus pycnocephalus and Carduus tenuiflorus). Its range covers that of its primary host. Beneficial status: Moderate, widespread, regular in NSW. Identification: Adult weevils are 6–10 mm long with a medium-sized snout (rostrum). When they emerge in spring, they are brown and may be covered in a yellowish coating which is gradually lost with time (Fig. 16.9). Eggs are laid between the bracts (spines) of the thistle head and are covered with a brown faecal cap. Larva and pupae are legless and white, and may be easily observed if an attacked capitulum/flower head is opened up during development (Fig. 16.10). The best time to see this is December–early January.
Life cycle: The nodding thistle receptacle weevil has predominantly one generation per year (a small partial second generation has been observed in a good season). Adults become active in spring just before the thistle rosettes produce flowering stems. They chew holes in the leaves and stems of thistle plants and feed on pollen in their flowers. Females lay their eggs between bracts of the thistle head from late spring to early summer. Eggs are laid individually in a small hole chewed into the surface by the female and are then covered with a faecal deposit. Each female may lay up to 100 eggs. Females select green bud stage heads for oviposition. Newly hatched larvae bore directly into the head and begin to feed on the receptacle tissue, destroying developing seeds. Following pupation in the head, adults emerge in late January–February and seek protected sites where they remain dormant until the following spring. Active period: Adults are active from November until January. Larvae feed in the thistle head from December to January. Damage: Adults chew shot-holes in the thistle leaves causing minor damage. Larvae feed in the receptacle of the thistle head, destroying the developing seed. Each larva can prevent the production of 30 seeds and at peak attack times larvae can destroy all of the developing seed in a capitulum. Unfortunately the peak attack period (December–January) in Australia falls outside of period of peak seed production (January– February). This results in the receptacle weevil reducing seed production by between approximately 15 and 35% annually (Woodburn and Cullen 1996).
Nodding thistle rosette weevil Fig. 16.10. Receptacle weevil pupae and damage caused to developing seed of nodding thistle by larval feeding. (CSIRO Entomology)
Trichosirocalus mortadelo (Panzer) AlonsoZarazaga & Sanchez-Ruiz Coleoptera: Curculionidae 449
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emerged, they feed on the flowering plants for a short period before returning to the soil/litter where they remain dormant until autumn. Active period: Adults are active from March until September with the new generation emerging in December. Larvae feed in the thistle rosette from March until October.
Fig. 16.11. Adult nodding thistle rosette weevil, 4 mm long. (CSIRO Entomology)
Distribution: A native of the Mediterranean basin and central Europe. First released in NSW in 1993. It has also been released onto Cirsium vulgare in Vic. and WA, but failed to establish. Beneficial status: Major, widespread, regular in NSW. Identification: Adult weevils are brown and 3– 4 mm long (Fig. 16.11). Eggs are laid primarily into the underside of rosette leaves between March and September. The larva is legless with a brown head and may be easily observed in the crown of a damaged rosette. To find larvae, look for soft black necrosis in the centre of the rosette as a sign of attack, then simply scratch around in the damaged area to see larvae. May be confused with: Unlikely to be confused with any other insects occurring on nodding thistle. Host range: The nodding thistle rosette weevil has been observed attacking thistles in the genus Carduus and Cirsium, although populations of the weevil only persist on nodding thistle. Life cycle: The nodding thistle rosette weevil has one generation per year. Adults become active in autumn and females lay several hundred eggs into rosette leaves from autumn until early spring. Once hatched, the larvae mine within the petiole towards the crown (centre) of the rosette where they feed until development is complete, at which time they leave the rosette to pupate in the soil. While oviposition and larval-feeding is staggered over several months, emergence from pupation is a relatively synchronous event, with the new generation of adults emerging form the soil during December. Once new adults have
Damage: Adults chew shot-holes in the thistle leaves causing minor damage. Larvae feed in the crown of the rosette damaging new leaves as they develop, under high attack rates they may destroy the meristem. Meristem damage can lead to the formation of multiple crowns that may also be attacked; when secondary meristem growth is attacked rosette death may result. Another effect of heavy larval attack is that rosettes lose their spines, becoming more dandelion-like, making them more accessible to grazing animals. The rosette weevil reduces seed production indirectly in nodding thistle through reducing plant size or killing plants before the onset of flowering. Field experiments have quantified this effect to reduce seed rain by up to 72% in a single year (Woodburn 1997).
Nodding thistle seed fly Urophora solstitialis Linnaeus Diptera: Tephritidae Distribution: A native of the Mediterranean basin and central Europe. First released in NSW in 1991. Beneficial status: Major, widespread, regular in NSW. Identification: Adult flies are small (5–7 mm long) with yellow and black markings. Females have a long ovipositor protruding from their abdomen (Fig. 16.12). Larvae are white and feed within a gall that is produced as a result of their feeding, which forms in the receptacle of the capitulum/flower head. May be confused with: Unlikely to be confused with any other insects occurring on nodding thistles. Host range: The nodding thistle seed fly is restricted to nodding thistle. Life cycle: The nodding thistle seed fly has one full (December–early January) and a second
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Fig. 16.12. Female nodding thistle seed fly. The body is 7 mm long. (CSIRO Entomology)
partial generation per year (late January– February). The female lays up to 100 eggs which she lays individually into a floret (each thistle ‘flower’ head is made up of hundreds of florets), while the flower head is at the ‘green bud’ stage. Once hatched, the larvae feed through the embryonic seed into the receptacle where it’s feeding induces the plant to form a woody gall around the larva. It is common to find large woody galls containing multiple larvae within a single flower head. Larvae overwinter inside the gall before pupating and emerging as adults in the late spring to start the cycle over again. Active period: Adults from the two generations are active from late November until February. Damage: The adult fly causes no damage to the thistle. An individual larva feeding within the gall causes the failure of seven seeds to set. Across a flowering season, the seed fly can reduce seed set by up to 70% (Woodburn 1996, Woodburn and Cullen 1996).
Paterson’s curse
been introduced to North and South America, Australia, South Africa and New Zealand. All species are weeds in NSW, SA, Vic., WA and southern Qld. Paterson’s curse is the most widespread and is the dominant pasture weed in winter rainfall regions of Australia, particularly in NSW. Vipers and Italian bugloss are native to Mediterranean and Continental Europe and have been introduced to South Africa, Australia and New Zealand. In Australia, viper’s bugloss is a common plant in tableland country in the eastern states, while Italian bugloss is rare and has been recorded in NSW and SA.
Paterson’s curse crown weevil Mogulones larvatus Goeze Coleoptera: Curculionidae Distribution: A native of the Mediterranean basin and Continental Europe. First released in NSW in 1992, it has been re-distributed to SA, Vic. and WA. Its range is still spreading. Beneficial status: Major, localised, regular in NSW, Vic. and SA; minor, restricted, irregular in other states. Identification: Adult weevils are small (3–4 mm long) with a medium-sized rostrum. When they emerge in spring, they feed on the foliage and flowers of Paterson’s curse (Fig. 16.13) until early summer when they are dormant. Weevils become active in autumn and females lay small (0.8 × 0.5 mm) yellow eggs into the leaf stalk of Paterson’s curse. The mature larva is white, legless with a brown head and is 10 mm long by 3–4 mm wide. It feeds in the leaf stalk and crown of the rosette.
Echium plantagineum
Viper’s bugloss E. vulgare
Italian bugloss E. italicum Boraginaceae: Boraginoideae Paterson’s curse, originally occurring widely in Mediterranean Europe and North Africa, has
Fig. 16.13. Adult crown weevils on an opened flower of Paterson’s curse (body is 3.5 mm long). (CSIRO Entomology)
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May be confused with: Unlikely to be confused with any other insects occurring on Paterson’s curse. Host range: The Paterson’s curse crown weevil is restricted to herbaceous plants in the genus Echium, which also includes the weeds Viper’s bugloss and Italian bugloss. Life cycle: The crown weevil has a single generation each year. Adult weevils become active from February to April after heavy rain stimulates Paterson’s curse germination. After feeding for a week on rosette leaves, females become sexually mature and lay eggs into the leaf stalk of the rosette. Eggs are laid from February–November with females producing 550 eggs on average. The weevil larvae hatch and feed within the leaf stalk. As larvae mature, they mine towards the crown (the growth point of the rosette) and the top of the taproot. After feeding, larvae leave the rosette to pupate in the soil. The adults emerge in spring, feeding on the leaves and flowers of Paterson’s curse until the plant is dead or until late December. During this time, adults disperse to seek out new sources of Paterson’s curse. After feeding, adults move into the soil and leaf litter and become dormant to escape the summer heat. The adults do not feed during summer, living off fat reserves accumulated during spring. Only the healthiest adults survive. Adult weevils remain dormant in the soil until autumn rain stimulates them to become active and start a new generation. The early activity of the crown weevil will limit its ability to survive in regions where late breaks to the season are common. Active period: Adults are active from late February until November with the new generation of adults emerging from October and feeding until December. Larvae feed in the leaf stalk and crown of the rosette from March to December. Damage: Adults chew shot-holes in the leaves. Larvae feed in the crown of the rosette and 10– 20 larvae can kill a rosette of 15–20 cm diameter before flowering (Fig. 16.14). Plant mortalities of greater than 80% have been observed at some sites with high weevil densities. Integrated control: Use of herbicides does not kill adults but larvae will starve as their food source has been destroyed. The use of
Fig. 16.14. Damage to a Paterson’s curse rosette caused by larval crown weevil feeding. (CSIRO Entomology)
insecticides will kill all stages of the insect directly. If insecticides need to be used for redlegged earth mite, application of insecticide using TIMERITE® will minimise mortality, as the crown weevil will be pupating below ground (the use of a non-systemic insecticide will further limit mortality). Grazing pressure that removes Paterson’s curse foliage and particularly the crown will severely limit the activity of the crown weevil.
Paterson’s curse root weevil Mogulones geograghicus Goeze Coleoptera: Curculionidae Distribution: A native of the Mediterranean basin and Continental Europe. First released in NSW in 1994, it has been re-distributed to SA, Vic. and WA. It is still spreading. Beneficial status: Major, restricted, in NSW, SA, Vic. and WA. Identification: Adult weevils are small (3.5– 4.5 mm long) with a medium-sized rostrum (Fig. 16.15). When they emerge in spring, they feed on the foliage and flowers of Paterson’s curse until early summer when they go dormant. Weevils become active in autumn and females lay small white (0.9 mm × 0.4 mm) eggs into the leaf stalk of Paterson’s curse. The mature larva is white, legless with a brown head and is 10–12 mm long by 3–4 mm wide and feeds in the tap-root of the rosette. May be confused with: Unlikely to be confused with any other insects occurring on Paterson’s curse. Host range: The Paterson’s curse root weevil is restricted to herbaceous plants in the genus
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Fig. 16.15. Adult root weevil on an opened flower of Paterson’s curse. The body is 4.5 mm long. (CSIRO Entomology)
Fig. 16.16. Damage to a Paterson’s curse root caused by larval-feeding. The body of the larva is 8 mm long. (CSIRO Entomology)
Echium, which also includes the weeds Viper’s bugloss and Italian bugloss. Life cycle: The root weevil has a single generation each year. Adult weevils become active from March to May after heavy rain stimulates Paterson’s curse germination. After feeding for a week on rosette leaves, females become sexually mature and lay eggs into the leaf stalk of the rosette. Eggs are laid from March to October with females producing 250 eggs on average. The weevil larvae hatch and feed within the leaf stalk. As larvae mature they mine towards the tap-root. After feeding, larvae leave the rosette to pupate in the soil. The adults emerge in spring, feeding on the leaves and flowers of Paterson’s curse until the plant is dead or until late December. During this time adults disperse to seek out new sources of Paterson’s curse. After feeding, adults move into the soil and leaf litter and become dormant to escape the summer heat. Adults do not feed during summer, living off fat reserves accumulated during spring. Only the healthiest adults survive. Adults remain dormant in the soil until autumn rain stimulates them to become active and start a new generation. Feeding belowground, the root weevil will tolerate grazing better than the closely related crown weevil. Active period: Adults are active from March until October with the new generation of adults emerging from October and feeding until December. Larvae feed in the tap-root of the rosette and in the flowering plant from March to December. Damage: Adults chew shot-holes in the leaves. Larvae feed in the tap-root of the rosette, where
20–30 larvae can kill a rosette of 15–20 cm diameter before flowering (Fig. 16.16). Plant mortalities of greater than 80% have been observed during the laboratory breeding of this insect. Integrated control: The use of herbicides does not kill adults but larvae will starve as their food source has been destroyed. The use in insecticides will kill all stages of the insect directly. If insecticides need to be used for redlegged earth mite, application of insecticide using TIMERITE® will minimise mortality, as the root weevil will be larvae and pupae belowground (the use of a non-systemic insecticide will further limit mortality). Grazing pressure that removes large amounts Paterson’s curse foliage and particularly the crown will slow the activity of the root weevil.
Paterson’s curse flea beetle Longitarsus echii Koch Coleoptera: Chrysomelidae Distribution: A native of the Mediterranean basin and Continental Europe. First released in NSW in 1996, it has been re-distributed to SA, Vic. and WA. It is still spreading. Beneficial status: Major, restricted in NSW, SA, Vic. and WA. Identification: Adult beetles are small (3–4 mm long) and are a shiny metallic black in colour (Fig. 16.17). The adults have large hind legs that they use to jump a metre or more when disturbed. When adult beetles emerge from the 453
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Fig. 16.17. Adult flea beetle on a leaf of Paterson’s curse. The body is 3.5 mm long. (CSIRO Entomology)
ground in winter, they feed on the foliage of Paterson’s curse and females lay small (0.9 mm × 0.3 mm) orange–red eggs onto the tap-root of Paterson’s curse. The mature larva is white, with three pairs of small legs near the black head and is 10 mm long by 1–2 mm wide. The larva feeds in the tap-root and secondary roots of the rosette and flowering plant. May be confused with: Unlikely to be confused with any other insects occurring on Paterson’s curse. Host range: The Paterson’s curse flea beetle is restricted to herbaceous plants in the genus Echium, which also includes the weeds Viper’s bugloss and Italian bugloss. Life cycle: The flea beetle has a single generation each year. Adult beetles become active from May to August. After feeding for a week on rosette leaves, females become sexually mature and lay eggs onto the taproot of the rosette. Eggs are laid from May to November with females producing 250 eggs on average. The beetle larvae hatch and feed within the taproot. After feeding, larvae leave the root to pupate in the soil and remain there belowground over summer. Adult beetles remain dormant in the soil until winter when they become active and start a new generation. Feeding belowground and emerging late in the season, the flea beetle will tolerate heavy grazing and dry autumns better than the other Paterson’s curse insects (Ireson et al. 1991). Active period: Adults are active from May until November. Larvae feed in the taproot of the rosette from May to December. Damage: Adults chew shot-holes in the leaves. Larvae feed in the taproot of the rosette, where 30–40 larvae can kill a rosette of 15–20 cm
Fig. 16.18. Damage to a Paterson’s curse root caused by flea beetle larval-feeding. The body is 10 mm long. (CSIRO Entomology)
diameter before flowering (Fig. 16.18). Plant mortalities of greater than 40% have been observed in the field under continuous grazing and > 80% have been observed during the breeding of this insect. Integrated control: The use of herbicides does not kill adults but larvae will starve as their food source has been destroyed. The use in insecticides will kill all stages of the insect directly. If insecticides need to be used for redlegged earth mite, application of an insecticide using TIMERITE® will minimise mortality, as the flea beetle will predominantly be larvae belowground (the use of a nonsystemic insecticide will further limit mortality). Grazing pressure that removes large amounts Paterson’s curse foliage even if the crown is grazed has minimal impact on the activity of the flea beetle.
Paterson’s curse pollen beetle Meligethes planiusculus Koch Coleoptera: Nitidulidae Distribution: A native of the Mediterranean basin and Continental Europe. First released in NSW in 1996, it has been re-distributed to SA, Vic. and WA. It is still spreading. Beneficial status: Moderate, restricted, in NSW, SA, Vic. and WA. Identification: Adult beetles are small (2–2.5 mm long), black (Fig. 16.19) and become active in early spring when Paterson’s curse rosettes start to bolt and flower. After a week of feeding, females become sexually mature and lay on average 130 eggs between the unopened flower buds of
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Fig. 16.19. Adult pollen beetle on an opened flower of Paterson’s curse. The body is 2 mm long. (CSIRO Entomology)
Fig. 16.20. A pollen beetle larva feeding in a Paterson’s flower. The body is 2 mm long. (CSIRO Entomology)
Paterson’s curse. Larvae hatch from the eggs and mine into the flower bud where they feed on the pollen and ovules (the female part of the flower). Larvae then move between flowers feeding on immature green seed. When larval feeding is complete, larvae pupate in the soil and after 1–2 weeks new adults emerge from the soil and feed on the buds and immature seed until the end of flowering. At the end of flowering, adults enter the soil and become dormant to escape the heat of summer until autumn rain stimulates germination. Adults then emerge from the soil and feed on the young rosette leaves before entering winter hibernation until spring.
is complete, larvae drop from the flowering plant and pupate in the soil. After 1–2 weeks, new adults emerge from the soil and feed on the buds and immature green seed until the end of flowering. This life cycle enables the pollen beetle to attack the first flowers through to the last seed produced by Paterson’s curse. At the end of flowering, adults enter the soil and become dormant to escape the heat of summer until autumn rain stimulates germination. Adults then emerge from the soil and feed on the young rosette leaves before entering winter hibernation until spring.
May be confused with: Unlikely to be confused with any other insects occurring on Paterson’s curse. Host range: The Paterson’s curse pollen beetle is restricted to herbaceous plants in the genus Echium, which also includes the weeds Viper’s bugloss and Italian bugloss. Life cycle: Pollen beetle adults become active in early spring when Paterson’s curse rosettes start to bolt and flower. Adults can be seen congregating on the first flowering plants in a paddock and feeding on the unopened flower buds. After a week of feeding, females become sexually mature and lay on average 130 eggs between the unopened buds of Paterson’s curse. Larvae hatch from the eggs and mine into the flower bud where they feed on the pollen and ovules (the female part of the flower), destroying the bud and preventing it from producing seed. Larvae then move between flowers, feeding on immature green seed preventing it from forming viable hard seed. When larval-feeding
Active period: Adults are active from late August until December. Larvae feed in the unopened buds, flowers and on the green seed of Paterson’s curse. Damage: Adults chew holes in the flower buds and in green seed of Paterson’s curse. Larvae feed in the flower buds, flowers (Fig. 16.21) and on the green seed of Paterson’s curse. Seed production has been reduced by 60% under experimental conditions in Europe. Integrated control: The use of herbicides does not kill adults but larvae will starve as their food source has been destroyed. The use in insecticides will kill all stages of the insect directly. If insecticides need to be used for redlegged earth mite, even the application of insecticide using TIMERITE® will kill the pollen beetle, as it is active at this time of year. Leaving areas of Paterson’s curse unsprayed each year (if possible) will ensure the survival of the beetle into the future. Grazing pressure that removes large amounts of flowering Paterson’s curse will limit the activity of the pollen beetle. 455
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Paterson’s curse leafminer Dialectica scalariella Zeller Lepidoptera: Gracillariidae Distribution: A native of the Mediterranean basin and Continental Europe. First released in Vic. in 1988, it has been re-distributed to NSW, SA and WA. Its can now be found across the range of Paterson’s curse. Beneficial status: Major, widespread, regular in NSW, SA and Vic.; minor, restricted, irregular in other states. Identification: Adult moths are small (8–10 mm long by 1 mm wide) with brown and white markings (Fig. 16.21). They are active all year except for the coldest winter months. Females lay small clear (0.3 mm × 0.3 mm) eggs into the leaves of Paterson’s curse. The caterpillar feeding within the leaf has a brown head and white body and is 10–12 mm long by 1.5 mm wide. The feeding of the caterpillar causes the leaf to blister, and when a leaf like this is opened several caterpillars can usually be found feeding. May be confused with: Unlikely to be confused with any other insects occurring on Paterson’s curse. Host range: The Paterson’s curse leafminer is restricted to herbaceous plants in the genus Echium, which also includes the weeds Viper’s bugloss and Italian bugloss. Life cycle: The leaf miner has five to seven generations each year. Females lay on average 120 eggs on the leaf surface; caterpillars hatch and mine into the leaf, feeding on the green plant material. Caterpillars pupate within the
mine inside a white silken cocoon. In summer, the leafminer can develop from egg to adult in less than a month and populations can build up rapidly if there is green plant material for them to feed on. Active period: Moths and caterpillars can be found throughout the year as long as there is green plant material for them to feed on. The activity of the moth is greatly reduced in winter. Damage: Caterpillars feed inside the leaves reducing leaf area for photosynthesis (Fig. 16.23). The leafminer is much more damaging on the perennial Echium weeds (Viper’s and Italian bugloss) as it provides green feed for them through the warm summer months when the moth is most active. Integrated control: The use of herbicides does not kill adults but larvae will starve as their food source has been destroyed. The use of insecticides will kill all stages of the insect directly. If insecticides need to be used for redlegged earth mite, application of an insecticide using TIMERITE® will minimise mortality, as the leafminer is most active in summer–autumn. Grazing pressure that removes large amounts Paterson’s curse foliage particularly if the crown is grazed will limit the activity of the leafminer.
Ragwort Senecio jacobaea Asteraceae: Senecioneae Ragwort is a native of Europe and western Asia. It has become a serious agricultural and environmental weed on the northwest coast of the United States, in the maritime provinces of Canada and in New Zealand, Argentina and Australia. In Australia, it has become widely established in the high rainfall regions of Tas. and Vic. Three biological control agents now established on ragwort in Australia are expected to have an impact.
Ragwort flea beetle Longitarsus flavicornis (Stephens) Coleoptera: Chrysomelidae Fig. 16.21. Paterson’s curse leafminer moth in its characteristic ‘alert’ position. Body is 8 mm long. (CSIRO Entomology)
Distribution: A native of Europe, the ragwort flea beetle was first released in Tas. and Vic. in
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1979. It has now spread throughout most of the ragwort infestations in Tas. and has only established at high altitude, high rainfall locations in Vic. Beneficial status: Major, widespread, regular in Tas.; minor and restricted in Vic. Identification: The beetle (Fig. 16.22) is about 3 mm long. Newly emerged beetles are a light brown colour, becoming darker as they mature. The eggs are elongated, about 0.2 mm in diameter and 0.6 mm long and light brown in colour. Larvae are whitish in colour and pass through three moults or instars. Newly hatched larvae are about 1.5 mm in length and have dark heads. Mature larvae are about 5 mm in length and have a reddish-brown head. The pupae from which adults emerge are white and just over 3 mm in length. May be confused with: In Tas., adults of beetles belonging to the genus Haltica (metallic flea beetles) occasionally feed on ragwort foliage in large numbers and are mistaken for the ragwort flea beetle. They are about the same size and shape, but the iridescent blue colour of the Haltica species easily distinguishes it from the brown ragwort flea beetle. Host range: In Australia, the ragwort flea beetle feeds only on ragwort and has no other host. Life cycle: One generation of the ragwort flea beetle is produced each year. The beetles start to emerge from pupae in the soil about midDecember (sometimes earlier) and eggs are laid, usually in the surface soil around the plants, 3–4
Fig. 16.23. Damage to the lower root crown caused by larvae of ragwort flea beetle. (TIAR)
weeks after mating. Hatching is temperaturedependent and occurs in about 17 days at 20°C. The newly hatched larvae feed by mining into the petioles, roots and root crown. Larvae are also commonly found feeding externally on roots in the soil around the plants. Larvae feed from late summer until the end of the following spring when they pupate in the soil around the plants. Active period: Much of the beetles’ life cycle takes place belowground and their presence is often not evident to the casual observer; however, during summer and autumn, the adult beetles are active and can be observed feeding on the rosettes, particularly on warm, still and overcast days. Damage: The action of the larvae burrowing into roots and root crowns (Fig. 16.23) causes the most significant damage to ragwort. As many as 400 larvae may feed on a single plant, severely weakening the plant and ultimately causing its death. Although the feeding of the adult stage (beetle) on the rosettes is not regarded as having a significant detrimental effect on the plants, large numbers of beetles have caused extensive leaf damage at some sites with only ragged portions of the leaves remaining on some plants. Integrated control: The ragwort flea beetle can be successfully integrated into a management plan using sheep-grazing, mowing and wickwiped herbicides.
Ragwort plume moth Fig. 16.22. The ragwort flea beetle. The body is 3 mm long. (TIAR)
Platyptilia isodactyla (Zeller) Lepidoptera: Pterophoridae 457
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Distribution: A native to central Europe and the Mediterranean, it was first released in Tas. and Vic. during 1999–2000. Beneficial status: Continuing to spread in Tas. and Vic. As yet, no studies have been conducted on its impact. Identification: The moths (Fig. 16.24) are about 9 mm in length with a wingspan of 21 mm. The wings are pale fawn or brown in colour with dark bands at the ends and variable V or double V-shaped dark brown bands approximately one third of the wing length from the wing tip. The body and legs are lighter brown. The forewing is divided into two lobes and the hindwing into three feather-like plumes. Eggs are flat, ovoid and smooth, 1 mm long and 0.5 mm wide. They are bright shiny green when first laid but gradually turn a pale yellow. Larvae have a black head. Newly hatched larvae are ivory or pale yellow. Larger larvae are pale green, with small dark spots, and sparsely hairy. Larvae become darker green as they develop further. Mature larvae are around 20 mm long. Pupae are light to dark brown, 18–19 mm long and 5–6 mm wide. May be confused with: Damage caused by the ragwort plume moth could be confused with that caused by the blue stemborer and the ragwort stem and crownboring moth. The main distinguishing features of these species are discussed under this heading for the ragwort stem and crownboring moth. Host range: The moth is restricted to species within the Senecionae tribe. In Australia, ragwort is the moth’s preferred host as it has been shown to have very limited survival
Fig. 16.25. Damage to the upper root crown caused by larvae of the ragwort plume moth. The photo shows mature larva together with a newly formed pupa below. (TIAR)
capabilities on any other Senecio species. Native species are not considered at risk. Life cycle: The plume moth has two generations per year. Moths fly in spring and autumn. At 20°C, females lay an average of 100 eggs during a lifetime of about 12 days. Newly hatched larvae burrow into the leaf petioles until they reach the crown. The larvae pass through five stages (instars). Older larvae tunnel in the crown, stem and roots. Larvae eject their frass (faecal deposits) and shed heads from a small hole in the stem and this debris accumulates on silken webbing spun around the hole by the larva. The pupal stage lasts approximately 1 week. Damage: The amount of damage this agent will inflict on ragwort under field conditions will need to be determined by future research; however, larval-feeding has the potential to severely damage the stem and crown of the plant (Fig. 16.25), reducing vigour and the number of flowers and seeds produced. Integrated control: Field studies are now being planned to determine a more effective integrated control system for ragwort using combinations of the biological control agents with traditional farm management practices (grazing, mowing and herbicides).
