Table of Contents

Table of Contents

Authors, Reviewers, Draft Log ...... 4 Introduction to Reference ...... 5 Introduction to Corn ...... 10

Arthropods ...... 12 Primary Pests of Corn (Full Datasheet) ...... 12 Autographa gamma ...... 12 suppressalis ...... 21 Diabrotica speciosa ...... 30 armigera ...... 39 Heteronychus arator ...... 52 Ostrinia furnacalis ...... 61 Spodoptera littoralis ...... 73 Spodoptera litura ...... 83 albicosta ...... 95 Thaumatotibia leucotreta ...... 105 Secondary Pests of Corn (Truncated Pest Datasheet)...... 116 Copitarsia spp...... 116 ...... 123 Eutetranychus orientalis ...... 128 Metamasius spp...... 132 ...... 138 Planococcus minor ...... 142 Rhabdoscelus obscurus ...... 146 Stenchaetothrips biformis ...... 153

Mollusks ...... 158 Primary Pests of Corn (Full Pest Datasheet) ...... 158 None at this time ...... 158 Secondary Pests of Corn (Truncated Pest Datasheet)...... 158 Achatina fulica ...... 158 vulgaris (A. lusitanicus) ...... 163

Nematodes ...... 170 Primary Pests of Corn (Full Pest Datasheet) ...... 170 Punctodera chalcoensis ...... 170 Secondary Pests of Corn (Truncated Pest Datasheet)...... 180 Meloidogyne fallax ...... 180

Plant Pathogens ...... 186 Primary Pests of Corn (Full Pest Datasheet) ...... 186

2 Table of Contents

Cochliobolus pallescens ...... 186 Harpophora maydis ...... 194 Peronosclerospora maydis ...... 203 Peronosclerospora philippinensis...... 209 Sclerophthora rayssiae var. zeae ...... 216 Secondary Pests of Corn (Truncated Pest Datasheet)...... 222 None at this time ...... 222

Appendix A: Characteristics of the downy mildew fungi found on corn and other hosts ...... 222

Appendix B: Diagnostic Resource Contacts ...... 224

Appendix C: Glossary of Terms ...... 226

Appendix D: FY09 CAPS Prioritized Pest List and Commodity Matrix ...... 240

Appendix E: FY10 CAPS Prioritized Pest List and Commodity Matrix ...... 243

Appendix F: 2008 Exotic Pest Detection Survey Order Form ...... 247

Appendix G: Plastic Bucket Trap Protocol ...... 249

3 Authors, Reviewers, Draft Log

Authors, Reviewers, Draft Log

Authors CAPS Commenters Melinda Sullivan Nancy S. H. Richwine Pathologist Plant Pathologist USDA-APHIS-PPQ-CPHST PA Department of Agriculture 2301 Research Blvd., Suite 108 2301 N. Cameron St. Fort Collins, CO 80526 Harrisburg, PA 17110-9408

Daniel MacKinnon Nancy K. Osterbauer Biological Science Technician Plant Health Program Manager USDA-APHIS-PPQ-CPHST Commodity Inspection Division 2301 Research Blvd., Suite 108 Oregon Department of Agriculture Fort Collins, CO 80526 635 Capitol St. NE Salem, OR 97301-2532 Talitha Price Biological Science Technician John F. Tooker USDA-APHIS-PPQ-CPHST Assistant Professor 1730 Varsity Dr., Suite 400 Center for Chemical Ecology/ Raleigh, NC 27606 Department of Entomology State University Robert J. Wright 501 ASI Building Professor of Entomology University Park, PA 16802 202 Entomology Hall University of Nebraska-Lincoln Lincoln NE 68583-0816 Draft Log

Feb. 2010– Draft sent for CPHST review Tamra J. Jackson

Assist. Professor of Plant Pathology April 2010 – Draft sent for CAPS review 448 Plant Science Bldg.