Ragwort stem and crownboring moth Fig. 16.24. The ragwort plume moth. Body is 9 mm long. (VDPI)
Cochylis atricapitana (Stephens) Lepidoptera: Cochylidae
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Distribution: A native of Europe, most abundant in Spain, France and Portugal. It was first released in Vic. in 1987 and Tas. in 1995. Beneficial status: Continuing to spread in Vic. and Tas., but its full impact is yet to be determined. Identification: The moth is 8–10 mm long and pale to dark brown in colour. It has a 1 mm wide dark blotchy band running diagonally across the middle and ends of the forewing. Moths are most active at dusk or dawn. Eggs are flat and oval in shape, 0.5 mm long and a translucent white in colour when first laid. They become yellow during development. The larvae are creamy white in colour. The young larvae have a black head, which becomes light brown or tan as it matures. Pupae are light brown, 7–8 mm long and 1.5–2 mm wide. May be confused with: The damage can be confused with that caused by larvae of a native insect that often attacks ragwort, the blue stemborer, Patagoniodes farinaria (Turner). The larva of the blue stem borer has bluish-grey stripes along its length, three of which are visible when viewed from above. The middle dorsal stripe is narrower than the other two. Younger larvae have the same stripes but they tend to be a lighter, brownish colour. The striping of these caterpillars easily distinguishes them from the uniform creamy white colour of the ragwort stem and crownboring moth. In addition, the excrement (frass) produced by the larvae of the blue stemborer is coarse and easily visible, hanging off the stems of ragwort. Although the ragwort stem and crownboring moth attacks similar parts of the plant, the frass produced is much finer and less obvious.
these moths feeding until autumn (May). Towards the end of autumn and the beginning of winter, mature larvae enter a state of dormancy until the following spring. Before entering dormancy, larvae spin silken cocoons and change from creamy white to pink in colour. Most larvae overwinter in this dormant phase in the crowns of rosettes or under the bark at the base of senescent stems. They have also been found in the leaf buds of senescent plants and among dead ragwort leaves and surrounding litter. With the increasing day lengths and higher spring temperatures, the larvae become active again and pupate (McLaren 1992). Active period: Moths may be present from spring until the onset of autumn, due to generation overlap. They conceal themselves under leaves during the day and begin to fly at dusk. Larvae can be found feeding actively on ragwort from November to May. Damage: After hatching, the larvae begin mining into the plant, boring into the leaves, stem or crown (Fig. 16.26). Attacked ragwort rosettes may be distinguished by blemished or blackened young central shoots and the wilted appearance of side shoots, caused by larval damage to the crown. In severely infested rosettes, larvae eat out the central crown and then start boring up leaf petioles, killing the plant. Some plants survive and re-grow, but have reduced foliage and do not flower that season. Bolting ragwort plants attacked in the stem can be identified by small tunnels entering at the leaf bud in the axils of side shoots. Attacked flowering plants can be identified by the
Host range: In Europe it is restricted to some close relatives within the genus Senecio, but only occurs on ragwort in Australia. Life cycle: There are at least two complete generations of the stem and crownboring moth annually. Moths of the first generation start to emerge in early spring with maximum emergence about October. The females lay up to 150 eggs. Most first generation larvae feed and develop on the ragwort plant from November with most pupating by mid-summer (January). The second generation of moths start to emerge during January with the larvae produced by
Fig. 16.26. Damage to the lower stem and upper crown caused by larvae of the ragwort stem and crown boring moth. (TIAR)
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blackened, wilted appearance of the flower buds and/or the multi-stemmed flower crown, which branches out from below the point where the flower stalk was affected. Heavy larval infestations on bolting ragwort plants may prevent or reduce flowering and subsequent seed production (McLaren et al. 2000). Integrated control: Field studies are now being planned to determine a more effective integrated control system for ragwort using combinations of the biological control agents with traditional farm management practices (grazing, mowing and herbicides).
a
b
Skeleton weed Chondrilla juncea Asteraceae Originally from west Asia and the Mediterranean areas of Europe and North Africa, it has spread to central Europe, North and South America, New Zealand and Australia. In mainland Australia, it is an important weed of winter rainfall cropping areas, particularly where soils are sandy. Here, it is a long-lived perennial and spreads by seed, from upgrowth of lateral roots and from roots fragmented by cultivation. Australia has three biotypes of the weed: a narrow-leafed form, which originally was the most common form, a broadleafed form, and an intermediate form of skeleton weed. A number of biological control agents are established in Australia, the most effective of which is a rust fungus, Puccinia chondrillina (Fig. 16.27) that controls the narrow-leafed form of skeleton weed in south-eastern Australia. Two arthropod agents supplement control by the rust fungus (Burdon et al. 1981).
Fig. 16.27. The rust Puccinia chondrillina is effective in controlling the narrow-leafed form of skeleton weed in eastern Australia. Damage symptoms may be seen on (a) leaf and (b) flower. (CSIRO Entomology: J. Green)
Identification: The mites are minute and barely visible to the naked eye. They are best identified by their galls of 1.5–2 cm (but may be as large as 5 cm) on vegetative and flower buds of the plant (Fig. 16.28). May be confused with: Chondrilla gall midge also produces galls but these are confined to leaves and are generally smaller than gall mite. Host range: Skeleton weed is the only known host.
Chondrilla gall mite Aceria chondrillae (Canestrini) Acarina: Eriophyidae Distribution: Cultures were imported from southern Europe in 1971 and released in southern Australian cropping areas during the following decade. The mite has established in south-eastern Australia. Beneficial status: Major and regular on narrowleafed skeleton weed in south-eastern Australia, minor on other forms of the weed.
Fig. 16.28. Galls on skeleton weed stems caused by Chondrilla gall mite. (CSIRO Entomology: J. Cullen and J. Green)
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Fig. 16.31. Stems with gall midge lesions. (CSIRO Entomology)
Fig. 16.29. Adult gall midge, 1–1.5 mm long. (CSIRO Entomology)
Life cycle: In spring, mites feed on flower and vegetative buds, causing them to form galls which shelter and nourish the mites. Mites reproduce within the gall and migrate to other parts of the plant or are blown to new plants. The mites overwinter on skeleton weed rosettes. Damage: Large numbers of galls on vegetative parts of skeleton weed reduce its viability and galls on floral parts reduce seed production. Damage is greatest on the narrow-leafed form of skeleton weed. Integrated control: Other biological control agents or current cultural practices in cereals do not appear to interfere with the action of chondrilla gall mite.
Chondrilla gall midge Cystiphora schmidti Rübsaamen Diptera: Cecidomyiidae Distribution: Southern Europe and west Asia. Cultures were imported from southern Europe.
In Australia, it was released at the same time as the gall mite and established in parts of southeastern Australia. Host range: Skeleton weed and related species in the genus Chondrilla. Beneficial status: This is the least effective of the biological control agents established in Australia, due in part to its patchy establishment and its parasitism by a native parasitoid, Tetrastichus sp. (Hymenoptera: Eulophidae). It appears to contribute to the effects of skeleton weed rust and gall mite. It attacks all forms of skeleton weed. Life cycle: The adult female midge lays eggs into the underside of newly growing leaves and on stems. The plant reacts by producing leaf galls (Fig. 16.29). Larval stages and pupa are contained within the gall. A generation takes 3–6 weeks during warm periods. The midge overwinters as larvae. Damage: Galls contribute to reducing the vigour of plants (Figs 16.30 and 16.31).
Scotch thistle Onopordum acanthium
Illyrian thistle O. illyricum
Stemless thistle O. acaulon Asteraceae: Cardueae Fig. 16.30. Skeleton weed rosette showing purplish leaf blotches caused by skeleton weed gall midge. (CSIRO Entomology)
Scotch thistle originally occurs widely in Europe and has been introduced to North and South America, Australia and New Zealand. Illyrian 4 61
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thistle is native to the Mediterranean basin and has been introduced to North America and Australia. Both species are weeds in NSW and Vic., while only Scotch thistle occurs in Tas. Hybrids of the two species also occur widely in NSW. Stemless thistle originates from the Iberian peninsula of Europe and was introduced to Australia where it occurs in NSW, SA, Vic., and WA. Four biological control agents are currently established on Onopordum spp. thistles (Briese et al. 2002a).
Onopordum capitulum weevil Larinus latus Linnaeus Coleoptera: Curculionidae Distribution: A native of the eastern Mediterranean basin and western central Asia. First released in NSW in 1992, it has been redistributed to Vic., SA and WA. Its range is still spreading. Beneficial status: Major, widespread, regular in NSW; minor, restricted, irregular in other states. Identification: Adult weevils are large (14–24 mm long), oval-shaped weevils with a medium-sized rostrum (Fig. 16.32). When they emerge in spring, they are dark brown and covered in a yellowish waxy coat which is gradually lost with time until they appear blackish by late summer. The creamy white eggs are covered with hard faecal material, which may be brown if laid on the outside of the capitulum or purplish if laid directly into the florets. The creamy white larva is legless with a brown head.
Fig. 16.32. Adult onopordum capitulum weevil on an unopened capitulum of Illyrian thistle. The body is 20 mm long. (CSIRO Entomology)
May be confused with: Unlikely to be confused with any other insects occurring on Onopordum thistles. Host range: The onopordum capitulum weevil is restricted to thistles of the genus Onopordum, which include Scotch and Illyrian thistle and their hybrids, and stemless thistle in Australia (Briese et al. 2002b). Life cycle: The onopordum capitulum weevil has one generation per year. Adults become active in spring just before the thistle rosettes produce flowering stems. They chew holes in the leaves and stems of thistle plants and feed on pollen in their flowers. Females lay their eggs on the underside, between bracts or directly into the florets of thistle capitula from late spring to early summer. Eggs are laid individually in a small hole chewed into the surface by the female and are then covered with a faecal deposit. Each female may lay up to 70 eggs. Oviposition occurs from the green bud stage until the flowers have dried. Newly hatched larvae bore directly into the capitulum and begin to feed on the receptacle tissue, destroying developing seeds. Following pupation in the capitula, adults emerge in late summer and seek protected sites where they remain dormant until the following spring. Active period: Adults are active from October until January. Larvae feed in the capitula of thistles from December to January. Damage: Adults chew shot-holes in the thistle leaves. Larvae feed in the base of the thistle capitulum, destroying the developing seed. A
Fig. 16.33. Damage to developing seed of Scotch thistle caused by larvae of onopordum capitulum weevil. (CSIRO Entomology)
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single larva can destroy all seed in a capitulum of 25 mm diameter or less, and several larvae may co-exist in larger capitula (Fig. 16.33). Over 90% reduction in seed rain has been observed at some sites with high weevil densities.
Onopordum crown weevil Trichosirocalus briesei Alonso-Zarazaga & Sanchez-Ruiz Coleoptera: Curculionidae Distribution: A native of the Mediterranean basin. First released in NSW in 1997. This agent is established at only a few sites, but is currently being re-distributed. Beneficial status: Minor, restricted, irregular. Identification: Adult weevils are mottled brown and rounded, 4–5 mm long with a mediumsized curved snout (rostrum) (Fig. 16.34). The creamy white eggs are laid into pockets that have been bored into the leaf petiole. The larva is white, legless and has a brown head. May be confused with: Unlikely to be confused with any other insects occurring on Onopordum thistles. Host range: The onopordum crown weevil is restricted to thistles of the genus Onopordum, which includes Scotch and Illyrian thistle and their hybrids, and stemless thistle in Australia (Briese et al. 2002b). Life cycle: The onopordum crown weevil has one generation per year. Adults become active in autumn, and lay eggs from autumn to spring in pockets in the midrib on the under-sides of thistle rosette leaves. Each pocket may contain up to four eggs and females may lay up to 700 eggs over this period. Upon hatching, the larvae
Fig. 16.35. Damage to Scotch thistles caused by onopordum crown weevil larval feeding on rosette meristem tissue. (CSIRO Entomology)
mine down through the midrib to the crown of the plant, where they feed in the base of the petiole and on meristem tissue. There are three larval instars. Mature third instars migrate from the crown into the soil, where they pupate inside earthen cells. The adults emerge in late spring, feed on Onopordum foliage and then enter a dormant period in the soil or litter until the following autumn. Active period: The main period of damage is from April to August, when mating and egglaying occur and larvae feed in the crowns of Onopordum thistle rosettes. Larval presence can first be detected from the presence of brown to black scarring along the midrib of a leaf. Damage: Adults chew small shot-holes in the thistle leaves. Larvae feed within the petiole of rosette leaves and can mine into and feed on meristem tissue within the rosette crown (Fig. 16.35). This can lead to black necrotic lesions within the rosette crown, which can reduce the subsequent size of bolting stems and even kill the plant at high larval densities.
Onopordum stemboring weevil Lixus cardui Ol. Coleoptera: Curculionidae Distribution: A native of the Mediterranean basin. First released in NSW in 1993, it has been re-distributed to Vic. Beneficial status: Major, widespread, regular in NSW; unknown in Vic. Fig. 16.34. Adult onopordum crown weevil on Scotch thistle. The body is 20 mm long. (CSIRO Entomology)
Identification: Adult weevils are elongate and up to 15 mm long with a medium-sized rostrum 463
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Fig. 16.36. Adult onopordum stemboring weevil on a Scotch thistle stem. The body is 20 mm long. (CSIRO Entomology)
(Fig. 16.36). When they emerge in spring, they are reddish-brown but darken with time until they appear blackish by late summer. The creamy white eggs are covered with raised plant frass on the thistle stems. The larva is white, legless and has a brown head. May be confused with: Unlikely to be confused with any other insects occurring on Onopordum thistles. Host range: The onopordum stemboring weevil is restricted to stem-forming thistles of the genus Onopordum, which include Scotch and Illyrian thistle and their hybrids in Australia (Briese et al. 2002b). Life cycle: The Onopordum stemboring weevil has one generation per year. Adults emerge in early spring and feed on the rosette leaves of Scotch and Illyrian thistle. As soon as the thistles bolt, adults feed voraciously on the new leaf tissue and females lay their eggs into the bolting stems. One to five eggs are laid in a hole chewed in the stem, secured by a frass plug. Females may lay up to 70 eggs over a period of 2–3 months. Newly-hatched larvae bore directly into the stem and feed on structural tissue, eventually pupating inside the stems by late summer. New adults develop, but remain in the dead stems over summer and winter until the following spring.
Fig. 16.37. Damage to Scotch thistles caused by adult onopordum stemboring weevil feeding on leaf and stem tissue. (CSIRO Entomology)
on structural tissue inside the thistle stems. Several hundred larvae can develop in a single stem. Heavy adult feeding can lead to up to 50% reduction in plant size and 70% reduction in seed production (Fig. 16.37).
Onopordum rosette moth Eublemma amoena (Hbn.) Lepidoptera: Noctuidae Distribution: A native of the Mediterranean basin. First released in NSW in 1998, where it is currently being re-distributed and WA in 2000. Beneficial status: Minor, restricted, irregular in NSW. Establishment uncertain in WA. Identification: Adult moths are 15 mm long and have mottled light brown and white wings with two wavy dark brown bars visible when resting (Fig. 16.38). Eggs are bluish-green and are laid into the cottony hairs of thistle leaves. Newly hatched larvae are dark olive-green, but turn
Active period: Adults are active from October until January. Larvae feed in the stems of thistles from November to January. Damage: Adults chew shot-holes in the thistle leaves and necrotic lesions may develop from these further destroying leaf tissue. Larvae feed
Fig. 16.38. Adult rosette moth on Scotch thistle. The body is 15 mm long. (CSIRO Entomology)
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Damage: Newly hatched larvae feed within the rosette leaf petiole causing the leaf to become deformed and die (Fig. 16.39). Older larvae may bore into the crown tissue and upper tap-root of the thistle if plant leaves are stressed, which can lead to plant death. Larvae may also attack stem leaves in summer.
Spear thistle Cirsium vulgare Asteraceae: Cardueae Fig. 16.39. Damage to Scotch thistles caused by larvae of onopordum rosette moth feeding in the leaf petioles. (CSIRO Entomology)
greyish-green as they develop within the leaf petioles, growing up to 20 mm long. The pupal case is a similar grey to the dead Onopordum leaves in which it frequently is found. May be confused with: Unlikely to be confused with any other insects occurring on Onopordum thistles. Host range: The onopordum rosette moth is restricted to stem-forming thistles of the genus Onopordum, which include Scotch and Illyrian thistle and their hybrids, and stemless thistle in Australia (Briese et al. 2002b). Life cycle: The onopordum rosette moth can pass through three generations per year. Firstgeneration moths emerge in mid- to late-spring and, after mating, lay their eggs into the fine hairs covering the leaves of Onopordum rosettes. Female rosette moths live from 2–4 weeks and lay over 80 eggs each. Newly hatched larvae feed within the leaf petiole before pupating among dry leaf material on or near the crown. The spring generation may last for 70 days. The summer generation is shorter, lasting for about 50 days. Female moths of this generation also lay on stem leaves as well as those of perenniating rosettes, and the larvae feed on these. Larvae of the third generation do not pupate, but overwinter as late instars inside hard cocoons within the leaf petioles or crown tissue. Active period: The moths are night-flying and adults of different generations are active from mid-spring to autumn, and larvae may be present in the leaf petioles from late spring to late autumn.
Spear thistle is native to Europe, western Asia and North Africa. It is an annual or sometimes biennial herb up to 1.5 m reproducing by seed. In Australia, it is an important weed of pastures and neglected lands, occurring in all states except the NT. The plants, which are not grazed by animals, grow vigorously in situations where nitrogen levels are high. Seeds are dispersed by wind, but some of them fall in the vicinity of the plants where they germinate after autumn rains. Seeds which do not germinate readily can remain dormant for years. One biological control agent is currently established on spear thistle.
Spear thistle gall fly Urophora stylata Fabricius Diptera: Tephritidae Distribution: A native of Europe. First released in Vic. in 1994, it has been re-distributed in Vic. and NSW. Beneficial status: Minor, restricted, irregular. Identification: A generally black fly with a brown slightly striped thorax and a pale yellow scutellum (rear thorax). The wings are clear except for three dark cross-bands. Wings are 4.0–5.0 mm long and the ovipositor is about 3 mm long (Fig. 16.40). The eggs are white,
Fig. 16.40. Female gall fly (7 mm long) on spear thistle leaf. (VDPI)
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0.7 × 0.2 mm, and laid into unopened thistle flower heads. The small white larvae live within a woody gall in the thistle head. The brown pupa also occurs in the thistle head. May be confused with: May be confused with other Tephritidae (fruit flies), though wing patterns are different. Host range: The spear thistle gall fly is restricted to thistles of the genus Cirsium. Life cycle: The first adults appear in February and females lay eggs among or inside the florets. Larvae feed on developing achenes (seeds) and the capitula receptacle where cells around the larvae form a hard gall. Pupation occurs inside the gall and adults emerge in April. Active period: Adults are active in the field in February and April and are known to live up to 2 months. Damage: The woody gall formed by larvalfeeding is a metabolic sink that sequesters resources from both within and outside the attacked head. The effect is to reduce growth, flowering and seed production. Damage varies according to fly density and the number of larvae developing per capitula. Seed reduction as high as 45% has been measured in Vic., but reduction in the range of 5–30% is more common at most sites.
St John’s wort Hypericum perforatum Clusiaceae St John’s wort originally occurs widely throughout Europe and northern Asia and has been introduced to North and South America, South Africa, Australia and New Zealand. It is a toxic plant, causing photosensitisation and liver damage to stock, and is considered a serious weed in NSW and Vic. St John’s wort is also considered an environmental weed due to its ability to invade areas of open sclerophyll forest. It is also present but less common in SA, Tas. and WA. Six biological control agents have been established on St John’s wort, but only four of these have any impact (Briese 1997). Biological agents against St John’s wort are best integrated with other methods: herbicides, cultivation, manual removal, properly timed
heavy grazing by sheep and goats, planting trees, use of clovers and superphosphate.
St John’s wort stunt mite Aculus hyperici (Liro) Acarina: Eriophyidae Distribution: A native of the Mediterranean basin. First released in 1991, it has been widely re-distributed in NSW and Vic. Beneficial status: Moderate, widespread, regular in NSW and Vic. Identification: This species is a minute mite 0.15 mm long × 0.05 mm wide needing a ×10 hand lens or ×20 microscope to be seen properly. It is elongate with four legs. There are two nymphal stages, similar in shape to adults but smaller: the first is 0.075 mm long, and the second is 0.11 mm long. Eggs are spherical and about 0.04 mm diameter. May be confused with: Unlikely to be confused with any other insects occurring on St John’s wort. Host range: The mites can reproduce on some other Hypericum species, including the Australian native Hypericum gramineum, but decline in numbers after some generations. Life cycle: At 20°C the life cycle is completed in 25 days, with potentially many generations per year. All stages may be found on the plant throughout the year. Males deposit sperm packages on leaves, which are picked up in the genital opening of the female mite, and the eggs are fertilised within the female. Unfertilised females produce all male progeny. Mite populations start increasing during spring and summer, starting in young plant buds and developing in flowers and fruit. Mites mainly disperse by wind but may also be carried by other insects. The mites overwinter in the terminal and axillary buds of the rosette. Active period: Spring and summer. Damage: Mite-feeding (over 20 per bud) and the possible production of a toxin, causes stunting of the plants and loss of vigour which, after several years results in plant death, particularly if the plant is under environmental stress or competition from other pasture species. Reduced seed production eventually limits regeneration
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of the weed. Previously dense stands of the weed become progressively shorter and less dense. Resistance: Populations of St John’s wort resistant to mite damage occur in NSW. These may be derived from different introductions to other Australian populations of the weed.
St John’s wort aphid Aphis chloris Koch Hemiptera: Aphididae Distribution: A native of the Mediterranean basin. First released in 1986, it has been widely re-distributed in NSW and Vic. Beneficial status: Minor, widespread, irregular. Identification: Adult aphids are 0.9–1.3 mm long and lime green in colour during spring and summer, but darker green over winter. Nymphs are pale green. St John’s wort aphid forms colonies on the stems of St John’s wort and winged forms are produced when colonies become crowded (Fig. 16.41). Colonies are often attended by ant species. May be confused with: Other aphids such as Myzus persicae and Macrosiphum euphorbiae occasionally colonise St John’s wort, but these are quite a bit larger than St John’s wort aphid and have longer legs. Host range: The St John’s wort aphid is restricted to St John’s wort (Hypericum perforatum) in Australia. Life cycle: The St John’s wort aphid has a short life cycle: 6 days at 25°C and 10–11 days at 18°C.
Between 10 and 20 generations per year can develop in different parts of southern Australia, depending on temperature. The strain introduced into Australia is parthenogenetic and viviparous (reproduces by giving birth to live nymphs). New colonies are established when dispersing winged forms settle on a plant and commence to produce nymphs. These develop into wingless reproductive adults and colonies build up on the stems and flower clusters of St John’s wort until overcrowding causes the production of more winged forms and further dispersal. Colony sizes peak in midsummer and decline toward winter, when adults darken and pass the winter on rosette foliage. Parasitism and predation often restrict the build-up of aphid populations (Briese and Jupp 1995). Active period: Adults are present all year round and nymphs are common during the warmer months in the aphid colonies. Both are more obvious on flowering stems in spring and summer. Winged forms have been more commonly seen in late summer and autumn. Damage: The adults and nymphs suck phloem from the foliage and stems of St John’s wort. The plants may wilt and die when attacked by a dense population. Lower population levels can lead to reduced growth and dwarfing of leaves.
St John’s wort leaf beetle Chrysolina quadrigemina (Suffrian) Coleoptera: Curculionidae Distribution: A native of the Mediterranean basin. First released in Vic. in 1939, it has been widely re-distributed throughout all states of southern Australia. Beneficial status: Major, widespread, irregular.
Fig. 16.41. Colony (mainly nymphs) of darker green overwintering St John’s wort aphids on a stem. The body of the largest is 1.2 mm long. (CSIRO Entomology)
Identification: Adult beetles are rounded, 6– 7 mm long, and either metallic bronze (most common), blue, green or purple in colour (Fig. 16.42). Adults rarely fly, unless temperatures are very high. The reddish-orange, 1.2 mm long, oval-shaped eggs are laid singly or in clusters on the prostrate rosette leaves. The larvae are orange, tending to greenish-grey as they mature, with a blackish head, a pronounced dorsal arch and broad abdomen. Resting, they remain curled, but lengthen when moving 467
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feed voraciously on the foliage of new flowering stems of the plant, building up fat reserves. After feeding, the adults pass the summer in a dormant state in the soil and litter or under bark. They are stimulated to resume activity by shortening day length, emerging in autumn to lay eggs of the next generation, before dying in late winter or early spring.
Fig. 16.42. Adult leaf beetles defoliating a flowering stem of St John’s wort. The body is 7 mm long. (CSIRO Entomology)
(Fig. 16.43). The pupae are orange and are found in pupal cells 3–5 cm deep in the soil. May be confused with: Chrysolina hyperici. This biocontrol agent was released in 1930, but has been largely displaced by C. quadrigemina. Both are of similar form but C. hyperici is slightly smaller. If beetles are all bronze, they may be either species; if other colours are also present, they are definitely C. quadrigemina. Host range: The St John’s wort leaf beetle is restricted to St John’s wort (Hypericum perforatum) in Australia. Life cycle: The St John’s wort leaf beetle has one generation per year. Females lay well over 1000 eggs each from March to September on prostrate rosette foliage. The larvae emerge and feed on rosette leaves before pupating in the soil near the plants. New adults emerge in spring and
Active period: Adults are active from October to December, when they are highly visible feeding on flowering stems of St John’s wort. They sometimes form distinctive feeding fronts, beyond which are undamaged yellow-flowering St John’s wort plants and behind which all plants have been completely defoliated. They are much less conspicuous when laying eggs from March to September. Larvae of the leaf beetle feed on rosette growth over winter and early spring. Damage: Both larvae and adults can reach densities that cause complete defoliation. Summer defoliation by the adults is striking, but less effective than larval defoliation of winter rosette growth. Defoliation may lead to the death of plants in a dry summer. In Mediterranean parts of Australia, control has been very successful, but in summer rainfall areas, plants can recover to some extent from defoliation and periodic cycles of control and weed recovery occur. The beetles are not effective in shaded situations.
St John’s wort midge Zeuxidiplosis giardi (Kieffer) Diptera: Cecidomyiidae Distribution: A native of the Mediterranean basin. First released in 1953, it has been widely re-distributed in south-eastern Australia. Beneficial status: Minor, widespread, irregular.
Fig. 16.43. Larvae of the leaf beetle feeding on winter growth of St John’s wort. The body is 7 mm long. (CSIRO Entomology)
Identification: Adults are slender greyish flies, 3 mm long, and females have a bright reddishorange distended abdomen when ready to lay. The eggs are red, curved and 0.3 mm long. Larvae are also red, growing to 3 mm long. Larval-feeding causes galling of growth apexes. The gall consists of two opposing leaves modified to form a thickened egg-shaped bivalve shell within which larvae live and feed singly or in small groups. Often a whitish pupal
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Host range: The St John’s wort gall midge is restricted to St John’s wort (Hypericum perforatum) in Australia.
Fig. 16.44. Galled leaves of St John’s wort with an empty pupal skin from where the adult midge has emerged . The skin is 3 mm long. (CSIRO Entomology)
skin can be seen protruding from the lips of a gall after the new adult has emerged (Fig. 16.44). May be confused with: Unlikely to be confused with any other insects occurring on St John’s wort.
Life cycle: The St John’s wort gall midge can pass through five generations per year. The midge hibernates over winter as pupae within galls, emerging during September and early October. The adults live for less than a week, during which time they mate and lay 100–200 eggs on leaves, stems and terminal shoots. Emerging larvae then form galls within which they feed and pupate. Two more generations of midges develop by mid-summer, when larvae emerging in January may enter summer dormancy until March. A fourth generation develops in autumn and the fifth generation hibernates over winter. Active period: Galling is evident throughout the year. Damage: Heavy galling can cause reduced growth of large plants and death of St John’s wort seedlings, although such densities are only occasionally reached in the field.
Sources of information Briese, D.T. (1997). Biological control of St John’s wort in Australia: Past, present and future. Plant Protection Quarterly 12: 73–80. Briese, D.T. and Jupp, P.W. (1995). Establishment, spread and initial impact of Aphis chloris Koch (Hemiptera: Aphididae), introduced into Australia for the biological control of St. John’s Wort. Biocontrol Science and Technology 5: 271–285. Briese, D.T., Pettit, W.J., Swirepik, A.E. and Walker, A. (2002). A strategy for the biological control of Onopordum spp. thistles in south-eastern Australia. Biocontrol Science and Technology 12: 121–137. Briese, D.T., Walker, A., Pettit, W. and Sagliocco, J-L. (2002). Host-specificity of candidate agents for Onopordum spp. thistles in Australia: An assessment of testing procedures. Biocontrol Science and Technology 12: 149–163. Burdon, J.J., Groves, R.H. and Cullen, J.M. (1981). The impact of biological control on the distribution and abundance of Chondrilla juncea in south-eastern Australia. Journal of Applied Ecology 18: 957–966. Ireson, J.E., Gourlay A.H., Kwong, R.M., Holloway, R.J. and Chatterton, W.S. (2003). Host specificity, release and establishment of the gorse spider mite, Tetranychus lintearius Dufour (Acarina: Tetranychidae), for the biological control of gorse, Ulex europaeus L. (Fabaceae), in Australia. Biological Control 26: 117–127. Ireson, J.E., Friend, D.A., Holloway, R.J. and Paterson, S. (1991). Biology of Longitarsus flavicornis (Stephens) (Coleoptera: Chrysomelidae) and its effectiveness in controlling ragwort (Senecio jacobaea L.) in Tasmania. Journal of the Australian Entomological Society 30: 129–141.
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McLaren, D.A. (1992). Observations on the life cycle and establishment of Cochylis atricapitana (Lepidoptera: Cochylidae), a moth used for the biological contol of Senecio jacobaea in Australia. Entomophaga 37: 641–648. McLaren, D.A., Ireson, J.E. and Kwong, R.M. (2000). Biological control of ragwort (Senecio jacobaea L.) in Australia. In: Proceedings of the X International Symposium on Biological Control of Weeds. (Spencer, N.R., ed.). Montana State University, Bozeman. pp. 67–79. Sagliocco, J.L. and Coupland, J. (1995). Biology and host-specificity of Chamaesphecia mysiniformis (Lepidoptera: Sesiidae), a potential biological control agent of Marrubium vulgare (Lamiaceae) in Australia. Biocontrol Science and Technology 5: 509–515. Scott, J.K. and Yeoh, P.B. (1999). Bionomics and the predicted distribution of the aphid Brachycaudus rumexicolens (Hemiptera: Aphididae). Bulletin of Entomological Research 89: 97–106. Stone, C. (1986). An investigation into the morphology and biology of Tetranychus lintearius Dufour (Acari: Tetranychidae). Experimental and Applied Acarology 2: 173–186. Weiss, J., Ainsworth, N. and Faithfull, I. (2000) Horehound Best Practice Management Guide for Environmental Weeds. No. 8. CRC for Weed Management Systems, University of Adelaide. Woodburn, T.L. (1996). Reduction of seed set in nodding thistle (Carduus nutans) by the seed-fly Urophora solstitialis, in Australia. In: Fruit Fly Pests: A World Assessment of their Biology and Management. (McPheron, B.A., and Steck, G.J., eds). St Lucie Press, Florida. pp. 165–169. Woodburn, T.L. (1997). Establishment in Australia of Trichosirocalus horridus a biological control agent for Carduus nutans, and preliminary assessment of its impact on plant growth and reproductive potential. Biocontrol Science and Technology 7: 645–656. Woodburn, T.L. and Briese, D.T. (1996). The contribution of biological control to the management of thistles. Plant Protection Quarterly 11: 250–253. Woodburn, T.L. and Cullen, J. (1996). Impact of Rhinocyllus conicus and Urophora solstitialis on achene set in Carduus nutans in Australia. In: Compositae: Biology and Utilization. Proceedings of the International Compositae Conference, Kew, 1994. (Caligari, P.D.S. and Hind, D.J.N., eds). Vol. 2. pp. 307–319.
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17 PASTURES—DUNG BEETLES P.B. Edwards and C. Pavri
(a) Accumulated cattle dung provides breeding sites for a number of species of flies
COMMON NAME AND SPECIES Introduced species in winter rainfall pastures Euoniticellus fulvus Southern sandy dung beetle, Euoniticellus pallipes Geotrupes spiniger Onitis aygulus Onitis caffer humpbacked dung beetle, Onthophagus binodis bullhorned dung beetle, Onthophagus taurus Introduced species in summer rainfall pastures greater sandy dung beetle, Euoniticellus africanus northern sandy dung beetle, Euoniticellus intermedius yellowshouldered dung beetle, Liatongus militaris bronze dung beetle, Onitis alexis Onitis caffer Onitis pecuarius emerald dung beetle, Onitis viridulus gazella dung beetle, Onthophagus gazella brown dungball roller, Sisyphus rubrus grey dungball roller, Sisyphus spinipes
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COMMON NAME AND SPECIES
PAGE
Native dung beetles in winter rainfall pastures western dung beetle, Onthophagus ferox five horned dung beetle, Onthophagus pentacanthus Native dung beetles in summer rainfall pastures Onthophagus atrox common northern dung beetle, Onthophagus consentaneus Onthophagus incanus Onthophagus muticus Onthophagus quadripustulatus Native dung beetles in summer and even rainfall temperate pastures southern dung beetle, Onthophagus australis granulose dung beetle, Onthophagus granulatus
Biological control of dung in pastures The aim of the CSIRO Australian Dung Beetle Project was to redress the ecological imbalance created by the accumulation of large amounts of cattle dung in Australian pastures following the arrival of the first European settlers and their cattle in 1788. Dr George Bornemissza recognised that, in contrast to other parts of the world, Australian pastures were littered with unburied cattle pads, some of which persisted for up to several years. Dung accumulation was attributed to lack of an effective dung beetle fauna working in cattle dung and Bornemissza’s proposed solution was to introduce a range of dung beetle species adapted to bury the moist dung pads of large ruminants. The project commenced in 1964 and lasted approximately 20 years. During that time nearly 50 species of dung beetles were introduced, mainly from Africa and Europe. Species were selected to suit different climatic regions of Australia, different soil and habitat types, and with different daily and seasonal activity patterns. Dung beetle introductions were aimed at controlling pest flies that breed in cattle dung, in particular the buffalo fly, Haematobia irritans exigua de Meijere (Diptera: Muscidae), and the bush fly Musca vetustissima Walker. Both adult and larval dung beetles feed on dung; the adults squeeze the dung through their mouth parts, extracting and feeding only on dung fluid, while larvae cut and chew the dung, feeding on both fibre and fluid. By burying dung rapidly, dung beetles destroy the breeding site of these flies. The life cycles of other pests of cattle, such as
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internal parasites, are also disrupted by dung beetle activity. In addition, rapid burial of dung pads reduces pasture fouling, and limits the growth of rank unpalatable grass around dung pads. The tunneling associated with good dung beetle activity assists root penetration and improves water infiltration into the soil, thus reducing run-off and pollution of waterways. Additionally, burial of dung results in accelerated and improved nutrient cycling. Thus, dung beetles contribute to improved soil health and structure, and improved pasture quality. They also provide substantial and sustainable environmental benefits.
Effectiveness of dung beetles A study of buffalo flies at Rockhampton in 1983– 1984 (Fay et al. 1990) showed that the dung fauna (dung beetles plus predators) killed about 80% of buffalo flies throughout the season, and 98–99% on three occasions during February and March. These early results are supported by recent anecdotal evidence from Qld cattle producers who are able to reduce or eliminate the use of chemicals to control buffalo fly during times of good dung beetle activity. A study of bush flies at Dardanup in WA (Ridsdill-Smith and Matthiessen1988) detected a 99.5% mortality of bush flies in December 1986 in the presence of dung beetles, compared with 21% mortality in dung pads from which dung beetles were excluded. Additionally, anecdotal evidence from around Australia indicates that, during periods of good dung beetle activity, people are able to enjoy outdoor pursuits that had previously been
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adult breaks out of dung ball and digs its way to the soil surface pupa
larva
young adult fresh dung pat
adults pair off egg dry remains of crust
soil casts
tunnel filled with soil
tunnel often lined with dung brood mass containing several eggs
brood ball containing one egg
ball is attached to base of surrounding vegetation
ball is rolled away
ball is cut out of dung pad
Fig. 17.1. The life cycle of a dung beetle (Tyndale-Biscoe 1990).
made unbearable by the annoyance of bush flies. The impact of dung beetles on pasture productivity is indicated by a small pot trial by Bornemissza and Williams 1970, who showed that dung beetle activity increased plant yield (including roots) by approximately 50%. The volume of dung buried or shredded by dung beetles has been measured in several studies (e.g. Tyndale-Biscoe 1994; Doube et al. 1988), and at times can reach 100%.
Breeding activity of introduced beetles There are two main types of breeding behaviour in dung beetles (Fig. 17.1). The most common is that of the ‘tunnelers’ or paracoprids. These
dung beetles bury dung beneath the dung pads. The female beetle digs a tunnel and the male passes dung down from the pad above. The female moulds dung into individual brood balls into which one egg is laid, or into longer, sausage-shaped brood masses into which between two and four eggs are laid. The larvae hatch from the eggs and feed on the dung in the brood balls. Larvae grow through three distinct stages, or instars, and at the end of the third instar they cease feeding. In some species, if conditions are unfavourable, development is delayed at this stage and the larvae remain inactive for many months. In due course, they change into pupae and then emerge as adult beetles that eat their way out of the brood ball, 473
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Table 17.1. Seasonal activity of introduced dung beetles in Australia
Species
No. generations per year
Onthophagus binodis Onthophagus taurus Onthophagus gazella Onitis alexis Onitis aygulus Onitis caffer (WR strain)
Several At least 2 Several 1–3 1–2 1
Onitis caffer (SR strain)
1
Onitis pecuarius Onitis viridulus Euoniticellus africanus Euoniticellus fulvus Euoniticellus intermedius
2–3 2–3 Up to 4 Up to 3 Up to 6
Euoniticellus pallipes Liatongus militaris Sisyphus spinipes Sisyphus rubrus
At least 2 NA Several Several
Adult activity Late spring–autumn Spring–autumn Spring–autumn Spring–autumn Spring–autumn Autumn, early winter, spring Autumn, early winter Spring–autumn Spring–autumn Late spring–autumn Spring–autumn Spring–autumn, warm winter spells Spring–autumn Spring–autumn Spring–autumn Spring–autumn
Adult life span in summer
Development period in summer (egg to adult)
Flight time
9 weeks 2–3 months 2 months 3 months > 2 months 3–4 months
4–6 weeks 6–8 weeks 4–6 weeks 2–5 months 2.5–3 months 10–12 months
Day Day Dusk and dawn Dusk and dawn Dusk and dawn Dusk and dawn
3–4 months
10–12 months
Dusk and dawn
2–3 months 2–3 months 2–3 months 2–3 months 1–2 months
NA 1.5–2 months 1–2 months 5–8 weeks 4–6 weeks
Dusk and dawn Dusk and dawn Day Day Day
2–3 months NA 6–11 weeks 2–6 months
5–7 weeks NA 7–11 weeks 5–16 weeks
Day Day Day Day
NA = not applicable, WR = winter rainfall, SR = summer rainfall.
dig up to the soil surface and disperse in search of fresh dung. The development time between egg-laying and adult emergence varies between 1 month to a year or more, depending on the species of beetle and seasonal conditions. The other main type are the ‘ball rollers’ or telecoprids. A male and female mould a ball of dung from the pad, and roll it away together. The female then buries the ball down a short shaft that she digs in the soil and lays an egg into it (e.g. Sisyphus rubrus), or attaches the ball to the base of grasses or other vegetation before laying an egg in it (e.g. Sisyphus spinipes). Adult beetles of most introduced species feed and breed throughout the late spring, summer and autumn and die in winter. It is usually the larvae of these species that overwinter in the soil, but in warmer areas adults may also survive the winter in the soil. A few species have adults that overwinter in the soil, then feed and breed in spring and die during summer. The larvae of these species emerge as adults the following autumn (Table 17.1).
I N T RO D U C E D S PE C I E S I N W I N T E R R A I N FA L L PA S T U R E S Introduced dung beetles that are common in winter rainfall and even rainfall pastures of southern Australia include the following.
Euoniticellus fulvus (Goeze) Coleoptera: Scarabaeidae: Scarabaeinae Strains from France and Turkey were introduced between 1978 and 1983 in NSW, SA, Tas., Vic., and WA, and the species has established in all these states. The adult is a brownish-yellow beetle, 8–12 mm in length with a light brown pronotum, which has no speckling. The wing covers may have darker brown markings, but no speckling (Fig. 17.2). Adult beetles are active between spring and autumn. This species can produce up to three generations a year. Development time from egg to adult is 5–8 weeks in summer and longer in winter. Adult lifespan in summer is between 2
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Fig. 17.4. Geotrupes spiniger female, 20–25 mm long. (CSIRO Entomology)
Fig. 17.2. Euoniticellus fulvus, 12 mm long. (CSIRO Entomology)
and overwintering probably takes place in the larval stage.
and 3 months. Flight activity of beetles to fresh pads occurs during the day.
The adults are most abundant in summer and autumn. They breed mainly from January to March and fly to fresh dung pads during the day.
Southern sandy dung beetle
Geotrupes spiniger Marsham Coleoptera: Geotrupidae
Euoniticellus pallipes (Fabricius) Coleoptera: Scarabaeidae: Scarabaeinae Strains from Iran and Turkey were introduced between 1977 and 1982 in NSW, SA and WA, and the species has since been recorded in NSW, SA, Vic. and WA. The adult is a brownish-yellow beetle, 9–12 mm in length with dark brown to black speckling on the pronotum and wing covers (Fig. 17.3). Development time from egg to adult during summer is 5–7 weeks and much longer in the winter. There are at least two generations a year
Introduced from France between 1979 and 1982, and now established in ACT, NSW, Vic. and Tas. It has recently been redistributed in SA. The adult beetle is 20–25 mm in length with black dorsal surface and metallic blue ventral surface (Fig. 17.4). Between spring and autumn, development time from egg to adult is about 6 months, while eggs laid in the autumn take up to a year to develop. There is only one generation a year. This species overwinters both in the adult and larval stage and probably oversummers in the larval stage. The adults are active in the autumn, early winter and again in spring, and adult beetles fly to fresh pads at dusk and dawn.
Onitis aygulus (Fabricius) Coleoptera: Scarabaeidae: Scarabaeinae Introduced from South Africa into Australia, releases were made between 1977 and 1982 in NSW, SA and WA and the species has established in drier grassland regions in all three states and Vic.
Fig. 17.3. Euoniticellus pallipes, 9–12 mm long. (CSIRO Entomology)
The adult is one of the larger dung beetles present in winter rainfall pastures with a length of 20–25 mm, it has a dark brown pronotum, sometimes with a green sheen, and the wing 475
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Fig. 17.7. Onthophagus binodis, 13 mm. (CSIRO Entomology)
Fig. 17.5. Onitis aygulus male, 25 mm long. Inset: double spur (arrowed) on hind femur of male, absent in females. (CSIRO Entomology)
covers are light brown (Fig. 17.5). Males have a characteristic double spur on the hind femur (Fig. 17.5 inset) and females have no spur. Development time from egg to adult during summer is 2.5–3 months and up to 10 months in winter. The adults usually emerge in late spring– early summer and are abundant for 1 month; the second generation emerges in late summer–early autumn. The species spends the winter as larvae in brood masses in the soil. Adult beetles are active from spring to autumn. Flight activity occurs at dusk and dawn.
Onitis caffer Boheman Coleoptera: Scarabaeidae: Scarabaeinae A winter rainfall strain was introduced from South Africa, and released in WA in 1982 and 1983. It has established at some sites in WA. It has recently been re-distributed into NSW and
SA. The adult beetle is shiny black in colour and 15–20 mm in length (Fig. 17.6) The males have a serrated ridge on the hind femur (Fig. 17.6 inset), while females have no serration. Adult beetles are active in autumn and early winter and become active again in spring in areas with winter rainfall. There is only one generation a year, with development time from egg to adult usually taking 10–12 months, with adults living 3–4 months. The species oversummers as larvae and overwinters in both larval and adult stages. Flight activity of beetles to fresh dung occurs at dusk and dawn.
Humpbacked dung beetle Onthophagus binodis Thunberg Coleoptera: Scarabaeidae: Scarabaeinae Introduced into Australia from South Africa between 1971 and 1983 in NSW, Qld, SA, Tas., Vic., WA, and Norfolk Island. It has since become established in temperate regions of all these states. The adult beetle is matt black in colour and 11–13 mm in length with a hump on the pronotum (pronotal lobe) that is pronounced in males (Fig. 17.7) and smaller in females. Adult beetles are active between late spring and autumn. There are several generations a year, with development time from egg to adult taking 4–6 weeks in summer and longer in winter, and the adult life span lasting around 9 weeks in summer. Flight activity of beetles to fresh pads occurs during the day.
Bullhorned dung beetle Onthophagus taurus (Schreber) Coleoptera: Scarabaeidae: Scarabaeinae Fig. 17.6. Onitis caffer, 20 mm long. Inset: the hind femur of males has a serrated edge (arrowed), absent in females. (CSIRO Entomology)
Strains from Spain, Greece, Italy and Turkey were released in Australia between 1975 and 1984 in NSW, SA, Tas., Vic. and WA and the
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Fig. 17.8. Onthophagus taurus major male, 10 mm long. (CSIRO Entomology)
species has become established in all these states.
Fig. 17.9. Euoniticellus africanus, 8–13m long. (CSIRO
The adult beetle is shiny black in colour and 8–10 mm in length (Fig. 17.8). Some (major) males have a pair of curved frontal horns extending backwards half way over the pronotum. Other (minor) males have very small horns projecting upwards from the head. The females have no horns.
Introduced from South Africa and released between 1971 and 1984 in NSW, Qld, SA, Tas., Vic. and WA. Established in the summer rainfall area of NSW and south-east Qld, west of the Great Dividing Range. Adults are 8–13 mm in length, have a narrow shape and are yellow–brown in colour. They have two shiny black spots on the thorax (Fig. 17.9). Neither sex has a horn on the head. Adults are active from October to April. Development from egg to adults takes 1–2 months in the summer, and it may complete three to four generations per year in the warmer parts of its range. Adults fly to fresh dung during the day.
The beetles breed in the spring and are most abundant in summer. Beetles are more abundant at sites with summer moisture or irrigation and breeding continues at these sites during the summer. Development time from egg to adult during summer is 6–8 weeks, but it is much longer in winter. The adult lifespan is 2–3 months. There are at least two generations a year. In the southern range of its distribution, the species overwinters in the larval stages. Adult beetles are active from spring to autumn and fly to fresh pads during the day. Other introduced species established in winter rainfall pastures include Onitis alexis (see next section), Euoniticellus intermedius (see next section), Bubas bison and Copris hispanus.
Entomology)
Northern sandy dung beetle Euoniticellus intermedius (Reiche) Coleoptera: Scarabaeidae: Scarabaeinae Introduced from South Africa, and released between 1971 and 1984 in all mainland states. The current distribution is NSW, NT, Qld, SA, Vic. and WA, but it is most widespread and abundant in summer rainfall regions. Adults are 7–9 mm in length with narrow bodies. They are yellow–brown with a faint diamond-shaped
I N T RO D U C E D S PE C I E S I N S U M M E R R A I N FA L L PA S T U R E S Introduced dung beetles that are common in summer rainfall pastures of eastern and northern Australia include the following.
Greater sandy dung beetle Euoniticellus africanus (Harold) Coleoptera: Scarabaeidae: Scarabaeinae
Fig. 17.10. Euoniticellus intermedius, 7–9 mm long. (CSIRO Entomology)
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Fig. 17.11. Liatongus militaris 8–10 mm long. (CSIRO Entomology)
pattern on the thorax. Males have a blunt horn on the head (Fig. 17.10). Adults are active between spring and autumn and during warm spells even in the cooler months. This species can produce up to six generations a year. Development time for egg to adult is 4–6 weeks in summer and longer in winter. Adult lifespan in summer is 1–2 weeks. Both the adults and larvae overwinter in the warmer regions, while in cooler regions, they overwinter in the larval stage. Flight activity of beetles to fresh pads occurs during the day.
Yellowshouldered dung beetle Liatongus militaris (Castelanu) Coleoptera: Scarabaeidae: Scarabaeinae Introduced from South Africa (via Hawaii), and released in NSW, NT, Qld, and WA between 1968 and 1979. It is established in NT, Qld, and subtropical parts of northern NSW. It is 8–10 mm long, and the overall colour is dark brown–black, with pale yellow patches on the sides of the thorax and broken stripes on the wing covers (Fig. 17.11) Adults are most abundant between November and April, and fly during the day.
Bronze dung beetle Onitis alexis Fabricius Coleoptera: Scarabaeidae: Scarabaeinae Occurs in Africa and southern Europe. Introduced from southern Africa between 1973 and 1984 into NSW, NT, Qld, and WA. Although established in winter rainfall and even rainfall regions, this species is most successful in the
Fig. 17.12. Onitis alexis, 15–20 mm long. Inset: males have a curved single spur (arrowed) on the hind femur. (CSIRO Entomology)
summer rainfall region. It is now present in NSW, NT, Qld, SA, Vic., and WA. Adult beetles are 15–20 mm long, the thorax is dark brown with a copper or greenish sheen, and the wing covers are light brown (Fig. 17.12). Males have a single curved spur on the hind leg (Fig. 17.12 inset); females have a small round bump at the back of the head. Adults emerge in spring and are abundant for about a month. The second generation emerges in autumn. The species spends the winter as adults or as larva in brood masses in the soil. Flight activity of beetles to fresh pads occurs mostly at dawn and dusk.
Onitis caffer Boheman Coleoptera: Scarabaeidae: Scarabaeinae A summer rainfall strain of this species was introduced from South Africa, and released at one site in Qld (Toowoomba) in 1983 and 1984. It established well, and in 2002 and 2003 was re-distributed throughout south-east Qld, and in 2004 was re-distributed to several sites on the northern NSW tablelands. Releases of an unknown strain were made in 1979 and 1980 at Moruya (southern NSW) where it has established. The adult beetle is shiny black in colour and 15–20 mm in length (Figure 17.6). Adult beetles are active in autumn and early winter, at a time when there is little other dung beetle activity. There is only one generation a year, with development time from egg to adult usually taking 10–12 months and adult lifespan of 3–4 months. They oversummer as larvae and overwinter in both larval and adult stages. Adults fly to fresh dung at dusk and dawn.
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Fig. 17.15. Male Onthophagus gazella, with a straight pair of short horns on the back of the head, 10–13 mm long. (CSIRO Entomology) Fig. 17.13. Onitis pecuarius, 18–23 mm. Males have a double spur on the hind femur (inset, arrowed) absent in females. (CSIRO Entomology)
Onitis pecuarius Lansberge Coleoptera: Scarabaeidae: Scarabaeinae Introduced from South Africa between 1976 and 1979 and released in NSW and Qld. It is well established in NSW, extending into the temperate south-east corner of Qld. Adults (Fig. 17.13) are very similar in appearance to Onitis viridulus, but the distribution of the two species overlaps in only a small region of southeast Qld. Males have a double spur on the hind femur (Fig. 17.13 inset), while females have no spur. Adults are active from spring to autumn, and fly to dung at dusk and dawn. Adults live for several months, and the species probably overwinters in the larval stage.