University of Nebraska-Lincoln June 2010 – Final Draft posted to CAPS site Lincoln NE 68583-0722 – incorporated CAPS comments and added

CPHST Reviewers new risk maps, Added datasheet for

Diabrotica speciosa Daniel MacKinnon Biological Science Technician July 2010 – Added datasheets for USDA-APHIS-PPQ-CPHST pallescens and Striacosta 2301 Research Blvd., Suite 108 albicosta as requested by two CAPS Fort Collins, CO 80526 reviewers

Lindsey Seastone August 2010 – Updated hyperlinks for links Biological Science Technician now on the CAPS Resource and USDA-APHIS-PPQ-CPHST Collaboration site; approved trap information 2301 Research Blvd., Suite 108 added for Chilo suppressalis Fort Collins, CO 80526 August 2016 – Removed outdated mapping Christina Southwick information, did not update other information. Identification Technology Program Technician USDA-APHIS-PPQ-CPHST 2301 Research Blvd., Suite 108 Fort Collins, CO 80526

4 Introduction to Reference

Introduction to Reference

History of Commodity-Based Survey The CAPS community is made up of a large and varied group of individuals from federal, state, and university organizations who utilize federal (and other) funding sources to survey for and (in some cases) diagnose exotic and invasive plant pests. By finding pests early, eradication efforts will likely be less expensive and more efficient. For more information on CAPS and other Plant Protection and Quarantine (PPQ) pest detection programs see: http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/index.shtml.

Traditionally, states have been given a list of pests. Each year, states choose (from this list) a number of pests to incorporate in their own specialized surveys. There is certainly value in surveying for plant health threats in terms of discreet pests. However, this may not always be the most efficient means of survey. For example, a single pest may occur on a myriad of different hosts, making a comprehensive survey too time consuming and expensive. An alternative method has been suggested. Grouping important pests under the umbrella of a single commodity could be a more efficient way to look for certain pests. The rationale for choosing a commodity survey in certain instances includes the following:

• Survey area will be smaller and targeted. • Resources can be better utilized with fewer trips to the field. • Commodities are easy to prioritize in terms of economic and regional (geographic) importance.

The Center for Plant Health Science and Technology (CPHST) has been charged to develop a commodity-based survey strategy in support of the CAPS program. There are two types of end products being developed for each commodity. Each product serves a valuable yet unique purpose. The result is a set of paired documents developed for each commodity. A description of these documents is provided below:

Commodity-Based Survey Reference (CSR): This document is composed of a series of pest data sheets, mini-pest risk assessments (PRAs), or early detection PRAs. The data sheets are highly graphic and illustrate the biology, survey, and identification of particular pests in appropriate detail for CAPS surveyors. The pests in this document are numerous. The pests were chosen primarily from the CAPS Analytic Hierarchy Process (AHP) prioritized pest list [Appendix D and E] and the Select Agent list (http://www.selectagents.gov/ or http://www.aphis.usda.gov/programs/ag_selectagent/ag_bioterr_toxinslist.html). Additional pests may be added if they are cited in the literature as being a primary pest of the given commodity and are exotic to the , or if specifically requested by the CAPS National Committee. States are not required to survey for all of the pests

5 Introduction to Reference in this document, but may choose those that are particularly relevant to include in their survey. In general, this document should serve as a desk reference for survey specialists as they plan their cooperative agreements. It may also be useful for obtaining high quality scientific information quickly during the field season.

Commodity-Based Survey Guidelines (CSG): This document is smaller. The list of pests is shorter than those chosen for the CSR. A subgroup of the CAPS National Committee determines which pests from the CSR will be included in the CSG. As such, states that participate in these surveys must survey for all organisms listed in the CSG that are relevant to the state in terms of risk. The CSG set forth guidelines for survey and identification from a broad scale (site selection, number of acres to survey, number of samples to collect, etc.) and a narrow scale (field methods, survey tools, transporting samples, etc.). States are encouraged to follow the procedure set forth in the CSG. The methods are intended to increase the homogeneity of the national data set and increase the statistical confidence in negative data (e.g., demonstration of “free from” status).

As a pilot project, citrus was undertaken as the first commodity in this initiative. The products were developed for implementation in the 2007 survey season. Citrus was chosen, because it is an economically important commodity that is equally distributed in both PPQ regions but is distributed in few overall states. To date, survey strategies for pests of citrus are also well documented. Shortly after completion of the citrus CSG, several other commodity survey guidelines were initiated, including soybean, , grape, potato, small grains, stone fruit, and oak and pine forests.