Emerald dung beetle Onitis viridulus Boheman Coleoptera: Scarabaeidae: Scarabaeinae
Fig. 17.14. Onitis viridulus 18–23 mm long. Males have an unequal double spur on the hind femur (inset, arrowed), absent in females. (CSIRO Entomology)
Introduced from South Africa, and released in NSW, NT and Qld between 1976 and 1980. It is now widespread in tropical and subtropical pastures of Qld, NT, far north-east NSW and the Kimberley region of WA. Adults are 18–23 mm in length, and are a uniform dark brown–black, with a greenish sheen (Fig. 17.14). Males have an unequal double spur on the hind leg (Fig. 17.14 inset), while females have no spur. Adults are active between September and May. Nests are made in the dung, just under the surface of the pad. Development time from egg to adult is 1.5 to 2 months, so the species can complete about three generations during the summer rainfall season. Adults fly at dusk and dawn.
Gazella dung beetle Onthophagus gazella (Fabricius) Coleoptera: Scarabaeidae: Scarabaeinae This species occurs over the hotter, drier parts of Africa south of the Sahara, also in Ethiopian and Oriental regions. Introduced throughout Australia and Norfolk Island between 1968 and 1984, it has established in all states except Tas. and Vic. It is widespread and abundant in Qld and NSW, and also occurs in NT and northern WA. Adult beetles are 10–13 mm in length, with a dark brown thorax and lighter brown wing covers. Males have a straight pair of short horns at the back of the head (Fig. 17.15). Adult beetles are active between spring and autumn and this species can produce several generations a year. Development time from egg to adult is 3–5 weeks in summer and much longer in winter. This species spends the winter in either the larval or adult stage. Flight activity of beetles to fresh pads occurs at dusk and dawn. 479
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A A B
B
Fig. 17.16. Sisyphus rubrus male, 6–8 mm long. The rear edge of the hind femur (A) is rounded and males have a pale spur (B) at the base of their hind leg. (CSIRO Entomology)
Brown dungball roller Sisyphus rubrus Paschalidis Coleoptera: Scarabaeidae: Scarabaeinae Introduced from South Africa and released in NSW, Qld and WA between 1973 and 1980. It is well established in subtropical Qld and northern NSW. It extends slightly further into the drier grassland regions of western Qld than does S. spinipes. It is 6–8 mm in length, and is light brown in colour with long spidery legs (Fig. 17.16). The rear edge of the hind femur is rounded (arrow A in Fig. 17.16) and males have a long pale spur at the base of the hind leg (arrow B on fig. 17.16). Most adult activity occurs between November and May, and adults can attain much higher densities than S. spinipes. A pair of male and female beetles rolls a ball of dung away from the dung pad and buries it at a depth of about 10 cm. The female lays a single egg in the ball. Development from egg to adult can be completed in 5 weeks in summer. Flight occurs throughout the warmer part of the day.
Grey dungball roller Sisyphus spinipes (Thunberg) Coleoptera: Scarabaeidae: Scarabaeinae Introduced from South Africa, and released in NSW, NT, Qld and northern WA between 1972 and 1978. It is well established in subtropical Qld and northern NSW. Adults are 8–10 mm in
Fig. 17.17. Sisyphus spinipes male, 8–10 mm long. The rear edge of the hind femur has an angle (A) and males have a dark spur (B) at the base of their hind leg. (CSIRO Entomology)
length, dark brown, with long spidery legs (Fig. 17.17). There is a distinct angle on the rear edge of the hind femur (arrow A on Fig. 17.17) and males have a dark spur at the base of the hind femur (arrow B on Fig. 17.17). Adults are most abundant between December and May. A male and female pair rolls a ball of dung away from the dung pad and attaches it to the base of a clump of vegetation. The female lays a single egg in the ball. Development from egg to adult takes about 6–11 weeks in summer. There are several generations a year, and overwintering occurs in the larval stage. Flight occurs throughout the warmer part of the day. Other introduced species that occur in summer rainfall pastures are Onthophagus sagittarius (within 100 km of Qld coast), Copris elphenor (established in only one region near Biloela, Qld, but re-distributed in 2001–2006 to 10 sites throughout Qld), Onitis vanderkelleni (in cooler, high rainfall regions), Onthophagus nigriventris (in cooler, high rainfall regions), and Onthophagus binodis (in temperate parts, see previous section).
N AT I V E D U N G B E E T L E S I N W I N T E R R A I N FA L L PA S T U R E S
Western dung beetle Onthophagus ferox Harold Coleoptera: Scarabaeidae: Scarabaeinae
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Fig. 17.19. Onthophagus pentacanthus, 14–20 mm long. (CSIRO Entomology)
Fig. 17.18. Onthophagus ferox, 12–20 mm long. (CSIRO
N AT I V E D U N G B E E T L E S I N S U M M E R R A I N FA L L PA S T U R E S
Entomology)
Present in WA; most abundant in the southwest of that state. The adult beetle is 12–20 mm in length, shiny black in colour with one horn on its head and two on the pronotum (Fig. 17.18). Adult beetles are active during the cool humid period of the year. To the north of Perth this is from May to September and to the south of Perth between May and December. Adult activity peaks in May and June. Adult beetles fly to fresh pads at night.
Five horned dung beetle Onthophagus pentacanthus Harold Coleoptera: Scarabaeidae: Scarabaeinae Present in ACT, NSW, SA, Vic., and into southern Qld. The adult beetle is 14–20 mm in length, entirely black in colour with reddish brown antennal clubs. A single, erect horn on the head is conical in shape and about 6 mm in major males and 1–2 mm in minor males. There are also four pronotal horns (projections) pointing upwards and forwards (Fig. 17.19). Females of the species have the same horns, which are proportionately smaller for a given individual size. Adult beetles are active throughout the year with most activity occurring in the winter months, even in summer rainfall regions. Adults fly to fresh pads at night.
Numerous native dung beetle species are active in cattle dung in summer rainfall pastures. Seventy-three species have been recorded from cattle dung in Qld. Some of the most widespread species are described below.
Onthophagus atrox Harold Coleoptera: Scarabaeidae: Scarabaeinae Widespread from northern NSW to northern Qld, particularly abundant in south-east Qld. The adult beetle is 12–19 mm in length, black in colour with reddish-brown antennal clubs and a raised ridge near the front of thorax, with a small tubercle on each side of ridge (Fig. 17.20). Adult beetles are active from September to June with a peak in April. Adults fly to fresh dung at night.
Common northern dung beetle Onthophagus consentaneus Harold Coleoptera: Scarabaeidae: Scarabaeinae The most widespread native Onthophagus species, it occurs in northern NSW, and
Fig. 17.20. Onthophagus atrox, 12–19 mm long. (CSIRO Entomology)
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Fig. 17.23. Onthophagus muticus. (CSIRO Entomology)
Fig. 17.21. Onthophagus consentaneus, 6–11 mm long. (CSIRO Entomology)
throughout Qld, NT and northern WA. The adult beetle is 6–11 mm long, and black in colour (Fig. 17.21), and is common in cattle dung, but also found under vertebrate carcasses and decaying mushrooms. It flies to dung during the day and at dusk and dawn.
Onthophagus incanus Macleay Coleoptera: Scarabaeidae: Scarabaeinae Common in much of Qld above the 400 mm isohyet, but not in the south-east corner, also in the Kimberley region of northern WA. It is a small beetle 4–7 mm in length, black with a greenish sheen, covered in fine hairs (Fig. 17.22). The species is common in cattle dung and wallaby dung and flies during the day.
Onthophagus muticus Macleay Coleoptera: Scarabaeidae: Scarabaeinae This species is common in northern coastal NSW, Qld above the 600 mm isohyet, the Darwin region of NT and the Kimberley region of WA. It is a shiny black beetle 6–9 mm in length (Fig. 17.23),
Fig. 17.22. Onthophagus incanus, 4–7 mm long. (CSIRO Entomology)
and is common in cattle dung. It has prehensile claws adapted for clinging to fur, and may congregate around the anal opening of wallabies, ready to grip onto an emerging dung pellet.
Onthophagus quadripustulatus (Fabricius) Coleoptera: Scarabaeidae: Scarabaeinae Distributed in New Guinea, throughout coastal Qld above the 600 mm isohyet and into far northern NSW, and the Kimberley region of northern WA. It is a small beetle 3.5–4.5 mm in length, black with a red patches at the outer front corner of each wing cover, and at the tip of the wing cover (Fig. 17.24).
N AT I V E D U N G B E E T L E S I N S U M M E R A N D E V E N R A I N FA L L T E M P E R AT E PA S T U R E S Native dung beetles in summer rainfall and even rainfall pastures in temperate regions include the following.
Southern dung beetle Onthophagus australis Guérin-Méneville Coleoptera: Scarabaeidae: Scarabaeinae
Fig. 17.24. Onthophagus quadripustulatus, 3.5–4.5 mm long. (CSIRO Entomology)
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adults commence activity in spring and continue throughout the summer until mid-autumn, with peaks during October and early in the following year. Adults fly to fresh pads during the day.
Monitoring beetle activity
Fig. 17.25. Onthophagus australis, 11–13 mm long. (CSIRO Entomology)
Distributed over most of south-eastern Australia, from southern Qld to SA and Tas. The adult beetle is 11–13 mm in length, black in colour with a green or coppery sheen. Females and minor males are without horns, while some males (major) have horns projecting upwards from the head (Fig. 17.25). This species has one generation a year. Development time from egg to adult is 3 months at 25°C and 7 months at 15°C. Adult emergence begins in the late summer and continues through autumn. Adult activity peaks in spring (September–October) and again in autumn (March–April). Overwintering occurs mainly in the adult stage. Spring is the main breeding period and the adults die off with hot summer weather. Beetles fly to fresh pads during the day.
Granulose dung beetle Onthophagus granulatus Boheman Coleoptera: Scarabaeidae: Scarabaeinae Present in NSW, Qld and Vic. The adult beetle is 6–8 mm in length with dark brown pronotum and light brown wing covers with dark brown markings and a granular surface (Fig. 17.26). This species has one generation a year. The
As dung beetle species are active for only certain periods of the year, a variety of species that are active at different times of the year is necessary to maintain dung burial throughout the year. Monitoring dung beetle populations involves assessing both beetle activity and the diversity of species present. A suggested monitoring program (based on notes prepared by NSW Agriculture) is as follows: 1
Monitor dung beetle activity and populations monthly.
2
Estimate the number of each species of dung beetle by following these guidelines: • Look for dung pads showing evidence of dung beetle activity, such as disturbed soil at the edge of the pad (1–2-day-old pads are preferable). Approach quietly, as dung beetles are sensitive to vibrations and will quickly descend down their tunnels. • Shovel up two dung pads into a bucket, taking about 25 mm of soil from under the pad. • Fill the bucket with water and stir well with a shovel to break up the dung. The dung beetles will float to the top; skim them off with a sieve. Stir again, skim again, and continue until no more beetles float to the surface. • Identify and count each species of dung beetle present. • Keep records of weather conditions each time you monitor. Weather has a huge impact on seasonal abundance, with most species being active in warm moist conditions.
Factors affecting dung beetle activity include:
Fig. 17.26. Onthophagus granulatus, 6–8 mm long. (CSIRO Entomology)
• Cold weather. Dung beetles are most active in warm moist conditions and many overwinter as larvae in the soil during the cool months of the year. 483
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• Dung from animals fed on high intake grain such as in feedlots. Cattle-growers have noticed very little dung beetle activity in dung produced from cattle in feedlots. Dung produced in a feedlot situation where cattle are fed grain has low pH and high nitrogen, which may repel the dung beetles. Beetles reproducing in feedlot dung have also shown a decrease in egg production, larval survival to adulthood and size of emerging larvae.
resulting in reduced larval survival. Some macrocyclic lactone chemicals, especially the avermectins, can also adversely affect the survival and development of larvae and can reduce egg-laying in newly emerged adults. If the use of these chemicals is necessary, try to treat animals at times of the year when beetle numbers are low (e.g. during the cooler months of the year when many of the species overwinter in the soil). Avoid using these chemicals, especially in spring, when dung beetles are first emerging after overwintering.
• Pesticides used for fly and parasite control in cattle. Many synthetic pyrethroid chemicals used for controlling external parasites leave residues in the dung of the treated animals. These residues are toxic to dung beetles that feed on the dung from these animals,
• Management of pastures to improve quality of dung. Improved quality of feed in pastures results in high quality dung with high moisture and nitrogen content. This results in higher rates of egg production by dung beetles.
• Predators, such as ibis, crows and foxes, which can have a significant impact on dung beetle populations.
Sources of information Bornemissza, G.F. and Williams C.H. (1970) An effect of dung beetle activity on plant yield. Pedobiologica 10: 1–7. Doube, B.M., Macqueen, A. and Fay, H.A.C. (1988) Effects of dung fauna on survival and size of buffalo flies (Haematobia spp.) breeding in the field in South Africa and Australia. Journal of Applied Ecology 25: 523–536. Fay, H.A.C., Macqueen A. and Doube, B.M. (1990) Impact of fauna on mortality and size of Haematobia spp. (Diptera: Muscidae) in natural dung pads in Australia and South Africa. Bulletin of Entomological Research 80: 385–392. Ridsdill-Smith, T.J. and Matthiessen, J.N. (1988) Bush fly, Musca vetustissima Walker (Diptera: Muscidae), control in relation to seasonal abundance of scarabaeine dung beetles (Coleoptera: Scarabaeidae) in south-western Australia. Bulletin of Entomological Research 78: 633–639. Tyndale-Biscoe, M. (1990). Common Dung Beetles in Pastures of South-eastern Australia. CSIRO Australia. Tyndale-Biscoe, M. (1994) Dung burial by native and introduced dung beetles (Scarabaeidae). Australian Journal of Agricultural Research 45: 1799–1808.
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18 LOCUSTS AND GRASSHOPPERS OF PASTURES AND RANGELANDS P. Walker, D. Hunter and R. Elder
PEST (major pests in bold)
PAGE
arid zone grasshopper complex
485
Australian plague locust
486
eastern plague grasshopper
489
migratory locust
490
mulga grasshopper complex
491
pyrgomorph grasshoppers
492
small plague grasshopper
493
spinifex grasshoppers
496
spur-throated locust
496
wingless grasshoppers
498
yellow-winged locust
502
Locusts are distinguished from grasshoppers by their behaviour. Australian locusts may change their behaviour from being solitary individuals to become gregarious when populations of nymphs are dense. The gregarious change in locusts usually presages large scale migrations from their breeding areas. Grasshoppers, on the other hand, generally remain solitary and usually do not move far from their breeding area. Locusts and a number of species of grasshoppers breed in pastures where they may compete with livestock for herbage, and in cropping areas where they may damage crops and pastures. In eastern Australian rangelands, migratory locusts predominantly damage native pastures (measured by the value of green plant material consumed). In cropping areas the value of locust damage is about equal between crops and pastures (Love and Riwoe 2005.) The most damaging is the Australian plague locust, which outbreaks somewhere in Australia every few years. Responsibility for operations to control
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locusts and grasshoppers rests with states, local government and landholders, but where migratory species cross state boundaries, operations are coordinated by the Australian Plague Locust Commission (APLC). The APLC is jointly funded by the Commonwealth (50%), NSW (32.5%), Victorian (10%), South Australian (5%) and Queensland (2.5%) governments. In addition to Australian plague locusts, the APLC also operates against spur-throated and migratory locusts when they are likely to cause damage in another state. Operations include forecasts based on remote sensing and ground surveys of breeding areas, and subsequent control of locusts in conjunction with state governments and local organisations. In WA, these functions are undertaken by the state government.
Arid zone grasshopper complex Peakesia hospita (Bolívar), Urnisa guttulosa Walker, Zabrala ceripes Sjöstedt and Yrrhapta
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a
Persistent Intermittent
b
Fig. 18.2. Breeding areas of the Australian plague locust and areas at risk during favourable seasons. (APLC)
limited grass response after drought, whole properties can be eaten out. Fig. 18.1. (a) Peakesia hospita adult (length 32 mm), and (b) Urnisa guttulosa adult (length 33 mm). (DCF Rentz)
spp. (Orthoptera: Acrididae) can occur in large numbers after heavy rain and cause substantial damage to pasture in some years. Distribution: Native to Australia. In some areas, like western Qld, high numbers of short winged grasshoppers such as Z. ceripes and Yrrhapta spp. can be locally common whenever it rains. In other regions, numbers can be high after drought. Pest status: Minor, restricted, irregular. Identification: A variety of small to moderatesized grasshoppers, some with wings. Others have only short wings as adults (Fig. 18.1). May be confused with: Several other species of grasshoppers may be confused with these pest species. Hosts: Most pasture plants. Life cycle: Little is known about the life cycle of these species. Most probably survive dry periods as eggs in the ground and then hatch out in large numbers after good rains in the arid zone. Risk conditions: After rain. Damage: These species usually hatch out after good rains when there is an abundance of grass and the damage is sustainable; however, when high numbers occur following localised rain and
Monitoring: Groups of small hoppers after rain indicate the possibility of damage. Action level: Control is usually not costeffective. Natural enemies: Little is known about the natural enemies of these species but they are probably similar to those of the Australian plague locust.
Australian plague locust Chortoicetes terminifera (Walker) Orthoptera: Acrididae Distribution: Native to Australia where it inhabits open plains of inland NSW, Qld, SA and WA; sometimes invades Vic. (Fig. 18.2). Pest status: Major, widespread, regular. Identification: Adult males are 25–30 mm long and females 30–45 mm long. Adults are most easily identified by the conspicuous black spot on the tip of the clear hindwings and the red shanks of the hindlegs (Fig. 18.3). There is no spur (throat peg) on the underside of the neck. The general body colour is usually brown, sometimes grey or green, often with a pale stripe down the middle of the back. Nymphs are usually brown but can be green. From above the thorax they have an ‘X’ mark, the hind part being more pronounced in the
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a
b
b
Fig. 18.3. (a) Adult Australian plague locust (female, length 35 mm), showing the black tip at the end of the hind wing and red shanks on the hind legs. (b) Adult female (top) and male (length 20 mm) showing light ‘X’ mark on top of the thorax. (DPI&F Qld: R. Elder)
early instars (Fig. 18.4). The rear leg femur has distinct bands which, when viewed from the side, are at 90 degrees across the femur. The rear tibia is dark except for a light band at the top. Bands of hoppers can be so dense that they can be seen from the air as an advancing front eating out the vegetation as they progress. May be confused with: Oedaleus australis (Saussure) (Fig. 18.8), Austroicetes cruciata (Saussure) (Fig. 18.16) and Heteropternis obscurella (Blanchard). Several grasshoppers have a hindwing with a smoky tip but only C. terminifera has the distinctive dark spot on the tip of the hind wing and red tibia on the hind legs. Host range: While the species shows a preference for gramineous (grass) crops such as pastures and related winter and summer cereal crops, under dry conditions or with dense populations, other plants may be eaten. Life cycle: In subtropical western Qld, it is warm enough for three generations between late winter and the next summer–autumn, but
Fig. 18.4. Australian plague locust nymphs: (a) firstinstar nymph (length 4 mm), and (b) fifth-instar nymph (length 20 mm). Arrows point to two arms of the ‘X’. (B. Lewis)
usually there are only two generations limited to the wetter summer. Eggs are laid in the soil, 3–10 cm deep, in pods of 30–50 eggs (Fig. 18.5). Females usually lay 2–3 egg pods. If the pasture is green, females lay their first pod 5–7 days after maturing and any subsequent egg pods are laid intervals of 5 days (summer) to 10 days (autumn). Eggs take about 2 weeks to develop in summer but development is arrested (quiescence) if conditions are too dry. Eggs laid in autumn enter diapause and hatch in spring. Nymphs take about 25–30 days to reach the adult stage in summer. The final moult to the winged adult is called fledging. The adult usually goes through three stages of development: growth, during which the wing muscles are developed and the exoskeleton hardens; fat accumulation; and oocyte (egg) development. Each stage can be suppressed if weather is dry. The growth stage usually lasts about a week. Fat is needed to fuel long-distance night migrations. Long distance migrations occur at night on the strong warm winds associated with rain-bearing fronts or lowpressure systems. Swarms can be displaced several hundred kilometres in a single night and such migrations make this locust a dangerous 487
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Fig. 18.5. The female bores a hole in the soil and deposits egg pods 3–10 cm deep. (Graphic Science: © Denis Crawford)
pest: cropping areas that had no locusts can be suddenly invaded, with substantial damage the result. Displacement of swarms may also occur through day flights; while such flights rarely exceed 10–20 km per day, they may occur for a week or more. Risk conditions: During most years, breeding on localised rains in the inland leads to small to moderate populations and only localised invasions of the agricultural zone during autumn. But in years of rain throughout the year, there can be three generations and the abundance of green grass results in large population increase and substantial invasions of the agricultural zone. Major breeding usually then follows in spring and in some years there can be significant return migrations to the inland allowing breeding to continue for several years. Damage: During outbreaks, nymphs form marching bands and adults form flying swarms that can cause substantial damage to pastures and cereal crops. Monitoring techniques: Locust populations are monitored and controlled by the APLC to reduce invasions of cropping areas. Warnings of outbreaks can be found on the APLC web site at www.daff.gov.au/animal-plant-health/locusts. When locusts are seen, they are reported to the APLC or local state authorities. Action level: At first evidence of bands or swarms. Chemical control: While direct damage to pastures may be minimal at times of heavy rain and abundant vegetation, the APLC initiates preventive control programs (Hunter 2004) to
Fig. 18.6. Aerial application of commercially formulated fungus Metarhizium anisopliae. (CSIRO: R. Milner)
reduce the size of invasions of cropping areas. Landholders treat small to moderate bands on the ground though large bands and most swarms require spraying by aircraft. Cultural control: Deep ploughing of egg-laying areas was once practised but the currently used minimum till ploughing is much less effective in destroying the eggs. Biological control: The naturally occurring fungus Metarhizium anisopliae var. acridium (formerly M. flavoviridae) (Deuteromycotina: Hyphomycetes) has been developed into a commercial product that can be applied by aircraft or ground equipment to locust bands (Fig. 18.6). There is high (> 80–90%) mortality in 8–18 days, depending on temperature, but the delay in mortality means locusts should be treated early as young nymphs to limit damage. Natural enemies: Eggs are parasitised by Scelio fulgidus Crawford (Hymenoptera: Scelionidae) (Fig. 18.7). Parasitism can be high in both rangelands and in agricultural zones, particularly during host outbreak recessions. In sub-coastal invasion areas, egg parasitism is supplemented by S. parvicornis Dodd. Larvae of Cryptomorpha flaviscutellaris Roberts (Diptera: Bombyliidae) have been listed as a predator of egg beds as have the larvae and adults of various beetles belonging to the Anthicidae, Melyridae, Carabidae and Tenebrionidae. Most egg predators appear to be opportunistic feeders and are rarely found. The bee fly, Trichopsidea oestracea Westwood (Diptera: Nemestrinidae) is a
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a
Fig. 18.7. Adult scelionid wasps emerging from a locust egg pod, which their immature stages have parasitised.
b
(NSWDPI: M .Hill)
widespread parasite of nymphs and adults, extending its range into semi-arid areas. Other fly parasitoids (mainly Blaesoxipha spp. Diptera: Sarcophagidae) are also sometimes found in nymphs and adults, mainly in higher rainfall areas. Various Dipteran and Hymenopteran predators of nymphs and adults are also known, but again they are usually opportunistic feeders and appear to have little effect on populations during outbreaks (Baker 1991).
Eastern plague grasshopper Oedaleus australis (Saussure) Orthoptera: Acrididae a
Fig. 18.9. Eastern plague grasshopper nymphs: (a) second instar (length 6 mm), and (b) fourth instar (length 11 mm). The face stripe on the nymphs distinguishes them from Australian plague locusts.
Distribution: Native to Australia, NSW, NT, Qld, southern SA, Tas. and eastern Vic. Pest status: Minor, restricted, irregular. May occur in large numbers and cause damage to pastures in drought years. Often associated with overgrazed or neglected farmland.
b
Fig. 18.8. The eastern plague grasshopper: (a) adult, 30 mm long, and (b) fifth-instar nymph (length 15 mm); arrows mark two arms of the ‘X’ on the thorax. (B. Lewis)
Identification: Adults are 22–31 mm long, brown or green, hindwings are lightly coloured from very pale to smoky yellow with a dark tip and a dark band along the centre. The pronotum with ‘X’ pattern (Fig. 18.8). The tibia on the hind legs are usually straw-coloured. The throat peg is absent. Nymphs are green or brown, have a prominent dark stripe from the antennae, across the eye, to the back of the head at a 45° angle (Figs 18.8 and 18.9). May be confused with: Both nymphs and adults are similar to the Australian plague locust. Adults can be distinguished by the markings on the hindwing. Nymphs can be distinguished by the face stripe. 489
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Host range: Grasses. Life cycle: Similar to the Australian plague locust. Risk conditions: Spring–summer. Drought typically precedes outbreaks of this species. Damage: Restricted to pastures. Monitoring techniques: Watch for groups of nymphs and adults in pastures, particularly during droughts. Action level: Usually not economically viable to treat, most outbreaks are short-lived. Natural enemies: Similar to Australian plague locust.
Migratory locust Locusta migratoria (Linnaeus) Orthoptera: Acrididae Distribution: Migratory locusts occur in Africa, Asia and Australia. In Australia, they are found in the tropical north but have become most common in the central highlands of Qld following clearing of the brigalow scrub and the introduction of African grasses such as rhodes grass and buffel grass (Fig. 18.10). Major outbreaks have occurred on the cracking clay areas of the central highlands of Qld from north of Clermont to south and east of Rolleston. Hopper bands and swarms are sometimes also found in central western Qld and along the east coast.
Persistent Intermittent
Fig. 18.11. Adult gregarious male migratory locust (length 45 mm). (DPI&F Qld: R. Elder)
Pest status: Minor, widespread, irregular. Identification: Adult migratory locusts are blockier in general appearance than the spurthroated locust, have hair on the under-side of the thorax and lack the spur between the front legs. The mandibles (jaws) are dark purple to black and there is a pale line stretching from the back of the eye. The shanks of the hindlegs are pink to straw-coloured and the insides of the legs are dark purple to mauve. Adults are between 35 and 65 mm long with the males being smaller than the females. The hindwings have no markings but may be faintly greenishyellow. General body markings vary depending on population density. Solitary individuals have green background colour overlain by darker markings. Gregarious adult females are grey– brown in colour while males may be strawyellow in colour (Fig. 18.11). Gregarious individuals of both sexes usually occur in swarms. Nymphs are similar to the adults, but are sexually immature and lack fully developed wings. Solitary nymphs are green, while gregarious forms are black and tan and usually occur in hopper bands (Fig. 18.12). Intermediate forms are dark grey to black. May be confused with: Spur-throated locusts, especially when seen flying as individuals, although migratory locusts are blockier in appearance. Host range: Pasture grasses, barley, maize, millet, oats, sorghum, sugarcane, triticale, and wheat.
Fig. 18.10. Breeding and outbreak areas of migratory locusts. (APLC)
Life cycle: Batches of eggs (egg pods) are laid in soil in a hole up to 90 mm deep by the
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a
The migratory locust has limited ability to survive dry conditions so in most years any increase in numbers during wet periods is balanced by mortality when it is dry, often in winter. Regular rains during winter, spring and summer lead to outbreaks with the rains allowing for maximum adult laying and good survival of eggs and nymphs. Risk conditions: Spring and summer following good winter rains.
b
Damage: Locusts eat grasses and graminaceous crops like sorghum and wheat. Monitoring: Inspect pastures weekly and note the locality of hopper bands and stage of development. Action level: In pastures, treat if hoppers are within a few kilometres of crops; in pastures far from crops, treat if hoppers are causing significant damage and stock feed is limited.
Fig. 18.12. (a) Migratory locust fifth-instar brown gregarious phase (length 35 mm), and (b) fifth-instar green, solitary phase (length 35 mm). The green form has a ridge along the back not present in the brown form. (B. Lewis)
female. The egg pods consist of about 60 eggs at the bottom of the tubular hole with a frothy plug from the top of the eggs up to the soil surface. All developmental stages may be present at one time throughout the year with up to four generations in good seasons. At 30–36°C, eggs take about 14 days to hatch, the nymph stage takes 25–30 days and adult females lay after 2 weeks. Development is slower in autumn and winter; for example, eggs take 59 days to hatch at the common autumn– winter temperature of 23°C. Eggs require moisture after being laid and obtain this from moist soil. Newly emerged nymphs require fresh green grass or green crop, while older nymphs can survive and develop on drier, less nutritious vegetation. When population densities are high, the locusts aggregate to form bands of nymphs and swarms of adults. Most outbreaks of the migratory locust originate in the central highlands and adjacent areas of Qld. Swarms seldom move more than a few kilometres a day.
Chemical control: In Qld, preventive control by landholders or by the government is conducted to limit movement onto crops. Control of hopper bands before they reach the winged adult stage is much more effective than later treatment of adult swarms. Biological control: A commercial product containing spores of the fungus Metarhizium anisopliae var. acridum has been developed for use against this species. See Australian plague locust for further details. Natural enemies: Similar to the Australian plague locust. Scelio fulgidus Crawford and S. gobar Walker (= S. bipartitus Kieffer) (Hymenoptera: Scelionidae) parasitise egg pods. Various dipteran parasitoids attack nymphs and adults.
Mulga grasshopper complex Adreppus sp. 6, Macrolobalia ocellata (Tepper) and Coryphistes interioris (Tepper) Orthoptera: Acrididae Distribution: Native to Australia, Qld. Pest status: Minor, irregular. Identification: Large, mottled, grey–green grasshoppers with an angled head, antennae 491
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Natural enemies: Unknown, but probably similar to the Australian plague locust and spurthroated locust.
Pyrgomorph grasshoppers Orthoptera: Pyrgomorphidae Blistered pyrgomorph, Monistria pustulifera (Walker) Fig. 18.13. Adreppus adult (length 50 mm).
thickened at the base and broad hindwings usually with a brown tinge (Fig. 18.13). May be confused with: Many other similar species exist within the same genera and within Euophistes and Microphistes. Hosts: Mulga (Acacia aneura), several species of Eucalyptus and other common semi-arid trees. Life cycle: Very little is known about these species and only one outbreak has been recorded. In 1992–1993, about one million hectares of mulga and gidgee in south-west Qld was severely defoliated, mainly by Adreppus species 6; however, other species of grasshoppers were also common, particularly M. ocellata and C. interioris, which caused significant damage by ringbarking small twigs. The outbreak was first reported in the Adavale-Quilpie area of Qld in April 1992 and then spread to other areas peaking in February and March 1993. The grasshoppers were inactive during the day and fed only at night.
Common pyrgomorph, M. discrepans (Walker) Distribution: Native to Australia, NSW, NT, Qld, SA, Vic., WA. M. discrepans is more widespread in the southern portion of the continent, while M. pustulifera extends further north. Pest status: Minor, irregular. Identification: Aposematically marked, colour variable. The ground colour is beige, brown, purplish or black, with spots ranging from yellow to scarlet. They can be long- or short-winged. Males are 15–33 mm long, and females are 26–38 mm long. The hindwings of long-winged forms are mottled with a purple tinge (Fig. 18.14). Younger nymphs are generally paler than adults. May be confused with: There are six other species in this genus but all are relatively easy to separate. Hosts: Recorded on six plant families with the genus Eremophila (desert fuchia) being the most frequently recorded. M. pustulifera also attacks a variety of garden plants including Buddleia, dahlias, honeysuckle and privet. Large numbers
Risk conditions: Summer–autumn. The 1992–1993 outbreak followed a sequence of exceptionally mild winters in south-west Qld. Damage: Severe defoliation and ringbarking resulting in death of some shrubs and trees. The level of damage to mulga was of economic concern to graziers as the foliage is used as sheep fodder during droughts; however, this loss was compensated in some areas by a substantial increase in grass growth due to the thinning of the shrub layer. Monitoring: Monitor for population increase if a sequence of mild winters occurs. Action level: Not economically viable to treat. Chemical control: Not viable to treat.
Fig. 18.14. Adult common pyrgomorph, winged form (length 27 mm). (D. Rentz)
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of M. discrepans have been collected from Westringia fructicosa in NSW. Life cycle: The following account of the biology of M. discrepans is taken from laboratory studies conducted by Allsopp (1977). Females lay up to four egg pods in the soil with an average of about 32 eggs per pod. Eggs enter an obligate diapause during winter and require a period of chilling to break diapause. Nymphs undergo four (males) or five (females) instars and take about 7–9 weeks to develop at 27.5°C constant. At this temperature, females take another 47 days after fledging before laying eggs. Risk period: Summer to autumn. Damage: In the 1970s, both species defoliated large stands of turpentine (Eremophila sturtii) and green turkey bush (E. gilesii) in the Charleville area, Qld. M. discrepans was considered as a potential biocontrol agent for E. gilesii, which is a woody weed of grazing lands in south-west Qld. Large numbers of Monistria (probably M. discrepans) were found attacking garden plants and Eremophila near Mildura, Vic., in autumn 2002 and summer 2003. Monitoring: Monitor for population increase if a sequence of mild winters occurs. Action level: Usually not economically viable to treat except in gardens. Chemical control: Usually not viable to treat. Biological control: The fungus Metarhizium anisopliae var. acridum was successfully tested against this genus in 2002 and 2003. Natural enemies: Probably similar to other grasshoppers and locusts.
Southern pyrgomorph, Monistria concinna (Walker) Orthoptera: Pyrgomorphidae Distribution: Native to Australia, NSW, Tas. and Vic. Pest status: Minor, irregular. Identification: Aposematically marked, colour variable, eight geographical races recognised, each distinct in colour and morphology. The ground colour dull green to olive, brown to purple, or nearly black with yellow spots. Spots may be abundant or few, or form into longitudinal stripes. It is short-winged
Fig. 18.15. Southern pyrgomorph adult (length 40 mm). Only a short-winged form is known. (D. Rentz)
(Fig. 18.15). Males are about 20 mm long and females are about 40 mm long (Rentz et al. 2003). May be confused with: There are six other species in this genus but all are relatively easy to separate. Hosts: Found on many types of herbaceous and shrub-type plants belonging to six families. Life cycle: Adults and juveniles are found throughout the year. Risk period: Summer. Damage: When populations become dense, they can be of economic concern feeding on clover and lucerne. Monitoring: Monitor for population increase if a sequence of mild winters occurs. Action level: None recommended. Chemical control: Not cost-effective. Natural enemies: Unknown, but probably similar to other grasshoppers and locusts.
Small plague grasshopper Austroicetes cruciata (Saussure) Orthoptera: Acrididae Distribution: Australian native. Inland southern NSW, SA and WA; sometimes north-west Vic. Occurs in these states on southern fringe of the semi-arid zone associated with open grasslands, open mallee and pastures. Pest status: Major, restricted, irregular. Outbreaks are usually localised and irregular but damage can be severe when they develop in cropping districts. The low migratory capability of adults and the slow rate of increase due to the 1-year life cycle usually prevent outbreaks from 493
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a
a
b b
Fig. 18.17. The small plague grasshopper: (a) fourth instar (length 9 mm); (b) fifth instar (length 14 mm). (B. Lewis)
Fig. 18.16. The small plague grasshopper: (a) adult female (length 25 mm); (b) adult male (length 14 mm). Arrows indicate position of the three dark bands on the rear femur. (B. Lewis)
becoming widespread. Outbreaks occur mainly in SA, although damage can occur in NSW and WA where land use changes appear to have reduced the pest status of this species. Identification: The colour and pattern are highly variable, with two relatively distinct races, the eastern race and the western race, both exhibiting phases. The rear femur has distinct bands, which, appear as three dark and three light bands when viewed from above (Fig. 18.16). When viewed from the side, the bands are at an oblique angle, often as a triangle, across the femur. The rear tibia are completely dark in early instars but become light in later ones. Antennae are short, being approximately one to one and a half times the length of the eye. Adults are 14– 25 mm, usually brown to green, but swarming males turn bright yellow. The top of the thorax has a faint ‘X’ mark, with the rear half more prominent. The rear tibia vary from pale straw to pink and orange. The hindwing is clear with a dark margin on the forward edge only. Nymphs (Fig. 18.17) vary in colour from brown to green, with a distinct ‘X’ mark on the thorax when viewed from above.
May be confused with: Australian plague locust (C. terminifera) and other species of Austroicetes. Nymphs can be distinguished from Australian plague locusts by more oblique banding on the hind femur, shorter antennae (relative to the eyes) and by the absence of a light band at the top of the hind tibia that is typical of the Australian plague locust (Fig. 18.4). Adults (except yellow phase swarming males) may also be confused with Australian plague locusts but are readily distinguished by their smaller size, the lack of a distinct spot on the tip of the hindwing and by the colour of the hind leg tibia. There are eight other described species in the genus and it is most likely to be confused with A. pusilla (Fig. 18.18), but A. cruciata is the only member of this genus that commonly forms into bands or swarms. Host range: Most pastures and winter cereal crops (wheat, barley and oats). Life cycle: Single generation per year. Eggs are laid in late spring and early summer, and enter diapause which is broken by cold temperatures. The post-diapause egg is not resistant to desiccation but usually winter rain ensures survival and hatching in late winter–early spring. This species is mainly distributed where winter–spring rainfall is reliable enough to ensure the high survival of nymphs. Nymphs
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a
b
Fig. 18.18. A. pusilla: (a) fourth instar (length 9 mm); (b) fifth instar (length 12 mm). (B. Lewis)
complete development in approximately 6 weeks and fledge in mid-spring. Egg-laying begins in late spring and continues for about 5 weeks, with adults persisting until mid-summer. Risk period: Spring, following moderately dry years. Damage: Nymphs form marching bands. Adults form loose flying swarms that can cause substantial damage to pastures and cereal crops. Monitoring techniques: Watch for groups of nymphs in pastures; later-adult swarms will be very obvious.
Fig. 18.19. Other Austroicetes spp. may be found in semi-arid grasslands but they do not form bands or swarms. (B. Lewis)
Natural enemies: Eggs are parasitised by Scelio chortoicetes Froggatt and S. parvicornis Dodd (Hymenoptera: Scelionidae). Larvae of Cryptomorpha flaviscutellaris Roberts (Diptera: Bombyliidae) have been recorded as predators of egg beds. Various species of Diptera parasitise nymphs and adults, similar to those found attacking C. terminifera and other species of grasshopper. Outbreak collapse has been associated with periods of high rainfall, which may allow bacterial and fungal pathogens to develop.
Other species of Austroicetes, Austroicetes vulgaris (Sjöstedt), Austroicetes nullaborensis Key Distribution and pest status: Austroicetes vulgaris may occasionally cause damage to pasture in conjunction with other species of grasshopper. It is found in southern NSW, SA, Tas., Vic., and southern WA. Austroicetes nullaborensis occasionally swarms in regions of
Action level: At first evidence of bands or swarms. Chemical control: Treat bands and swarms directly; swarms require spraying by aircraft. Cultural control: Deep-ploughing of egg-laying areas was once practised but the currently used minimum-till ploughing is much less effective in destroying the eggs. Biological control: A commercial product containing spores of the fungus Metarhizium anisopliae var. acridum has been developed for use against this species. See Australian plague locust for further details.
Fig. 18.20. Adult Austroicetes nullaborensis (20 mm long) (top) and A. cruciata (25 mm long) (bottom). (PIRSA: P.R. Birks)
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South Australia but is not regarded as a serious pest.
some species (e.g. M. modesta) males are shortwinged.
Identification and confusion with other species: Similar to A. cruciata and other members of the genus (Fig. 18.19).
May be confused with: Unlikely to be confused with other species of grasshopper in spinifex areas.
Host range: Most pastures and winter cereal crops (wheat, barley and oats).
Hosts: Spinifex (Triodia spp.) and other grasses.
Life cycle: Not well studied but probably similar to A. cruciata. Risk period: Spring and summer. Damage: Occasional pests of pasture but rarely of economic importance. Monitoring techniques: Watch for groups of nymphs in pastures; later adult swarms will be very obvious. Action level: Usually not economically viable to treat. Natural enemies: See Austroicetes cruciata.
Spinifex grasshoppers Various species of Macrotona Brunner (Orthoptera: Acrididae), with M. modesta (Sjöstedt) often being the commonest. Distribution: Native to Australia, NSW, NT, Qld, SA, and WA, associated with many habitats in the arid and semi-arid interior particularly in areas of spinifex grass in Qld. Pest status: Minor, sporadic pest in Qld. Identification: Small to moderate (12–40 mm), slender grasshoppers with a sloping head when viewed in profile (Fig. 18.21), often brilliantly marked. Adults are usually fully winged, but in
Life cycle: Little is known about the life history of these species. Risk period: Summer, after high rainfall, can persist in high numbers over several years. Damage: During outbreaks, adults and nymphs can cause substantial damage to spinifex plants and even completely destroy tussocks. They eat part or all of the leaves and appear to chew through aerial roots or runners. Where only part of the tussock is eaten, the remainder wilts giving the impression that the plant is drying out. Affected tussocks turn light green to brown in colour and only stand at less than half the height of healthy ones. Monitoring: Groups of small hoppers after rain indicate an infestation is developing. Action level: Usually not economically viable to treat. Cultural control: Some landholders have practised burning spinifex tussocks to reduce grasshopper numbers, but this is often impractical when large areas are infested. Also, if spinifex is burnt, some soil moisture is needed to protect the root system from being destroyed. Natural enemies: Unknown.
Spur-throated locust Austracris (Nomadacris) guttulosa (Walker) Orthoptera: Acrididae Distribution: Native to Australia and Papua New Guinea, although similar species are found in Africa and Asia. They are found throughout northern Australia (Fig. 18.22), although the most commonly infested area is the cracking clay plains south of the Gulf of Carpentaria, where swarms commonly form during autumn and early winter.
Fig. 18.21. Adult spinifex grasshopper, Macrotona modesta (length 20 mm). (D. Rentz)
Pest status: Minor in pastures, major in crops, widespread distribution, irregular occurrence. Some treatments are required every few years in
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Intermittent Persistent
Fig. 18.24. Spur-throated locust: fifth instar (length 50 mm) showing black stripe along its back. (B. Lewis)
to light brown as they approach the adult (winged) stage. Fig. 18.22. Occurrence of spur-throated locusts. (APLC)
the central highlands of Qld but major outbreaks only occur every 10–20 years. Identification: Adults are between 50 and 80 mm long. All stages have a conspicuous spur between the front legs (Fig. 18.23). The hindwings are colourless or have a bluish tinge. The shanks of the hind legs have dark-tipped white spines. At rest, the insect has a white– cream stripe down the middle of the dorsal (upper) surface of the head, through the thorax and overlapping wings. Nymphs are pale green on hatching and later develop a black stripe down the middle of their backs (Fig. 18.24). The green colour may change
Fig. 18.23. Under-side of the head and thorax showing the ‘spur’ in the ‘throat’ between the two front legs. (DPI&F Qld: R. Elder)
May be confused with: Some solitary species of grasshopper also have a spur between the front legs, but are usually much larger (e.g. the giant grasshopper, Valanga irregularis (Walker)). There are also some grasshoppers that are pale green like mid-instar spur-throated locusts, but grasshoppers of the same size are older and have wing buds (mid-instar spur-throated locusts lack wing buds). Adults can be confused with migratory locusts, which are, however, much blockier in appearance. Host range: Pastures and most crops may be attacked, but summer crops are at greatest risk. Some of the hosts include citrus, macadamias, soy bean, corn, green wheat, young sunflower, cotton, sorghum, mung beans, palm trees, eucalyptus trees, and turkey bush. Life cycle: One generation per year. Adults live 10–12 months and survive the dry season by feeding on the leaves of shrubs and trees. They do not mature sexually until the onset of the summer wet season when day-lengths are greater than 13 hours, temperatures are greater than 32°C and humidity is greater than 60%. Adults develop their eggs by feeding on green vegetation abundant during the first half of the wet season (October–January). Spur-throated locusts do not lay into egg beds like other locust species but lay throughout grazing country as well as cropping and fallow areas. Eggs are laid in pods up to 80 mm deep in moist soil. The pods consist of up to 150 eggs at the bottom of the tubular hole with a frothy plug to the soil surface. After hatching, they normally do not form hopper bands, although there can be some marching behaviour at the height of plagues. 497
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During the summer wet season when maximum temperatures are 30–36°C, the approximate duration of the life history stages are: egg 18 days, and hopper 65 days, with adults not maturing their eggs until the beginning of the next wet season. Risk period: Some treatment is required every few years in the central highlands of Qld, but there are major outbreaks only every 10–20 years. Rainfall of > 40 mm early in the wet season followed by two or more rains of > 40 mm at intervals of 6 weeks or less favours egg-laying and survival of nymphs. There is risk of an outbreak when this sequence of rains is widespread and occurs in two successive years. West to east weather fronts during September– January may move large numbers of adults to cropping areas. Damage: The locusts mature rapidly on arrival in cropping areas and lay; the resultant hopper populations can cause serious crop losses. Monitoring: During outbreak years, landholders should inspect pastures weekly for large numbers of hoppers or adults. Action level: Action may be warranted if large populations are present and are causing significant damage when food for stock is limited. Chemical control: Landholders sometimes control hoppers when they are dense, although the normal lack of hopper bands means that most landholders do not treat until the adult stage when crops are directly threatened. The Qld government treats the dense swarms that form in trees over the winter, particularly when swarms are in or near the agricultural zone. Biological control: A commercial product containing spores of the fungus Metarhizium anisopliae var. acridum has been developed for use against this species. See Australian plague locust for further details. Natural enemies: Similar to Australian plague locust. Scelio gobar Walker (= S. bipartitus Kieffer) (Hymenoptera: Scelionidae) parasitises egg pods at low levels. Various dipteran parasitoids attack nymphs and overwintering adults, mainly Blaesoxipha spp. (Sarcophagidae) and Phorocerocoma postulans (Walker) (Tachinidae). High rates of parasitism by
dipteran parasitoids has been implicated in the short persistence of outbreaks of this species in eastern invasion areas of Qld. One mammal and four bird predators have also been recorded, but none significantly affected populations during outbreaks. The strain of fungus Metarhizium anisopliae var acridum developed for the control of Australian locusts and grasshoppers was originally isolated from a spur-throated locust.
Wingless grasshoppers Orthoptera: Acrididae Phaulacridium vittatum (Sjöstedt), and robust phaulacridium, P. crassum Key Distribution: Native to Australia. The wingless grasshopper is widely distributed in moist (annual rainfall above 500 mm) areas of NSW, SA, Tas., Vic., southern Qld and southern WA. Robust phaulacridium is recorded from parts of south-west WA. Pest status: Wingless grasshoppers: major, widespread, irregular. Robust phaulacridium: minor, restricted to coastal areas, irregular. Identification: Colour and pattern are highly variable (Figs 18.25 and 18.26). Nymphs are pale pink on hatching but quickly darken to black (Fig. 18.25). Later instars are grey–brown, and the pronotum is flat on the dorsal surface (Fig. 18.26). Adult females are about 20 mm long. The hind femur has a black mark at mid-point and the hind tibia is orange (Fig. 18.26). Males are smaller than females; most have short wings but a variable proportion have functional long wings. Wing veins on short-winged morphs run laterally, in line with the body, as opposed to radiating outwards as in late-instar locust nymphs. A small proportion of adults may have a white lateral body stripes. May be confused with: Early instar nymphs may be confused with Brachyexarna lobipennis Sjöstedt (Fig. 18.29) and Austroicetes pusilla (Walker) (Fig. 18.18) as these can hatch at the same time and are commonly associated with wingless grasshoppers during outbreaks. Adults and nymphs of Phaulacridium vittatum may be confused with P. crassum in WA, but both species rarely occur at the same locality. P. crassum is larger than P. vittatum, the body surface is
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a
b
Fig. 18.27. Wingless grasshopper adult female (20 mm long) (top) and male (bottom). Both these specimens have short wings (‘wingless’) but some individuals have functional wings. (QPI&F Qld: R. Elder)
smoother and shinier, and their colour is duller and less variable than P. vittatum (Key 1992). Host range: A wide range of native and introduced forbs, field crops, horticultural crops and trees. Grasses are not preferred. Fig. 18.25. Wingless grasshopper nymphs: (a) first instar (length 2–3 mm), and (b) second instar (length 5 mm).
a
Life cycle: One generation per year. Eggs are laid in the soil in pods at a depth of approximately 2 cm. Egg pods are 1.5–2.0 cm long, tear-shaped and usually contain 10 to 16 eggs, each 4.5 mm long (Fig. 18.28). Females lay at least three times, but under favourable conditions may lay more than six times. Eggs enter diapause during the winter and hatch in mid–late spring (October– November). Nymphs develop through five instars in approximately 7 weeks and fledge between mid-December and early January.
b
Fig. 18.26. Wingless grasshopper nymphs: (a) third instar (length 8 mm), and (b) fourth instar (length 13 mm). (B. Lewis)
Fig. 18.28. Wingless grasshopper eggs (length 4.5 mm) laid in pods. (SARDI: P.T. Bailey)
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Oviposition commences by February and continues until April. Early instars do not move far from hatching sites. Late-instar nymphs and short-winged adults may exhibit gregarious behaviour and march in streaming bands. Dispersal is usually downhill, with populations concentrating on moist flats and springs. Following rainfall, populations may move back to suitable egg-laying sites on the crest of ridges. Long-winged adults are extremely active and fly at the first opportunity following fledging. Flights are often less than half a kilometre but may result in substantial downwind displacement of populations over time. Risk conditions: Most damage to pastures occurs in mid-summer and autumn, following moderately dry years. Wingless grasshoppers are severe pests of improved pasture with outbreaks occurring every 5 years or so on the western slopes and every 8–10 years in the tablelands of NSW and mid-western Vic. In some districts of SA, outbreaks appear to occur in cycles lasting 6 or more years. Sporadic outbreaks also occur in Qld, Tas. and WA. Localised outbreaks often occur after changes in land use when new pastures or crops are being established. The abundance of favoured food plants in improved pastures and the absence of natural enemies following the disruptive change in land use, leads to significant wingless grasshopper infestations after colonisation. Widespread outbreaks occur during seasons of below-average spring rainfall, attributable to increased nymphal survival. Survival of nymphs is favoured by relatively abundant growth of broadleafed host plants, higher temperatures favouring development and adverse conditions for natural enemies, particularly nematodes. Damage: Preferential feeding on the legume component of improved pastures may damage newly established areas so severely that resowing is necessary. Following mid-season drying-off of vegetation and/or exhaustion of food sources in pastures, dispersal of grasshoppers may result in the invasion and severe defoliation of pasture legumes, fodder crops (particularly lucerne), horticultural crops, trees and gardens. New pine plantations are also susceptible to attack. Wingless grasshoppers directly compete with sheep during late summer in SA and Vic.; densities of 30 per square metre
eat a similar biomass of clover as 2.6 dry sheep equivalents (Bailey et al. 1994). Monitoring techniques: Watch for groups of nymphs in pastures; later-flying adults will be very obvious. Action level: At first evidence of gregarious populations of nymphs in high-value pastures. Concentrated infestations in the vicinity of crops should be controlled with barrier treatments. Treatment of low-value pastures is usually not cost-effective. Chemical control: Ground-spraying is practised to protect valuable pastures and irrigated fodder crops, particularly when stock feed is limiting. Barrier treatment of lucerne and horticultural crops such as vineyards, orchards and gardens may be necessary to prevent invasion. Baits of wheat bran mixed with insecticide can also be used to control concentrations. Baits are best applied in a band of about 2 m wide to bare ground but they are not very effective in dense pasture. Re-invasion of treated areas often occurs during regional outbreaks, particularly by long-winged adults, and several applications of baits or sprays may be necessary in a single season. Cultural control: Cultural control is difficult on pastures, but maintaining a dense grassy pasture without bare areas where grasshoppers can lay eggs can help to keep numbers in check. Control of weeds that act as wingless grasshopper food sources can also help to reduce the intensity of outbreaks. Reafforestation of denuded hillsides that act as source areas of wingless grasshoppers has also been proposed as a long-term solution to decrease outbreaks in some areas; however, this can be difficult as young trees are very susceptible to attack. Biological control: A commercial product containing spores of the fungus Metarhizium anisopliae var. acridum has been developed for use against this species. See Australian plague locust for further details. An exotic protozoan disease, Nosema locustae, was tested against wingless grasshoppers during the late 1970s but was of limited success (Moulden and D’Antuono 1984). Natural enemies: Up to 30% of the eggs may be parasitised by Scelio improcerus Dodd
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and S. parvicornis Dodd (Hymenoptera: Scelionidae). Nymphs and adults are parasitised by several dipterans (e.g. Trichopsidea oestracea Westwood, Nemestridae) similar to the ones attacking the Australian plague locust. Nematodes (Nematoda: Mermithidae) are important regulators of wingless grasshopper populations in NSW and are often the cause of the collapse of outbreaks. The two nematode genera most commonly found are Amphimermis and Mermis; both are highly dependant on rainfall for successful parasitism. Amphimermis spp. lay their eggs in the soil and on hatching, the free-swimming larvae move to the surface where they locate and penetrate a host. Mermis spp. lay their eggs on vegetation and infect the host when the grasshopper accidentally eats it (Baker 1995). Birds (e.g. crows, magpies and ibis) prey on wingless grasshoppers, and guinea fowls may be useful in gardens. Cannibalistic feeding in wingless grasshopper adults transmits a natural amoebic disease, caused by Malameba locustae.
Other pest species of wingless grasshoppers Yellow-bellied grasshoppers, Praxibulus insolens Rehn, Praxibulus eurobodallae Key, Praxibulus duplex Rehn, Praxibulus stali Key Stripewinged meadow grasshopper (= mountain brown), Brachyexarna lobipennis Sjöstedt Robust fipurga, Fipurga crassa Sjöstedt Orthoptera: Acrididae
Fig. 18.29. Stripewinged meadow grasshopper, Brachyexarna lobipennis (length 12 mm). (D. Rentz)
winged but may produce long-winged morphs when in high densities. Praxibulus spp. (Fig. 18.30) are generally brown–green, often with a yellow under-side and short wings. Fipurga crassa is a robust-looking, brown–green, short-winged grasshopper with hind tibia variable in colour but often blue–green with purple feet. May be confused with: Early instar nymphs may be confused with Phaulacridium vittatum and other wingless species that can occur together during outbreaks. Several species of Praxibulus exist and they are notoriously difficult to distinguish, requiring detailed study of genitalia for certain identification (Rentz et al. 2003). Host range: Various native grasses of short growth habit, Praxibulus prefer rat’s tail fescue (Volpa bromoides). Life cycle: Similar to P. vittatum. Usually univoltine, but a partial second generation can
Distribution: Moist (annual rainfall above 500 mm) areas of south-east Australia (NSW, SA, Vic.). Pest status: Sporadic. These grasshoppers may occur in large numbers, often as a suite of wingless species, sometimes in association with the wingless grasshopper Phaulacridium vittatum. These grasshoppers cause significant damage to pastures in certain years. Identification: Colour and pattern are variable. Generally small (< 20 mm) with a throat peg and often a broad, flattened pronotum. Brachyexarna lobipennis is brown (Fig. 18.29), usually short-
Fig. 18.30. Praxibulus insolens may be found with other species of wingless grasshoppers. (D. Rentz)
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occur during exceptionally warm springs. Populations of Praxibulus are usually ephemeral as they are susceptible to fungal infections that may decimate populations after rain.
a
Risk period: Spring–summer. Drought typically precedes outbreaks of these species. Damage: Restricted to pastures. Unlike P. vittatum, they are not known for invading fodder crops or gardens. Monitoring techniques: Watch for groups of nymphs and adults in pastures, particularly following droughts.
b
Action level: Usually not economically viable to treat, most outbreaks are short-lived. Natural enemies: Similar to P. vittatum; however, Praxibulus spp. appear to be less susceptible to parasitism by nematodes but more susceptible to epizootics such as the fungus Entomophaga grylli. The microsporidian disease Nosema locustae has been recorded in populations of Praxibulus in the northern tablelands of NSW. Various species of Scelio (Hymenoptera: Scelionidae) parasitise egg pods. Praxibulus nymphs and adults are prone to attack by parasitic flies, which results in grasshoppers having an intersex appearance and renders them sterile (Rentz et al. 2003).
Yellow-winged locust Gastrimargus musicus (Fabricius) Orthoptera: Acrididae Distribution: The yellow-winged locust is found throughout coastal and sub-coastal eastern, western and northern Australia in districts with an annual rainfall above 500 mm. Pest status: Minor in pastures, major in crops, widespread and irregular occurrence. Identification: A moderate-sized locust (35– 50 mm), easy to identify from the distinctive yellow colour of the hindwings. The hindwings are bright yellow at the base, with a broad black curved band, and a transparent and colourless wing tip (Fig. 18.32). Males make a distinctive clicking sound in flight. Colour and body shape vary with population density. Solitary individuals have predominantly green and dark brown colouration, while gregarious individuals are predominantly dull brown in colour.
Fig. 18.31. The yellow-winged locust: (a) second instar (length 9 mm), and (b) third instar (length 14 mm). (B. Lewis)
Nymphs are similar to adults in colour and shape but lack fully formed wings (Fig. 18.31). There are gregarious and solitary forms. The tip of the mandibles in large hoppers is bright red. May be confused with: Nymphs are sometimes confused with migratory locusts, but yellow-winged locust nymphs have a distinct ridge behind their head (Fig. 18.31) even when in bands. Nymphs of the eastern plague locust (Oedaleus australis) also have the ridge but they are smaller. Adults of O. australis are also smaller than yellow-winged adult locusts and have a hindwing that is pale yellow with a dark band. Host range: Gramineous crops such as pasture grasses, oats, wheat, barley, triticale sugarcane, sorghum, maize and millet. Life cycle: Batches of eggs (egg pods) are laid in soil in a hole up to 85 mm deep drilled by the female. Eggs are laid in a pod at the bottom of the tubular hole with a frothy plug up to the soil surface from which hatched hoppers escape by moving up its centre. At normal
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a
Fig. 18.33. A band of yellow-winged locusts. (DPI&F Qld: R.
b
Elder)
Risk period: Damaging outbreaks occur many years apart. Damage: Both nymphs and adults may eat all the grass component of pastures. Monitoring: During outbreak years, landholders should inspect pastures weekly for large numbers of hoppers or adults. Fig. 18.32. Yellow-winged locust adult (length 38 mm) in (a) green solitary phase colouration (D.Rentz) , and in (b) gregarious colour (45 mm long). (DPI&F Qld: R. Elder)
summer temperatures (30–36°C) the minimum duration of the life history stages is: egg about 17 days, hopper 30–40 days, and adults lay after about 2 weeks. With regular rainfall there are two to three generations per year. Hoppers have five instars. The species probably survives the winter in the egg stage. When population densities are high, hopper bands (Fig. 18.33) and adult swarms form. Eggs are laid in beds with up to 3000 egg pods per square metre. Bands and swarms move only short distances, although they can invade adjacent crops.
Action level: Action may be warranted if large populations are present and are causing significant damage when food for stock is limited. Chemical control: Usually only warranted to protect crops. Natural enemies: The most effective natural enemy is thought to be the wasp egg parasitoid Scelio gobar Walker (= S. bipartitus Kieffer) (Hymenoptera: Scelionidae). Other natural enemies include beetle egg predators, Dermestes ater (Coleoptera: Dermestidae), and predators of the hoppers including robber flies Bathypogon spp. (Diptera: Asilidae), frogs (Limnodynastes peronii) and cane toads (Bufo marinus), birds (ibis and crows). The egg parasitoids are believed the most important in reducing populations.
Sources of information Allsopp, P.G. (1977). Biology and capacity for increase of Monistria discrepans (Walker) (Orthoptera: Pyrgomorphidae) in the laboratory. Journal of the Australian Entomological Society 16: 207–213. Bailey, P., Frensham, A., Hincks, A. and Newton, M. (1994). Competition for herbage by Phaulacridium vittatum (Sjöstedt) (Orthoptera: Acrididae) and sheep during summer drought. Journal of the Australian Entomological Society 33: 175–179.
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Baker, G.L (1991). Parasites of locusts and grasshoppers. Agfact AE2 (second edition). NSW Agriculture. Baker, G.L. (1995). Wingless Grasshopper. Agfact AE1 (third edition). NSW Agriculture. Hunter, D.M. (2004). Advances in the control of locusts (Orthoptera: Acrididae) in eastern Australia: from crop protection to preventive control. Journal of the Australian Entomological Society 43: 293–303. Key, K. (1992). Taxonomy of the genus Phaulacridium and a related new Genus (Orthoptera: Acrididae). Invertebrate Taxonomy 6: 197–243. Love, G. and Riwoe, D. (2005). Economic Costs and Benefits of Locust Control in Eastern Australia. ABARE Report. Canberra. Moulden, J.H. and D’Antuono, M.F. (1984). Evaluation of Nosema locustae for control of wingless grasshoppers (Phaulacridium spp.) in Western Australia. In: Proceedings of the 4th Australian Applied Entomological Research Conference, Adelaide. pp. 387–393. Rentz, D.C.F., Lewis, R.C., Su, Y.N. and Upton, M.S. (2003). A Guide to Australian Grasshoppers and Locusts. Natural History Publications (Borneo), 419 pp.
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GLOSSARY
Acaricide: Pesticide used to control mites. Action threshold: An estimate of the level of pest abundance (e.g. number per square metre, number per plant or leaf) or signs of damage at which intervention is likely to be cost-effective. ‘Bestavailable’ estimates used in this book vary in reliability between those based on careful field experiments and those based on informed guesses. All thresholds are likely to vary between plant varieties, between geographical areas with markets, with changes in commodity price and with costs of control. Alate aphids: Winged. Antennal club of adult beetles: The thickened terminal end of the antenna. Aposematic colouration: Warning colouration, often to advise predators that the wearer is unpalatable. Apterous aphids: Wingless. APVMA: Australian Pesticides and Veterinary Medicines Authority. Operates the national system which evaluates, registers and regulates agricultural and veterinary chemicals. Current pesticide registrations are detailed at the APVMA website. Arthropods (Phylum Arthropoda): Invertebrates with a jointed external skeleton. The group includes crustaceans, millipedes (four legs on most segments), centipedes (two legs on most segments), spiders and mites (eight legs), and insects (six legs).
preferences or biotypes that may be resistant and non-resistant to a pesticide. Boll: Developing cotton fruit. Bolls have a hard outer covering protecting the developing lint and seeds inside. Bollgard II®: Transgenic cotton varieties containing two genes from Bacillus thuringiensis, one of which expresses the Cry1Ac protein, the other the Cry2Ab protein, both of which are toxic to Helicoverpa species and other noctuids (rough boll worm, cotton tipworm, cotton loopers). Break (of season in winter rainfall areas): Autumnal rains, which may allow crops to be sown. Bt: Bacillus thuringiensis Berliner, a bacterium that occurs naturally in some soils which may form crystals that are toxic to some species of arthropods. Commercial formulations of these act as stomach poisons when ingested by insects. Broad-spectrum insecticides: Non-selective insecticides that kill a wide range of insect groups including both target and non-target insects and mites. Capitulum (of a thistle): Head of a thistle. Carbamate insecticides: Inhibit acetylcholine esterase at nerve synapses. Cauda (of aphids): A single pipe at the tip of the tail through which honeydew is voided.
Awn: Outer cover of cereal seed. Beat sheet: See monitoring methods. Biocide: Biological agent applied in the same way as an insecticide. Typically, mass-produced parasitoids, diseases or bacterial products are released on crops for immediate reduction of a target pest. Biotype: Variant populations of the same species. Some biotypes may be distinguishable by colour, size or some other observable character. Other biotypes may not be morphologically distinguishable, such as those with differing host
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The relative lengths of the cauda and siphunculi at the tip of an aphid’s abdomen are characters often used to help distinguish between aphid species. (Graphic Science: © Denis Crawford)
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Chorium: Part of the front wing of the group of true bugs that have a front wing divided into hard and soft parts (Heteroptera). The chorium is the larger, hard part of the wing. Clean fallow: Bare ground kept free of weeds before a crop is planted. Climex: Program which predicts distribution and abundance of pests from climate data (Hearn Ltd). Common name: Common names of insects in this book follow, with few exceptions, those listed in Naumann, I. (1993). CSIRO Handbook of Australian Insect Names, updated on the website: www.ento. csiro.au/aicn CottonLOGIC: Decision support tool that runs on Windows-based computers and handheld computers. Accepts pest sampling information and evaluates pest numbers against thresholds. Models predict Helicoverpa abundance for the following 3 days and predict potential yield loss from mites. The program also acts as a repository for information on operations (e.g. sprays applied, when and at what price). Coxa (plural coxae): The first segment of the leg next to the arthropod body, to which it is articulated. Cracked grain baits: Used for field control of adults of a number of soil insects. Cracked wheat, sorghum or bran is impregnated with insecticide and broadcast over the soil surface after sowing. Addition of a vegetable oil may improve bait attractancy. Ingredients to make your own insecticidal grain bait include: • 100 mL chlorypyrifos (500 g L–1) EC • 125 mL crop or vegetable oil • 2.5 kg of cracked wheat, sorghum or standard pellets. Crawler: First-instar mobile stage of scales and whiteflies (Hemiptera). Cremastal spines or tail spines, of some Lepidoptera pupae: Small, fine projections from the last abdominal segment. The shape may be used for identification (Fig. 3.22). Crop dilution of insecticides: Rapid growth of foliage after insecticide application may reduce the effective insecticide dosage on plant surfaces. Crown (of plant): The central area of a rosette from which new leaves are produced.
Cry1Ac and Cry IIAb: See Bollgard II®. Cultural control: Modification of agronomic practices to reduce the incidence of pests in a crop; for example, by changing the dates of sowing or harvest or fallowing to remove weed hosts of pests. CV: Cultivar. Determinate: Crop cultivar or variety with synchronous flowering and podding; that is, where these reproductive stages do not overlap. See also Indeterminate. Detritivore: An organism that feeds on detritus (decaying organic matter). Diapause: A physiological state of arrested metabolism, growth, and development programmed to occur during a particular stage of development when triggered by environmental changes such as temperature and/or day length. Diapause may be triggered during an earlier stage of development (e.g. egg) to occur in a later stage (e.g. pupa). Diapause enables the insect to survive regular unfavourable environmental conditions (e.g. winter). Dispersive clay soils: Clay soils that disperse to produce muddy water when inundated for rice production. Dorsal surface: The upper surface of an insect. Drill-sowing: Crop sowing through the use of a seed drill (an alternative method to aerial sowing in rice production). Elytra: The thickened, hardened forewings that protect the membranous hindwings in beetles. These forewings are not used for flight. Endophyte: A fungus that infects the leaf and crown of ryegrasses. The fungus secretes alkaloid compounds which appear to act as natural ‘insecticides’ within the plant. Entomopathogenic: A micro-organism causing an insect disease. Entomopathogenic nematodes: Small parasitic roundworms that infect and often quickly kill insects that live underground. Epigeal emergence (of germinating pulses): The seed leaves (cotyledons) appear above the ground when the seedling has emerged; for example, lupins. (The epicotyl is the portion of the seedling stem above the cotyledons.) See also Hypogeal emergence.
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GLOSSARY
Eye (of sugarcane): The sugarcane bud. New shoots sprout from the eyes when the sett is planted.
seedling stem below the cotyledons.) See also Epigeal emergence.
Fallow (of sugarcane): Ground that does not contain a sugarcane crop within a cane farm for at least 6 months.
IGRs: Insect growth regulators. Synthetic insecticides that disrupt the development of insects. Usually slower acting than nerve or stomach poisons.
Fill-in (of sugarcane): Bringing more soil in the open drill when plants are about 60 cm tall.
Improved pastures: Sown pastures. Native grasses are supplemented or replaced by exotic grasses and forbs (usually pasture legumes). These pastures usually require fertiliser.
Ferrosols: Also known as krasnoszems and chocolate soils. Soils of volcanic origin with high free iron and clay contents. They occur along the eastern coastline, in northern parts of Western Australia and the Top End (NT). In higher rainfall zones, they may be very deep and well drained. Wellknown agricultural regions with Ferrosol soils include the Atherton Tablelands and the South Burnett in Queensland. Flare, flaring (after insecticide application): Rapid increases in pest abundance, usually of aphids, mites or whitefly, following application of broadspectrum insecticides targeted against other pests. The mechanism is thought to be largely due to the loss of natural enemies of these pests, allowing them to increase unchecked, but may also include insecticidal stimulation of the pests’ growth rate or fecundity (egg production). Fledging (of locusts): Final moult from nymph to winged adult. Generation time: Average time between birth or egglaying and death of a population of insects. Genitalia: External parts of the reproductive system of arthropods. Germinating seed baits: See Monitoring methods. Glabrous: Free from hairs. Heliothine moths: A sub-family of the Family Noctuidae. Includes the genera Adisura, Australothis, Heliocheilus, Heliothis and Helicoverpa. Heliothis: Sometimes used as a common name for native budworm and/or corn earworm. Heliothis was the generic name for these two species before they were re-assigned to the genus Helicoverpa. Honeydew: Sugar rich waste excreted by feeding aphids and whitefly. Hypogeal emergence (of germinating pulses): The seed leaves (cotyledons) remain underground when the seedling has emerged; for example, field peas, faba beans. (Hypocotyl is part of the
Indeterminate: Crop cultivars or varieties with asynchronous flowering and podding; that is, where these reproductive stages overlap. In many indeterminate cultivars, rain events can trigger further flowering flushes. See also Determinate. Insecticide resistance management strategy (IRMS) (for cotton): An area-wide voluntary strategy developed by the cotton industry to manage resistance to insecticides and miticides. The strategy, which is updated annually and appears on the NSW DPI website, includes: (i) restricting the number of insecticide applications for any one group of insecticides which have the same mode of action; (ii) restricting the time period over which specific insecticides can be used, thereby reducing the duration of selection; (iii) preferentially placing more disruptive insecticides later in the season; (iv) recommending destruction of diapausing H. armigera populations under cotton by cultivation. The strategy currently includes H. armigera, spider mites, cotton aphids and silverleaf whitefly. Insects and allied forms: Members of the Class Insecta. In this book, ‘allied forms’ include other arthropods such as crustaceans, collembola, mites, spiders and millipedes, together with some molluscs, including snails and slugs. Instar: The period between insect moults; between egg hatch and first moult, and between any two subsequent moults. IPM: Integrated pest management. Originally conceived as combining non-insecticidal methods such as resistant plant varieties, cultural and biological control. Now extended to include intervention with insecticides generally compatible with the above methods. Larva (plural larvae): The immature stage between the egg and pupa. The larvae of some groups may be called caterpillars, grubs or maggots. ‘Larva’ is
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GLOSSARY
also applied to the stage of some mites and spiders that emerge from eggs, often with a reduced number of legs, with the final legs appearing when they moult into nymphs. The term is also used for immature thrips (Order Thysanoptera). Lodging (of crop plants): Falling or collapse of plants to ground level due to heaviness of plants, weakness of roots or softness of soil. One cause of lodging may be insect activity weakening the roots or stems of plants. Major and minor male dung beetles: Adult dung beetles can be separated into two distinct classes: large, ‘major’ males, and ‘minor’ males. Larval growth is usually dependent on how much dung is available to them during the larval phase. Larvae growing larger than a genetically determined threshold size metamorphose into ‘major’ male adults with long horns, and larvae not reaching this size metamorphose into ‘minor’ males lacking or with very short horns. Minimum tillage (conservation tillage, zero tillage, stubble retention): Residue of the previous season’s crop or pasture is left largely undisturbed in the soil. Seeds of the new crop are planted in a narrow furrow; the space between furrows is sprayed with herbicide for weed control. Monitoring methods Beat sheet-sampling: A standard beat sheet measuring 1.5 m × 2 m (or 1.5 m × 1.5 m in short crops such as dryland mung beans) is placed in the furrow between rows and extended up and over the adjacent row of crop. A metre stick may be used to beat the plants 10 times against the beat sheet, moving from the base to the tops of the plants. The insects dislodged from the plant onto the sheet are quickly counted. Results can be expressed as number per plant or metre of row. Counts can be converted to insects per square metre by dividing counts by the row spacing, in metres. Beat sheets are not suitable for use after irrigation or heavy dew, or during windy weather. For lucerne insects and mites, a white tray can be used in which to beat foliage. Germinating seed baits: For monitoring soil insects prior to planting. Immediately following first rains and prior to planting, a quantity of the insecticide-free crop seed is soaked in water for at least 2 hours to initiate germination and a small
Sampling with a beat sheet; the plants in front of the operator are beaten to dislodge insects onto the sheet. (DPI&F Qld: J. Wessels)
handful of the germinating seed is buried under 1 cm of soil at each corner of a 5 m × 5 m square at five widely spaced sites per 100 ha. One day after seedling emergence, dig up the plants and count the insects. Pre-soaked sorghum seed emerges after 3–4 days in summer and 5–6 days in autumn–spring. See Soil insect rating (SIR) for interpretation of results. Pheromone traps: For monitoring activity of (usually male) adult insects. Used for detecting stored-product moths. Sticky cardboard pheromone traps for some field crop pests are commercially available but numbers caught in traps are sometimes difficult to interpret and may not be good indicators of damage risk or the need for control. Pitfall traps: For trapping insects moving over the ground. A typical trap consists of a container sunken into the ground so that its rim is level with the ground surface. A bait (e.g. fresh dung) may also be used to attract insects. Shelter bags: For monitoring field crickets. Hessian bags placed on cricket-prone pastures indicate relative numbers of crickets in pastures. Spade-sampling: For soil insects. A spade is used to dig a defined area of soil. Typically, a block of soil 30 × 30 cm is extracted down to the moist layer and searched for soil insects. Many insects will usually be located near the moist–dry interface. Counts may be converted to numbers per square metre or per hectare. Spear-sampling for stored grain insects: Trucksampling spears are generally stainless steel with
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GLOSSARY
a double tube construction. The inner tube opens progressively from the bottom first, to ensure a representative sample from the whole load. Some tubes have openings only at the bottom end and are designed to take a sample from the bottom of the truck-load, others have openings over the entire length so that a cross-section sample is taken. The spears are 1.5–2 m long and 3–3.5 cm inner diameter. Suction-sampling: Used in cotton for sampling mobile arthropods, including predators. A suction machine (D-vac Homelite Textron, Inc., NC, USA or Stihl) with a 120 mm diameter suction tube is used to suck insects from plants and collect in a gauze bag The tube of the machine should be drawn from the top to the base of the plants. A 20 m of row of cotton is sampled, which should be divided into four random samples of 5 m to cover the field. After each sampling, the contents of the D-vac should be transferred to separate plastic bags and taken to the laboratory and counted. Suction sampling is also used to sample insects on prostrate peanuts. The sampling unit is as for cotton; that is, 20 m of row, but the sampling tube is held vertically and passed through the top of the canopy (to avoid sucking up dirt). Sweep net-sampling: A sweep net made from fine mesh or a calico bag (approximately 60 cm deep) is attached to a hoop of 45 cm in diameter with a handle 1.5 m long. The net is used to sweep insects from foliage. The sample is taken by walking briskly and sweeping the net in front so that the bottom of the sweep strikes the canopy about 20–30 cm from the top. For pulse crops, the sampler typically makes 20 sweeps in 20 m over two rows, after which the contents are counted or transferred to a plastic bag. At least five to six sites are sampled per field. Counts are expressed as numbers per sweep or, in some cases, sweeps can be calibrated to numbers per square metre or per metre of row. For summer pulses, factors to convert insect counts to beat cloth equivalents per square metre are available from agronomists. The technique can be modified for sampling insects with edge distributions, such as pea weevil. Visual-sampling: The whole plant, including buds, flowers, fruit, leaves and terminals, is checked and all predators and pests on the plant
Sweep nets are used to sample insects from a wide range of field crops. (DPI&F Qld: J. Wessels)
are counted. Counts can be expressed as number per plant, number per metre of row or number per square metre. Sample size can also be calculated from the standard error of the mean, time taken to collect the samples and labour costs involved in collecting the samples. This method is mainly used in cotton. A hand lens is often useful. Neonate: A larva recently hatched from an egg. Non-persistent transmission of viruses by aphids: Plant viruses may contaminate an aphid’s mouth parts after it probes or feeds on an infected plant. The virus may be transmitted to other plants when the aphid subsequently probes or feeds on ‘clean’ plants. Non-persistent viruses remain viable on mouth parts for only a short time (up to 4 hours), may be lost after a few probes and do not survive a moult of the aphid. These viruses do not replicate in the aphid’s tissues. See also Persistent transmission. Nymph: Immature stage that is a small copy of the adult form. NPV: Nuclear polyhedro virus. A virus disease naturally infecting some Lepidoptera larvae. In Australia, Helicoverpa spp. are often diseased by NPV. Commercial formulations of the virus are used to control H. armigera. Okra leaf (of cotton): Cotton varieties with a deeply lobed leaf shape (Fig. 3.12). Oligochaetes: A group of annelid worms that includes earthworms. Organophosphate insecticides: Inhibit acetylcholine esterase at nerve synapses. Oviposit: To lay eggs.
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GLOSSARY
Ovipositor: Projection from the abdomen of female insects through which eggs are laid.
Pleopod: An abdominal swimming appendage present in some crustaceans.
Paddock: Field.
Plough-out (of sugarcane): Complete removal of the crop, including the root mass, from the soil.
Paddy rice: Unprocessed (unhulled) rice grain. Panicle: The branched inflorescence that bears the grain in cereal crops. Parasitoid: An insect whose immature stages feed on a host insect over a period of time but which eventually kills its host. In contrast, a parasite does not usually kill its host while a predator kills its prey quickly. Parthenogenesis: Reproduction without mating. Adults are all females and offspring are genetically identical to the parent. This is common in the life cycles of aphids and in some weevil species. Pectinate antennae: Comb-shaped. Perenniation (of weed): Living for more than 1 year. Persistent transmission of plant viruses by aphids: Aphids acquire viruses from feeding on infected plants and retain the virus for over 100 hours. After a latent period during which some types of virus propagate within the body of the aphid and others simply circulate within the body, the aphid is capable of transmitting the virus through its salivary glands for the remainder of its life. Semipersistent viruses are capable of being retained for 10–100 hours in the aphid’s foregut. See also Non-persistent transmission.
Pitfall trap: See Monitoring methods. Prehensile claws: Grasping claws. Pre-oviposition period: The time between emergence as an adult and the time of first egg-laying. Pre-pupa: Near the end of larval stage, the larva builds a cell or spins a cocoon as protection for its pupal stage. The external appearance of the larva may change during the pre-pupal stage. Press wheel: Rubber wheel attached to a crop seeder to compress the soil at furrow closure. Soil compaction provides better seed–soil contact, aids germination and restricts insect movement through the soil. Primary pest (of stored grain): Pests that can damage sound grain (see also Secondary pests).
Importance: Major: Likely to cause significant economic damage to the host crops unless controlled. Minor: Damage may result in some crop loss, for which it may or may not be economical to control.
Prolegs on some caterpillars: Fleshy non-segmented legs on caterpillar’s abdomen. They terminate in a set of microscopic hooks or crochets and are used for walking and clinging. Prolegs on abdominal segments 3–6 are called ventral prolegs (Fig. 2.33). The number of ventral prolegs for caterpillars listed in this book ranges from two to eight. The last pair of prolegs at the end of the abdomen are called anal prolegs or claspers (Fig. 2.33). All caterpillars (lepidopteran larvae) referred to in this book have anal prolegs so only the ventral prolegs are referred to when differentiating between species. Prolegs are additional to the three pairs of thoracic (or true) legs.
Range of pest: Widespread: Likely to be a pest over most of the area where the host crop is grown. Restricted: Damage is confined to a part/ parts of the area where the crop is grown.
Pronotum of an insect: The upper, often shield-like, hardened plate, located just behind the head (Fig. 2.59). The dorsal part of the first thoracic segment (prothorax).
Seasonal occurrence: Regular: Likely to be a pest in most years. Irregular: Unlikely to be a pest every year.
Prothorax: First segment of the thorax.
Pest status (as used in this book):
Phytophagous: Plant-eating organism. Plant names: Common and scientific names used in this book are generally consistent with those in Lazarides, M. and Hince, B. (1993). CSIRO Handbook of Economic Plants of Australia.
Pupa (plural pupae): Inactive, non-feeding stage between larval and adult stages in insect orders that undergo complete metamorphosis. Also used to describe the pre-adult stages of whiteflies and thrips. Pyrethroid insecticides: Act on the sodium channels of nerves.
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GLOSSARY
1
Genus (initial capital letter)
2 species
(all lower case)
Genus and species names are printed in italics. 3
Author who originally described the species (the name appears in parentheses if there has been a change from the original genus)
Side view of a scarab larva showing position of the raster on the last abdominal segment and (right) end view of the raster surrounding the anus. (NSWDPI: Alan Westcott)
Quadrat: An area used for sampling animals or plants. Typically, quadrats of known area may be defined by square, rectangular or circular frames. Raster: Hair pattern. In particular, the raster on the ventral surface of the last abdominal segment of scarab larvae is important in separating species. Ratoon (of sugarcane): A following crop that grows out of the root mass and subterranean parts of the stem of the previous crop that was left in soil after that crop was harvested. Ratooning (of sugarcane): Allowing the crop to regrow after harvest from the original root mass, for another season. Receptacle (of thistles): The tissue within the head to which seeds are attached. Reduced rates of pesticides: Use of a lower product rate than specified on the registered label or, where more than one rate is specified for a particular use, lower than the maximum label rate. Refuge (plural refugia): A pesticide-free area for (usually) beneficial arthropods to survive and reproduce. Rosette (of plant): Flat, non-flowering part of plants such as thistles and Paterson’s curse. Salt (sodium chloride as an additive to insecticides): Addition to some insecticides increases their efficacy so the insecticide rate can be reduced without loss of efficacy. Scientific name: The following is an example of how scientific names are written. Helicoverpa1 punctigera2 (Wallengren) 3 Lepidoptera4: Noctuidae5
4
Order
5
Family
Plant names in this book do not include the name of the author. Scarabaeiform: In the shape of a scarab larva. Sclerotised: Hard, darkened structure, area(s) or plate(s) on the body of an insect. Scutellum: The posterior part of the thorax, prominent and triangular in some groups of true bugs. Secondary pest (of stored grain): Insects which live on broken grain or that damaged by primary pests. Seed dressing: Seed is treated with an insecticide prior to planting. A contact insecticide may prevent the seed being eaten or carried away (e.g. by ants) or a systemic insecticide may protect the germinating seedling from insects and mites. Insecticide may be applied as a liquid, slurry or dust, with or without a sticker. sens. lat. following some scientific names: Latin sensu lato. Terminology sometimes used to indicate ‘as broadly defined’. Sett (of sugarcane): A 20–30 cm long piece of cane stalk with active buds (eyes) used for planting. Sibling species: Species that are morphologically very similar but genetically different. Siphunculi (singular siphunculus) of aphids: Pair of tubes at the rear sides of the abdomen through which defence chemicals and alarm pheromones are dispensed. See also Cauda. Sod-sowing: A rice-sowing method that involves drill-sowing seed into a pasture paddock prior to flooding. Soil insect rating (SIR) (for summer field crops): Assessment of need to control soil insects based on relative destructive equivalence of each species
5 11
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GLOSSARY
detected around 20 GSB. (See also Monitoring methods, Germinating seed baits.)
developed by CSIRO for the control of redlegged earth mite.
SIR = (1 × SW + 3.5 × LW + 5 × SR + 10 × LR + 5 × CR)/20
Tipping (of sugarcane): Cane plants falling over, leaving the root system exposed out of soil.
where
Tipping out (of cotton seedlings): Death of the apical meristem, often caused by insects.
SW = No. of small false wireworm beetles
SR = No. of cockroaches less than 20 mm length
Thorax: The middle of the three major divisions of the insect body (head, thorax, abdomen) to which the legs and wings are attached.
LR = No. of cockroaches more than or equal to 20 mm length
Topping (tobacco): Flower removal to maximise vegetative growth.
CR = No. of crickets around the 20 GSB.
Topping (weeds): The use of low rates of herbicide to reduce viable seed set of weeds.
LW = No. of large false wireworm beetles
For example, if there are a total of 10 of each type of insect around the 20 GSB then the SIR = 12.25. Control surface active insects if the SIR is more than 6. sp., spp.: species (singular, plural). Follows the generic name when the species is unknown or unspecified. Spade sample: See Monitoring methods. Spear sample: See Monitoring methods. Spider mites: The mite Family Tetranychidae, which includes a number of pest species, including twospotted mites. Spur: A large spine, for example, sometimes found on insect legs (Fig. 17.12).
Tubercle: A small knob-like projection on an insect. Univoltine: One generation per year. Ventral surface: Lower surface. Vertex: The area on the head of an insect between the eyes. Visual-sampling: See Monitoring methods. Volunteer plants: Plants from the previous season that grow unwanted in the following crop. Windrowing: The crop is cut to stop growth and even up maturity. The cut vegetation is placed in narrow rows (windrows) to be picked up with a special harvester front. Windrowed vegetation acts as a refuge for invertebrates that may contaminate grain.
Squaring (of cotton): Time period during which flower buds (squares) are produced. Stool (of sugarcane): The basal part of the plant with stem(s) and roots attached. Suction-sampling: See Monitoring methods. Sweep net: See Monitoring methods. Systemic insecticide: An insecticide absorbed into plant tissue. Effective against sucking insects. TIMERITE®: Management strategy, based on the critical timing of acaricide application in spring,
Windrowed barley. (SARDI)
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INDEX
Page numbers of scientific names are duplicated for common names. Where multiple page numbers are given, major entries are shown in bold. Abacarus hystrix 28, 394 Acacia nilotica 387 Acanthoscelides macrophthalmus 366 Acanthoscelides obtectus 214 Acarophenax tribooli 59 Acaropsis docta 59 Acarus siro 42 Aceria chondrillae 460 Aceria tosichella 28 Achyra affinitalis 227 Aconophora compressa 379 Acroelytrum muricatum 190 Acrossidius pseudotasmaniae 399 Acrossidius tasmaniae 17, 399, 401, 405, 407, 409, 411 Aculus hyperici 466 Acyrthosiphon kondoi 72, 265, 420, 428 Acyrthosiphon pisum 72, 266, 420, 428 Adelina sp. 324 Adelina triboli 59 Adelium brevicorne 23, 141, 263 Adoryphorus couloni 401 Adreppus sp. 491 African black beetle 128, 321, 347, 397, 402 Agathis sp. 230 Agrotis infusa 25, 83, 166, 364 Agrotis ipsilon 23, 242, 406 Agrotis munda 24, 28, 38, 166, 242, 347 Agrypnus sp. 21 Agrypnus variabilis 80, 124, 125, 210, 310 Ahasverus advena 52 Alternanthera spp. 76 amaranths 226 Amaranthus sp. 226 Ambrosia artemisiifolia 372 Amnemus quadrituberculatis 218, 357 amnemus weevil 218, 357 Amphibolus flavipes 59 Amphimermis spp. 501 Amrasca terraereginae 74 Amyna axis 247 Angoumois grain moth 55 Anisopteromalus calandrae 59 annual ragweed 372 annual ryegrass 28, 394, 398, 417, 419 Anomis flava 88 Anoplognathus suturalis 404 Anthrenus flavipes 49 Anticarsia irrorata 239 Antitrogus consanguineus 332
030701•Pests of Field Crops and 513 513
Antitrogus parvulus 326 Antitrogus planiceps 331 Antitrogus rugulosus 330 Anystis wallacei 395, 417 Apanteles glomeratus 154 Apanteles sp. 33, 34, 362 Apanteles tasmanica 435 Aphaenogaster pythia 339, 364 aphid transmission of viruses 73, 151, 267, 345 Aphidius colemani 74 Aphidius ervi 428 Aphis chloris 467 Aphis craccivora 71, 178, 264, 420 Aphis glycines 179 Aphis gossypii 71 Apina callisto 424 Apion congestum 217 apple dimpling bug 86 Aproaerema simplexella 221 aquatic earthworm 281 Arachnodima bribbarensis 21 Arachnodima opaca 21 Arachnodima ourapilla 21 Arachnodima xenicon 21 Araneus spp. 103 Arcototheca calendula 12, 72, 138, 144, 145, 148, 150, 151, 346, 394, 416, 417, 424, 434 Argentine stem weevil 396 arid zone grasshoppers 485 Aristotelia sp. 377 armyworms 30, 38, 83, 130, 240, 290, 295, 300, 337, 363 asilid predation 324, 405, 503 Aspergillus and peanuts 174, 209, 229 assassin bugs 78, 104, 246, 320 Asynonychus cervinus 144 Athetis tenuis 242 Atrichonotus taeniatulus 144, 430 Atriplex spp. 424 Austracris guttulosa 106, 127, 316, 345, 496 Australian crop mirid 182 Australian plague locust 29, 96, 127, 290, 316, 485, 486 Austroagilla torrida 75, 429 Austroasca alfalfae 181, 429 Austroasca viridigrisea 74, 181, 429 Austroicetes cruciata 30, 290, 493 Austroicetes nullaborensis 495 Austroicetes vulgaris 495 Automolius depressus 404 Axonopus compressus 360 Axonopus spp. 359, 360, 398 Baccharis halimifolia 374
Bacillus thuringiensis (Bt) 59, 65, 96, 114, 240, 243, 253, 273 Balaustium medicagoense 138, 413 balaustium mite 138, 413 Balclutha 345 barley yellow dwarf virus (BYDV) 15 Baryopadus corrugatus 358 Bathurst burr 373 Bathypogon spp. 503 Bathytricha truncata 291, 336 Bdellodes lapidaria 395, 418 bean bruchid 213 bean flower caterpillar 234 bean leafroller – see legume webspinner bean looper 248 bean podborer 224 bean yellow mosaic virus 267 beanfly 219 Beauvaria spp. 21, 188, 202 beet webworm 226 beet western yellow lutreovirus (BWYV) 151 Bemisia tabaci 66, 158, 176 Bidens pilosa 254 big eyed bug 104, 245 bitter peas 433 black cutworm 23, 242, 406 black field cricket 66, 159, 175, 309, 419 black field earwig 122, 176, 298 black leaf beetle 333 black soil scarab 18 black sunflower scarab 156, 208 blackberry nightshade 349 blackheaded pasture cockchafers 17, 399, 401, 405, 407, 409, 411 black-keeled slug 11, 137 bladder ketmia 70, 72, 101 blady grass 313, 323, 329, 339 Blaesoxipha spp. 489, 498 Blattisocius tarsalis 59 blistered pyrgomorph 492 bloodworms 286, 292 blue oat mites 11, 139, 164, 262, 394, 415, 426 bluegreen aphid 72, 265, 420, 428 bogong moth – see common cutworm Bollgard II® 65, 77, 81, 86, 89, 96, 98, 101 Bourletiella hortensis 165 Bourletiella viridescens 165 Brachiaria spp. 249, 323, 356, 373 Brachycaudus helichrysi 151 Brachycaudus rumexicolens 442 Brachyexarna lobipennis 501 Bracon hebitor 59 Bradysia spp. 219
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INDEX
Brassica spp. 74, 84, 98, 200 Brevicoryne brassicae 149 broad-leaf dock 442 broad-leafed carpet 360 brokenbacked bug 76, 183 bronze dung beetle 478 bronzed field beetle 23, 141, 263 brown clover mite – see bryobia pasture mite brown cutworm 24, 28, 38, 166, 242 brown dungball roller 480 brown field slug 164 brown flea beetle 81 brown lacewing 107, 428 brown mirid 76, 160, 184 brown pasture looper 147, 266 brown planthopper 292 brown shield bug 196 brown smudge bug 105, 245 brown sugarcane cicada 312 brown wheat mite 28 Bruchidius mackenziei 214 Bruchidius sahlbergi 387 Bruchophagus roddi 435 Bruchus pisorum 268 bryobia pasture mite 139, 143, 414 Bryobia praetiosa 139, 143, 414 Bt – see Bacillus thuringiensis Bucculatrix gossypii 88 Bucculatrix ivaefolia 377 Bucculatrix parthenicola 387 buffel grass seed caterpillar 361 bullhorned dung beetle 476 Bundaberg canegrub 324 BYDV – see barley yellow dwarf virus cabbage aphid 149 cabbage moth – see diamondback moth cabbage white butterfly 153 cabbage-centre grub 152 cactoblastis 389 Cactoblastis cactorum 389 cactus moth 390 Cadra cautella 57 Callosobruchus maculatus 214 Calolampra elegans 156 Calolampra solida 156 Calosoma schayeri 33, 107 Calycomyza lantanae 381 Campsomeris spp. 324 Campylomma liebknechti 86 canegrubs 322 cape gooseberry 349 cape gooseberry budworm 350 capeweed 12, 72, 138, 144, 145, 148, 150, 151, 346, 394, 416, 417, 424, 434 Capitophorus elaeagni 151 carabid predation 11, 21, 23, 25, 33, 107, 126, 138, 164, 405, 488 Carcelia illota 274 Carduus nutans 448 carpet grasses 359, 360, 398 Carpophilus spp. 97
Carpophilus hemipterus 51 caseworm 294 castor oil plant 200 catabena moth 381 Catochrysops panormus 235 caudata canegrub 326 Celama taeniata 291 Celosia sp. 226 Cenchrus spp. 361 Cephalonomia gallicola 59 Cephalonomia meridionalis 59 Cephalonomia waterstoni 59 Cercospora sp. 201 cereal curculio – see spinetailed weevil cereal rust mite 28, 394 Cermatulus nasalis 105, 245 Cernuella virgata 7, 136, 261 Chaetocnema sp. 81 Chamaesphecia doryliformis 443 Chamaesphecia mysiniformis 447 Chauliognathus lugubris 211 Chauliognathus pulchellus 108 Chauliognathus spp. 211 Cheiroplatys latipes 404 Chenopodium sp. 226 Cherax destructor 286 chevron cutworm 148 Cheyletus malaccensis 59 chickpea chlorotic dwarf virus 267 Childers canegrub 326 Chilean predatory mite 427 Chilo polychrysa 294 Chilo suppressalis 294 Chironomus spp. 286, 292 Chironomus tepperi 286 Chloris spp. 360 Choetospila elegans 59 choko 194 chondrilla gall midge 460 chondrilla gall mite 460 Chondrilla juncea 460 Chortoglyphus gracillipes 59 Chortoicetes terminifera 29, 96, 127, 290, 316, 485, 486 Chrysodeixis argentifera 149, 254, 350 Chrysodeixis eriosoma 254 Chrysodeixis spp. 266 Chrysolina quadrigemina 467 Chrysopa spp. 74, 106, 154 Ciampa arietaria 147, 266 Cicadetta crucifera 312 Cicadetta multifascia 312 Cicadulina bimaculata 123 cigarette beetle – see tobacco beetle Cirsium vulgare 465 clover casebearer 423 clover seed moth – see clover casebearer cluster caterpillar 83, 240, 349 Cnaphalocrocis medinalis 294 cobbler’s pegs 254 Coccinella transversalis 108 Coccus longulus 365
Cochlicella acuta 9, 136 Cochylis atricapitana 458 cockscomb 226 Coelophora inaequalis 108 Coleophora alcyonipennella 423 Colpochila obesa 404 common armyworm 30, 38, 130, 153, 240, 290, 295, 300, 337, 361 common brown leafhopper 75, 345, 429 common crowfoot 346 common cutworm 25, 83, 166, 364 common garden snail 356 common garden springtail 165 common grass blue butterfly 234 common hoverfly 150 common northern dung beetle 481 common pyrgomorph 492 common spotted ladybird 108 common white snail 7, 136, 261 confused flour beetle 54 Conogethes punctiferalis 101, 133, 303 consobrina canegrub 327 Contarinia sorghicola 301 Coranus triabeatus 104 corbie 406 Cordyceps aphodii 401, 405 Cordyceps heteropoda 313 Cordyceps oncoperae 408 Cordyceps spp. 210 corn aphid 14, 16, 17, 28, 72, 127, 299, 317 corn earworm – see cotton bollworm Corrhenes stigmatica 212, 432 Corticaria spp. 51 Coryphistes interioris 491 Cotesia rubecula 154 Cotesia sp. 337 cotton aphid 71 cotton bollworm 37, 65, 89, 131, 152, 161, 241, 243, 274, 300, 350 cotton bunchy top disease (CBT) 73 cotton harlequin bug 97 cotton leaf perforator 88 cotton leafhopper 74 cotton looper 88 cotton seedling thrips – see onion thrips cotton tip borer 81 cotton tipworm – see cotton tip borer cotton webspinner 227 CottonLOGIC 94 cottonseed bug 87 cowpea aphid 71, 178, 264, 420 cowpea bruchid 214 crabgrass leaf beetle 356 Creontiades dilutus 75, 185 Creontiades pacificus 76, 160, 184 Crocidosema plebejana 81 crotalaria moth 237 Crotalaria spp. 70, 178, 186, 194, 195 200, 204, 228, 229, 230, 235, 237, 334 crusader bug 191
514
030701•Pests of Field Crops and 514 514
29/6/07 17:40:02
INDEX
Cryptoblabes adoceta 231, 302 Cryptolaemus montrouzieri 108, 181 Cryptolestes spp. 50 Cryptomorpha flaviscutellaris 488, 495 Cryptostegia grandiflora 391 cucumber mosaic virus 267 Cucumis spp. 72 cucurbit shield bug 192 curled dock 442 cutworms 23, 28, 39, 83, 126, 145, 148, 158, 166, 242, 299, 347, 364, 406 Cylindracheta psammophila 13, 395 Cystiphora schmidti 460 Dactylopius ceylonicus 388 Dactylopius confusus 388 Dactylopius opuntiae 388 Dactylopius tomentosus 388 damsel bug 86, 105, 185, 245 damselfly 293 dark cerulean – see bean flower caterpillar darkheaded riceborer 294 Datura sp. 74 Daviesia spp. 433 dayfeeding armyworm 130, 295, 300, 337, 363 Deraeocoris signatus 105, 245 Dermestes ater 503 Dermolepida albohirtum 329 Deroceras parnormitanum 164 Deroceras reticulatum 11, 137, 164 Desiantha caudata 20, 57, 144 Desiantha diversipes 21, 144 devil’s apple 349 Dialectica scalariella 456 diamondback moth 146 Diarsia intermixta 148 Dicranolauis bellulus 110 Dictyotus caenosus 196 Dinarmus basalis 59 Diomus notescens 108 Diphucephala colaspidoides 404 dock aphid 442 dock moth 443 docks 424, 442 dragonfly 293 driedfruit beetle 51 drugstore beetle 44 dusky pasture scarab 402, 403 Dysdercus sidae 96 dytiscid water beetle 293 Earias huegeliana 97, 101 Earias vittella 101 earwigs as predators 33, 58, 124, 126, 164, 211, 246 eastern false wireworm 23, 80, 141 eastern plague grasshopper 489 Ebor grassgrub 359 Echium italicum 451 Echium plantagineum 451 Echium vulgare 451
Ectropis despicata 232 eight spots moth 247 Elasmolomus pallens 189 emerald dung beetle 479 emu foot 197, 222 Encarsia formosa 71, 159 Endotricha puncticostalis 227 Entomophaga grylli 502 Entomophthora sp. 240 Ephestia kuehniella 58, 352 Ephysteris promptella 315 Epiblema strenuana 373, 382, 386 Epinota lantana 381 Epiphyas postvittana 82 Eremophila spp. 492 Eretmocerus hayati 178 Eretmocerus sp. 71 Erharta sp. 408 Ericeia inangulata 243 Eriocerus martini 378 Eriocerus tortuosus 378 Erodium spp. 166, 346, 424 Etiella behrii 228, 270, 432 etiella moth 228, 270, 432 Eublemma amoena 464 Eublemma dimidialis 243 Eubrostis spp. 236 Euclasta whalleyi 391 Eukerria saltensis 281 Eumargarodes laingi 317 Euoniticellus africanus 477 Euoniticellus fulvus 474 Euoniticellus intermedius 477 Euoniticellus pallipes 475 Euphorbia pulcherrima 70 European earwig 56, 124, 140, 165 Euryaulax carnifex 320 Evippe sp. 382 Exapion ulicis 445 Eysarcoris distinctus 205 Eysarcoris trimaculatus 205, 290, 293 Fabrictilis gonagra 192 false wireworms 23, 80, 141, 143, 157, 211, 263, 298 Ferrisia virgata 86 fescues 360, 406 Festuca spp. 360, 406 fiddle dock 442 Fiji leaf gall virus 320 Fipurga crassa 501 fivehorned dung beetle 481 flat grain beetles 50 flatheaded pasture webworm 359 flour mite 42 flower beetles 97 flower wasps 405 food supplements 113 foreign grain beetle 52 Forficula auricularia 56, 124, 140, 165 forget-me-not – see pale pea blue butterfly Frankliniella occidentalis 78, 86, 207, 268
Frankliniella schultzi 78, 86, 152, 207, 268, 429 Frankliniella williamsi 125 Fraus polyspila 408 Fraus simulans 408 French’s canegrub 327 Froggatt’s canegrub 328 Fuller’s rose weevil 144 funnel ant 339, 364 furniture carpet beetle 49 garden springtail 165 Gastrimargus musicus 316, 502 gazella dung beetle 479 Geocoris lubra 104, 245 Geotrupes spiniger 475 geranium plume moth 223 ghost moth 408 giant pigweed 226 giant sensitive plant 373 giant termite 309 glossy shield bug 105, 245 Glyptophysa sp. 283 Gonocephalum macleayi 23, 81, 157, 298 Gonocephalum spp. 211, 263, 347 goosefoot 226 gorse 444 gorse seed weevil 445 gorse spider mite 444 gorse thrips 445 granary weevil 47 granulose dung beetle 483 Graptostethus servus 188 grass anthelid 29 grass caterpillar – see sod webworm grata canegrub 328 greater sandy dung beetle 477 green carab beetle 33, 107 green cicada 312 green cutworms 148 green lacewings 74, 106, 154 green mirid 75, 185 green muscardine fungus 30, 315, 316, 324, 403, 488, 491, 493, 495, 498, 500 green peach aphid 72, 74, 149, 151, 345 green soldier beetle 108 green stink bug 99, 197 green vegetable bug 98, 127, 198, 290, 347 green vegetable bug parasite 202 greenhouse whitefly 67, 158 greenhouse whitefly parasite 71, 159 grey cluster bug 151, 160 grey dungball roller 480 grey false wireworm 23, 141, 263 greyback canegrub 329 grisea canegrub 329 ground cherries 72 groundselbush 374 groundselbush gall fly 375 groundselbush leaf-tying moth 377 groundselbush stemboring moth 376 groundsel leafeating beetle 375
515
030701•Pests of Field Crops and 515 515
29/6/07 17:40:03
INDEX
groundsel stem borer 374 Gryllotalpa spp. 310, 395 Guinea grass 329, 323, 336 Gymnoscelis lophopus 233 hairy flower wasps 405 hairy fungus beetle 53 hairy rice caterpillar 291 hairy scarab 402, 404 Halotydeus destructor 12, 28, 139, 141, 164, 261, 415, 426, 512 Hanseniella spp. 307 Haplothrips victoriensis 427 Harmonia conformis 108 Harmonia octomaculata 108 harrisia cactus 378 harrisia cactus mealybug 378 Hednota spp. 26 Helea tuberculatus 23, 141 Helicoverpa armigera 37, 65, 89, 131, 241, 243, 274, 300, 350 Helicoverpa assulta 350 Helicoverpa punctigera 37, 89, 166, 243, 271, 350, 435 heliothine moths 89 Heliothis punctifera 89, 275 Helix aspersa 356 Hellula hydralis 152 Hemianax papuensis 293 Heteronychus arator 128, 321, 347, 397, 402 Heteronyx obesus 18 Heteronyx piceus 209 Heteronyx rugosipennis 209 Heteronyx spp. 128, 208 Heteronyx tasmanicus 404 Heteropelma scaposum 274 Heteropsylla cubana 365 Heteropsylla spinulosa 373 Hibiscus heterophyllus 70 hibiscus mealybug 86, 180 Hibiscus trionum 70, 72, 101 Hippodamia variegata 108 Holepyris sylvanids 59 horehound 446 horehound clear-wing moth 447 horehound plume moth 447 hoverfly 17, 151, 180, 234 humpbacked dung beetle 476 Hydrellia mareeba 292 Hydrellia michelae 288 Hypena laceratalis 381 hypena moth 381 Hypericum perforatum 466 Hypogeococcus festerianus 378 Ichneumon prommisorius 274 Illyrian thistle 461 Imperata cylindrica 313, 323, 329, 339 Indian meal moth 56 inland armyworm 32, 36 inland field cricket 419 Inopus flavus 313
Inopus rubriceps 313, 359 Invermay bug 160 Iphiaulax sp. 213 Ipomoea spp. 70, 74 Isidorella newcombi 283 Isopteron punctatissimus 23, 141 Isopteron sp. 263 Israelius carthami 59 Italian bugloss 451 Ithome lassula 366 Jamides phaseli 234 Josephine burr 70 joyweed 76 jumping spiders 103 khapra beetle 49 kikuyu grass 356, 359, 360, 361, 363, 398 lablab bean 191 Lablab purpureus 191 Laccotrephes tristis 293 lacewings 16, 71, 74, 105, 106, 127, 151, 154, 175, 181, 207, 246, 428 Lampides boeticus 236 lantana 379 Lantana camara 379 lantana flower caterpillar 381 lantana lace bug 380 lantana leaf-folding caterpillar 382 lantana leafmining beetles 380 lantana leafmining fly 381 lantana mealybug 379 lantana plume moth 382 lantana seed fly 381 lantana sucking bug 379 Lantanophaga pusillidactyla 382 large brown bean bug 193 Larinus latus 462 Lariophagus distinguendus 59 Lasioderma serricorne 351 Latheticus oryzae 55 lawn armyworm 295, 363 leafcurl plum aphid 151 leafhopper transmitted diseases 124, 267, 345, 429 Lecithocera sp. 333 legume webspinner 230 Lepidiota caudata 326 Lepidiota consobrina 327 Lepidiota crinita 324 Lepidiota frenchi 327 Lepidiota froggatti 328 Lepidiota grata 328 Lepidiota grisea 329 Lepidiota negatoria 330 Lepidiota noxia 330 Lepidiota picticollis 331 Lepidiota rothei 332 Lepidiota sororia 332 Lepidiota squamulata 333 Lepidogryllus parvulus 159, 419
Lepidogryllus comparatus 159 Leptocorisa acuta 194, 293 Leptopius maleficus 324 lesser armyworm 83 lesser budworm 89, 275 lesser ghost moth 408 lesser grain borer 45 leucaena flower eating caterpillar 366 leucaena psyllid 365 leucaena seed bruchid 366 Leucania convecta 30, 38, 130, 153, 240, 290, 295, 300, 337, 361 Leucania loreyi 35, 337 Leucania separata 33, 240, 295, 300, 337 Leucania stenographa 35, 240, 300, 337 Liatongus militaris 478 lightbrown apple moth 82 Limotettix incertus 345 linear bug 321 Lipaphis erysimi 149 Liparetrus sp. 404 Liposcelis spp. 43 Listroderes difficilis 144 Listronotus bonariensis 396 Listronotus setosipennis 385 Litomastix sp. 43, 253 little evening cricket 419 Lixus cardui 463 Locusta mirgratoria 30, 96, 316, 490 Lolium perenne 28, 321, 356, 360, 396, 406 Lolium rigidum 28, 394, 398, 417, 419 long soft scale 365 longheaded flour beetle 55 Longitarsus echii 453 Longitarsus flavicornis 456 lucerne aphid parasite 428 lucerne crown borer 211, 432 lucerne flea 13, 28, 140, 165, 263, 417, 427 lucerne leafhopper 181, 429 lucerne leafroller 223, 434 lucerne seed wasp 435 lucerne seed web moth – see etiella moth Lycosa spp. 104 lynx spiders 103 Lysiphlebus testaceipes 74 Maconellicoccus hirsutus 86 Macrolobalia ocellata 491 Macroptilium atropurpureum 194, 195, 214, 215, 246, 357 Macroptilium lathyroides 186, 194, 195, 204, 212, 220 Macrosiphum euphorbiae 149 Macrotona modesta 496 Macrotona spp. 496 maculate weevil – see spotted vegetable weevil maize leafhopper 123
516
030701•Pests of Field Crops and 516 516
29/6/07 17:40:04
INDEX
maize thrips 125 maize weevil 48 Malameba locustae 501 Malva spp. 72, 74, 76, 81, 82, 84, 98, 200, 346, 424 Mampava rhodoneura 361 Maravalia cryptostegiae 391 Marrubium vulgare 446 marshmallow 72, 74, 76, 81, 82, 84, 98, 200, 346, 424 Maruca vitrata 224 Mastotermes darwiniensis 309 Mattesia dispora 59 mealybug ladybird 108, 181 Mediterranean flour moth 58, 352 Megacyllene mellyi 374 Megymenum affine 192 Melanacanthus scutellaris 194 Melanagromyza sojae 220 Melanaphis sacchari 316 Melangyna viridiceps 150 Meligethes planiusculus 454 Mermis spp. 501 Merophyas divulsana 223, 434 mesquite leaf-tying moth 382 mesquites 382 Metanastes vulgivagus 321 Metarhizium anisopliae 316, 324, 358, 403, 488, 491, 493, 495, 498, 500 Metarhizium sp. 30, 315 Metopolophium dirhodum 17, 28 Micraspsis frenata 108 Microctonus aethiopoides 422 Micromus tasmaniae 107, 428 Microplitis demolitor 111, 246 Mictis profana 191 migration of native budworm 93 migrations of cereal pests 38 migratory locust 30, 96, 316, 490 Milax gagates 11, 137 milky disease 324, 405 Mimosa invisa 373 mimosa psyllid 373 Mimosestes ulkei 383 minimum tillage and insects 24, 139, 140, 142, 143, 263 minute mould beetles 51 mite-eating ladybirds 108, 427 Mocis alterna 248 Mocis frugalis 250, 337 Mocis trifasciata 249 Mogulones geograghicus 452 Mogulones larvatus 451 mole crickets 310, 395 Monistria concinna 493 Monistria discrepans 492 Monistria pustulifera 492 monolepta – see redshouldered leaf beetle Monolepta australis 87, 129, 216 morning glory vines 70, 74 mould mite 43 mulga grasshoppers 491
mustard 146, 346 Mythimna convecta – see Leucania convecta Myzus persicae 72, 74, 149, 151, 345 Nabis kinbergii 86, 105, 185, 245 Nabis tasmanicus 274 Nala lividipes 122, 176, 298 Nambour canegrub 330 native budworm 37, 89, 152, 161, 166, 243, 271, 350, 435 native rosella 70 Naupactus leucoloma 129, 218, 335, 431 Naupactus peregrinus 431 negatoria canegrub 330 Neogalea sunia 381 Neomolgus capillatus 418 Nephotettix malayanus 292 Netelia producta 274 Neumichtis niderrima 148 Neumichtis saliaris 148 Neumichtis spumigera 148 New Guinea weevil borer – see sugarcane weevil borer Nezara viridula 98, 127, 198, 290, 347 Nicotiana spp. 200, 348 nightfeeding sugarcane armyworm 35, 337 Nilaparavata lugens 292 Nisotra sp. 81 Nodaria externalis 238 nodding thistle 448 nodding thistle receptacle weevil 448 nodding thistle rosette weevil 449 nodding thistle seed fly 450 Nomuraea rileyi 240, 258 noogoora burr 200, 373, 382 noogoora burr longicorn 382 northern armyworm 33, 240, 295, 300, 337 northern rough bollworm 101 northern sandy dung beetle 477 Nosema heterosporum 59 Nosema locustae 500, 502 Nosema oryzaephili 59 Nosema plodiae 59 notonectid backswimmer 293 noxia canegrub 330 NPV – see nuclear polyhedrosis virus nuclear polyhedrosis virus 237, 240, 242, 245, 246, 253, 273, 274, 275, 301, 362, 435 Nupserha vexator 382 Nymphula depunctalis 294 Nysius clevelandensis 151, 160 Nysius turneri 160 Nysius vinitor 78, 151, 159, 190, 301 oat aphid 15, 16, 17, 28, 72 Octotoma championi 380 Octotoma scabripennis 380 Oechalia schellenbergii 33, 200, 245, 274
Oedaleus australis 489 Oidoematophorus balanotes 376 Oligonychus zanclopes 315 Omiodes diemenalis 230 Oncocoris coelebs 202 Oncocoris hackeri 202 Oncopera alboguttata 359 Oncopera brachyphylla 359 Oncopera fasciculata 409 Oncopera intricata 406 Oncopera mitocera 359 Oncopera rufobrunnea 406 Oncopera tindalei 359 onion thrips 78, 86, 152, 206, 268, 347, 429 Onitis alexis 478 Onitis aygulus 475 Onitis caffer 476, 478 Onitis pecuarius 479 Onitis viridulus 479 Onopordum acanthium 461 Onopordum acaulon 461 onopordum capitulum weevil 462 onopordum crown weevil 463 Onopordum illyricum 461 onopordum rosette moth 464 onopordum stemboring weevil 463 Onthophagus atrox 481 Onthophagus australis 482 Onthophagus binodis 476 Onthophagus consentaneus 481 Onthophagus ferox 480 Onthophagus gazella 479 Onthophagus granulatus 483 Onthophagus incanus 482 Onthophagus muticus 482 Onthophagus pentacanthus 481 Onthophagus quadripustulatus 482 Onthophagus taurus 476 Onychiurus spp. 164 Ophiomyia lantanae 381 Ophiomyia phaseoli 219 Opogona glycyphaga 311 Opuntia spp. 388 orbweavers 103 Orgyia australis 236 Oriental grassroot aphid 316 Orius spp. 80, 86, 106, 206 Orosius argentus 75, 345, 429 Oryzaephilus surinamensis 51 Othnonius batesii 18 Oulema rufotincta 356 Oxycanus antipoda 408 oxycanus grassgrub 408 Oxycarenus luctuosus 87 Oxyopes spp. 103 paddy bug 194, 293 paddymelons 72 Paddy’s lucerne 72 Paenibacillus popilliae 324, 405 painted pine moth 236 pale cotton stainer 96
5 17
030701•Pests of Field Crops and 517 517
29/6/07 17:40:04
INDEX
pale pea blue butterfly 235 Panicum maximum 323, 329, 336 Pantydia capistrata 250 Pantydia metaspila 250 Pantydia sparsa 250 Paraplonobia sp. 174 parkinsonia 382 Parkinsonia aculeatea 382 parkinsonia leaf bug 383 parkinsonia seed beetles 383 Parnara sp. 295 Parnkalla muelleri 312 Parthenium hysterophorus 384 parthenium leaf-mining moth 387 parthenium seed-feeding weevil 385 parthenium stem-boring weevil 385 parthenium stem-galling moth 373, 382, 386 parthenium weed 384 paspalum 321, 336, 359, 363, 398 Paspalum spp. 321, 336, 359, 363, 398 passionvine bug 192 pasture day moth 424 pasture planthopper 356 pasture snout mite 395, 418 pasture tunnel moths 410 pasture webworms 26, 359 Paterson’s curse 451 Paterson’s curse crown weevil 451 Paterson’s curse flea beetle 453 Paterson’s curse leafminer 456 Paterson’s curse pollen beetle 454 Paterson’s curse root weevil 452 pea aphid 72, 266, 420, 428 pea blue butterfly 236 pea weevil 261, 268, 273 Peakesia hospita 485 peanut mealybug – see hibiscus mealybug peanut mite 175 peanut trash bug 189 Pectinophora gossypiella 97, 100 Pectinophora scutigera 100 Pennisetum alopecuroides 361 Pennisetum clandestinum 356, 359, 360, 361, 363, 398 Penthaleus falcatus 11, 262, 394 Penthaleus major 11, 164, 262, 394 Penthaleus tectus 11, 262, 394 Penthobruchus germaini 383 perennial ryegrass 28, 321, 356, 360, 396, 406 Perkinsiella saccharicida 319 Perkinsiella thompsoni 319 Persectania dyscrita 36 Persectania ewingii 35 Petrobia latens 28 Phaenacantha australiae 321 phasey bean 186, 194, 195, 204, 212, 220 Phaulacridium vittatum 498 Pheidole anthracina 344 Phenacoccus parvus 379 Phenacoccus solani 86
Philobota productella 410 Philobota spp. 410 Phorocerocoma postulans 498 Phthorimaea operculella 349 Phyllotocus bimaculatus 404 Phyllotocus macleayi 404 Phyllotocus nigripennis 404 Phyllotocus rufipennis 404 Physalis spp. 72, 349 Phytoseiulus persimilis 427 picticollis canegrub 331 pie-dish beetle 23, 141 Pieris rapae 153 Piezodorus oceanicus 100, 132, 203 Pimelopus nothus 404 pink bollworm 97, 100 pink cutworm – see brown cutworm pink ground pearl 317 pink sugarcane mealybug 318 pinkspotted bollworm 100 pirate bugs 80, 86, 106, 206 plague soldier beetle 211 plague thrips 207, 268, 429 planiceps canegrub 331 Platyptilia isodactyla 457 Platyzosteria sp. 156 Plautia affinis 99, 197 Plodia interpunctella 56 Plutella xylostella 146 pod weevil 217 poinsettia 70 pointed snail 9, 136 Polygonum sp. 84 Porphyrosela aglaozona 221 Portulaca spp. 226 potato aphid 149 potato moth 349 potato virus Y 345 Praxibulus duplex 501 Praxibulus eurobodallae 501 Praxibulus insolens 501 Praxibulus stali 501 predator:pest ratio 112 predatory shield bug 33, 200, 245, 274 prickly acacia 387 prickly acacia seed beetle 387 prickly pear cochineals 388 prickly pears 388 Prietocella barbata 10, 136, 426 Pristhesancus spp. 104, 246 Pristomeris sp. 424 Promargarodes australis 317 Prosopis spp. 382 pruinose scarab 404 Pseudoheteronyx basicollis 156, 208 Pseudoheteronyx sp. 128 pseudoscorpions 59 psocids 43 Psoralea tenax 197, 222 Pternemobius regulus 419 Pterohelaeus alternatus 23, 81, 141, 157, 298 Pterohelaeus darlingensis 23, 80, 141
Pterohelaeus sp. 125, 157, 211, 263 Pterolocera amplicornis 29 Pteromalus puparum 154 Pteromalus semotus 59 Ptochostola microphaella 26, 360 Pyemotes tritici 59 Pyemotes ventricosus 59 pygmy cricket 419 pyrgomorph grasshoppers 492 ragwort 456 ragwort flea beetle 456 ragwort plume moth 457 ragwort stem and crown boring moth 458 Raphanus raphanistrum 146, 150, 200, 204, 263, 346 rat’s tail fescue 501 ratoon shoot borer 315 rattle pods 70, 178, 186, 194, 195 200, 204, 228, 229, 230, 235, 237, 334 red and blue beetle 110 redbanded shield bug 100, 132, 203 redheaded flea beetle 81 redheaded pasture cockchafer 401 redlegged earth mite 12, 28, 139, 141, 164, 261, 415, 426, 512 redshouldered leaf beetle 87, 129, 216 refuges for beneficials 113 reticulated slug 11, 137, 164 Rhabdoscelus obscurus 324 Rhinacloa callicrates 383 Rhinocyllus conicus 448 Rhodes grass 360 rhopaea canegrub 331 Rhopaea magnicornis 331 Rhopalomyia californica 375 Rhopalosiphum maidis 14, 16, 17, 28, 72, 127, 299, 317 Rhopalosiphum padi 15, 16, 17, 28, 72 Rhopalosiphum rufiabdominalis 290 Rhynchosia spp. 70, 229 Rhyparida dimidiata 333 Rhyparida morosa 333 Rhyparida spp. 130 Rhyzopertha dominica 45 rice leaffolder 294 rice leafhopper 292 rice leafminer 288, 292 rice root aphid 290 rice skipper 295 rice stink bug 205, 290, 293 rice weevil 46 ricespotting bug 205, 293 Ricinus communis 200 Riptortus serripes 193 robust fipurga 501 root feeding springtails 164 rose-grain aphid 17, 28 Rothe’s canegrub 332 rough bollworm 97, 101 rough brown weevil 358 roundheaded pasture webworm 359
518
030701•Pests of Field Crops and 518 518
29/6/07 17:40:05
INDEX
rubber vine 391 rubber vine leaf rust 391 rubber vine leaf-feeding moth 391 Rumex crispus 442 Rumex obtusifolius 442 Rumex pulcher 442 Rumex spp. 424 rust red flour beetle 53 Rutherglen bug 78, 151, 159, 190, 301 rye 360 ryegrass mosaic virus 28 Saccharicoccus sacchari 318 Salbia haemorrhoiodalis 382 saltbushes 424 Salticidae 103 sandgroper 13, 395 Saragus sp. 22, 81, 141 Saulostomus villosus 402, 404 sawtoothed grain beetle 51 Scelio chortoicetes 495 Scelio fulgidus 488 Scelio gobar 491, 498, 503 Scelio improcerus 500 Scelio parvicornis 488, 495 sciarid flies 219 Scirpophaga innotata 294 Scitala sericans 402, 404 Scolothrips sexmaculatis 79 Scopula perlata 233 Scorpionida 59 Scotch thistle 461 Scrobipalpa heliopa 348 Secale cereale 360 Sechium edule 194 seedharvesting ants 344 Senecio jacobaea 456 Senna sp. 194, 334 Sericesthes micans 404 Sericesthis consanguinea 18 Sericesthis geminata 404 Sericesthis harti 18 Sericesthis ino 209 Sericesthis nigra 404 Sericesthis nigrolineata 402, 403 Sericesthis spp. 128, 208 Sericesthis suturalis 209 Sericothips staphylinus 445 Sesbania cannabina 194, 204, 212, 225 sesbania pea 194, 204, 212, 225 Setaria spp. 361 shiny pasture scarab 402, 404 Sida rhombifolia 72 Sidnia kingbergii 182 silverleaf whitefly 66, 158, 176 Silybum marianum 98 Simosyrphus grandicornis 150 Simplicia caeneusalis 238 siratro 194, 195, 214, 215, 246, 357 Sisymbrium orientale 146, 346 Sisyphus rubrus 480 Sisyphus spinipes 480 Sitona discoideus 422, 431
sitona weevil 422, 431 sitona weevil parasite 422 Sitophilus granarius 47 Sitophilus oryzae 46 Sitophilus zeamais 48 Sitotroga cerealella 55 six-spotted thrips 79 skeleton weed 460 slugs 11, 137, 143, 164, 356 small brown bean bug 194 small lucerne weevil 144, 430 small pasture scarab 404 small plague grasshopper 30, 290, 493 small pointed snail 10, 136, 426 Smicronyx lutulentus 385 Sminthurus viridis 13, 28, 140, 165, 263, 417, 427 sod webworm 26, 360 Solanum capsicoides 349 Solanum nigrum 349 Sonchus spp. 66, 70, 84, 263 sorghum head caterpillar 231, 302 sorghum midge 301 sororia canegrub 332 southern armyworm 35 southern dung beetle 482 southern false wireworm 23, 81, 298 southern one-year canegrub 332 southern pyrgomorph 493 southern sandy dung beetle 475 sow thistles 66, 70, 84, 263 soybean aphid 179 soybean leaf miner 221 soybean looper 251 soybean moth 221 soybean podfly 220 spear thistle 465 spear thistle gall fly 465 Sphenarches anisodactylus 223 Sphenarches zanclistes 223 spinetailed weevil 20, 57, 144 spinifex grasshoppers 496 spiny snout mite 418 Spodoptera exempta 130, 295, 300, 337, 363 Spodoptera exigua 83 Spodoptera litura 83, 240, 349 Spodoptera mauritia 295, 363 Spoladea recurvalis 226 spotted alfalfa aphid 72, 420, 428 spotted alfalfa aphid parasite 428 spotted leafhopper 75, 429 spotted vegetable weevil 21, 144 springtails 103, 165 spur-throated locust 106, 127, 316, 345, 496 squamulata canegrub 333 St John’s wort 466 St John’s wort aphid 467 St John’s wort leaf beetle 467 St John’s wort midge 468 St John’s wort stunt mite 466 Stegobium paniceum 44
stemless thistle 461 Stenocorynus spp. 311 stenocorynus weevils 311 Stethorus nigripes 427 Stethorus spp. 108 storksbill 166, 346, 424 striate false wireworm 23, 81, 141, 298 striped ladybird 108 striped mealybug 86 striped riceborer 294 stripewinged meadow grasshopper 501 subterranean clover red leaf virus 267 sugarcane and maize stemborer 291, 336 sugarcane aphid 316 sugarcane armyworm 35, 240, 300, 337 sugarcane bud moth 311 sugarcane butt weevil 324 sugarcane froghopper 320 sugarcane leaf beetle 333 sugarcane looper 250, 337 sugarcane mosaic virus 317 sugarcane planthoppers 319 sugarcane soldier fly 313, 359 sugarcane spider mite 315 sugarcane weevil borer 324 sugarcane wireworm 80, 124, 125, 210, 310 swamp foxtail 361 swarming leaf beetles 130 symphylans 307 tadpole shrimp 285 Taylorilygus pallidulus 76, 183 Tectocoris diophthalmus 97 Telenomus cyrus 198 Telenomus sp. 103, 110 Teleogryllus commodus 66, 159, 175, 309, 419 Teleogryllus oceanicus 159, 309, 419 Teleonemia scrupulosa 380 Telura sp. 404 Telura vitticollis 404 Temclucha sp. 222, 223 Tetraneura nigriabdominalis 316 Tetranychus lintearius 444 Tetranychus urticae 83, 132, 175, 344, 426 Theba pisana 9, 136, 261 Therioaphis trifolii 72, 420, 428 thistle aphid 151 thornapples 72, 74 threebanded ladybird 108 thrips 78, 86, 196, 125, 152, 187, 206, 268, 347, 427, 429, 445 Thrips imaginis 207, 268, 429 Thrips tabaci 78, 86, 152, 206, 268, 347, 429 thrips-transmitted diseases 207 Thysanoplusia orichalcea 251 TIMERITE® 13, 139, 262, 416, 451, 453, 454, 455, 456
519
030701•Pests of Field Crops and 519 519
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INDEX
Tindale’s grassgrub 359 tobacco beetle 44, 351 tobacco looper 149, 254, 350 tobacco moth – see Mediterranean flour moth tobacco stemborer 348 tobacco thrips – see onion thrips tobacco yellow dwarf virus (TYDV) 346 tomato spotted wilt virus (TSWV) 207 tomato thrips 78, 86, 152, 207, 268, 429 tortrix – see lightbrown apple moth Toya sp. 356 transverse ladybird 108 trap crops 240, 244, 275, 324 Trialeurodes vaporariorum 67, 158 Trianthema postulacastrum 226 Tribolium castaneum 53 Tribolium confusum 54 Trichogramma carverae 110 Trichogramma evanescens 59 Trichogramma pretiosum 110, 113 Trichogramma sp. near ivalae 273 Trichogramma spp. 273 Trichopoda giamocellii 202 Trichopsidea oestracea 488, 501 Trichosirocalus briesei 463 Trichosirocalus mortadelo 449 Trigonoides hyppasia 255 Triops australiensis 285 Trioxys complanatus 428 Trirhabda baccharidis 375 Trissolcus basalis 202 Trissolcus oenone 198 Trogoderma granarium 49 Trogoderma variabile 48 tropical warehouse moth 57 tubular black thrips 427 Tucumania tapiacola 390 turf 24, 318, 328, 360, 396, 406, 407 turnip aphid 149 tussock moths 236 twig caterpillar 233 two-spotted ladybird 108
two-spotted mite 83, 132, 175, 344, 426 Typhaea stercorea 53 Tyrophagus putrescentiae 43 Tytthus chiensis 185, 186, 188 Tytthus spp. 320 Ulex europaeus 444 underground grassgrub 409 Urnisa guttulosa 385 Urophora solstitialis 450 Urophora stylata 465 Uroplata fulvopustulata 380 Uroplata girardi 380 Utetheisa lotrix 237 variable ladybird 108 variegated ragweed beetle 372, 384 variegated thistle 98 vegetable leafhopper 74, 181, 429 vegetable looper 254 vegetable weevil 144 veldt grass 408 verbena 76 Verbena spp. 76 vineyard snail – see common white snail viper’s bugloss 451 Volpa bromoides 501 Voriella uniseta 435 wallaby ear 124 warehouse beetle 48 wart-eye mite damage 308 water scorpion 293 water snails 283 weed web moth – see cotton webspinner western dung beetle 480 western flower thrips 78, 86, 207, 268 Westringia fructicosa 493 wheat aphid – see oat aphid wheat curl mite 28 wheat root scarab 18 wheat streak mosaic virus 28
Wheeleria spilodactylus 447 white collared ladybird 108 white ground pearl 317 white grubs (scarab larvae) 18, 129, 322, 356, 397 white Italian snail 9, 136, 261 white rice stemborer 294 whitefringed weevils 129, 218, 335, 431 wild radish 146, 150, 200, 204, 263, 346 wild tobacco 200, 348 wild turnips 74, 84, 98, 200 wingless cockroaches 156 wingless grasshoppers 498, 501 winter corbie 406 wireweed 84 wireworms (Elateridae) 21, 80, 124, 125, 143, 210, 310, 315, 324 wireworms as predators 124, 126, 315, 324, 415 wolf spiders 104 Xanthium pungens 200 Xanthium spinosum 373 Xylocoris flavipes 59 yabby 286 Yarrita pakaria 419 yellow peach moth 101, 133, 303 yellow shouldered dung beetle 478 yellow soldier fly 313 yellow sugarcane cicada 312 yellowbellied wingless grasshoppers 501 yellowheaded cockchafer 18 yellow-winged locust 316, 502 Yrrhapta spp. 485 Zabrala ceripes 485 Zeteticontus utilis 59 Zeuxidiplosis giardi 468 Zizina labradus 234 Zoophthora radicans 435 Zygogramma bicolorata 372, 384 Zygrita diva 212, 432
520
030701•Pests of Field Crops and 520 520
29/6/07 17:40:07