Corn Commodity Survey Reference The Corn Commodity-based Survey Reference (CSR) is a companion document to the Corn Commodity-based Survey Guidelines (CSG). Both documents are intended to be tools to help survey professionals develop surveys for exotic pests of corn. The Corn CSR is a collection of detailed data sheets on exotic pests of corn. Additionally, the authors have identified native pests that may be easily confused with these exotic pests as well as potential vectors of exotic pests. These data sheets contain detailed information on the biology, host range, survey strategy, and identification of these pests. The commonly confused pests and vectors are included in a section of the pest data sheet dealing with the target pest.

By comparison, the Corn CSG companion document is intended to help states focus resources on survey efforts and identification of a smaller group of target pests (usually less than a dozen). The Corn CSG contains little information about biology. Instead, the guideline focuses on survey design, sampling strategies, and methods of identification. There is no single survey that would be wholly applicable to each location in the United States. Environment, personnel, budgets, and resources vary from state to state. Thus, the Corn CSG will provide a template that states can use to increase the uniformity and usability of data across political, geographic, and climatic regions while maintaining flexibility for specificity within individual regions.

6 Introduction to Reference

Purposes of the Corn CSR • To relate scientific information on a group of threatening pests. • To facilitate collection of pest data at a sub-regional, regional, and national level versus data collection from a single location. • To aid in the development of yearly surveys. • To help CAPS cooperators increase their familiarity with exotic pests and commonly confused pests that are currently found in a given commodity. • To aid in the identification and screening of pests sampled from the field. • To collate a large amount of applicable information in a single location. End Users As previously noted, this document may be used for many purposes. Likewise, it will be of value to numerous end users. As the document was developed, the authors specifically targeted members of the CAPS community who are actively involved in the development and implementation of CAPS surveys.

State Plant Health Director (SPHD): The SPHD is the responsible PPQ official who administers PPQ regulatory and pest detection activities in his or her state. The SPHD is also responsible for ensuring that the expanded role of CAPS is met in his or her state. In many states, the SPHD provides guidance for the state’s ongoing management of pest risk and pest detection. However, SPHD responsibilities will vary according to the extent to which each state carries out the various components of the CAPS program.

State Plant Regulatory Official (SPRO): These individuals are employees of their respective states and generally manage the expanded survey program. The SPRO is the responsible state official who administers state agricultural regulatory programs and activities within his or her respective state.

Pest Survey Specialists (PSS): The PSS, a PPQ employee, is supervised by the SPHD of the state in which he or she is assigned. A PSS may also be responsible for survey activities and may work with the SSC and the survey committee in more than one state.

State Survey Coordinators (SSC): The SSC is a state employee responsible for coordinating each state’s CAPS program, participating as a member of the state CAPS committee (SCC), and acting as liaison with the state PPQ office.

Diagnosticians: Diagnostic capabilities vary by state. Some states have advanced networks of diagnosticians, whereas other states access diagnostic support through National Identification Services (NIS) or through contracts with external partners. States are encouraged to utilize qualified diagnosticians in their respective states if expertise is available. PPQ offers diagnostic support for the CAPS program through NIS. A major

7 Introduction to Reference responsibility for NIS’s Domestic Identifiers is to provide diagnostic support to CAPS programs. There are plant pathology and entomology domestic identifiers in each of the regions. A Forest Entomology Domestic Identifier oversees both regions. To learn more about diagnostic resources available to you, discuss your diagnostic requirements and options with your State Plant Health Director, one of the regional Domestic Identifiers, and/or NIS. Appendix B has a listing of NIS and Domestic Identifier contact information.

Organisms Included in the Corn Survey Reference Organisms included in the corn survey reference are organized first by:

1. Pest type, (e.g., , plant pathogens, , and mollusks).

2. Organisms are then divided by their pest status on corn [e.g., primary pest (major pest) and secondary (minor pest)]. Primary and secondary is determined by reviewing the literature, host association, yield loss, and etc. associated with the pest on a given commodity

A. Primary Pests: Full pest datasheets will be developed for primary pests. All pests must be exotic to the conterminous United States.

• Pests found on the AHP Prioritized Pest List (for the fiscal year of interest) and that are major pests on the commodity will be considered primary pests.

• Additional exotic pests that the author finds in the literature that are major pests on the commodity will be included as primary pests and given the designation of “National threat”.

B. Secondary Pests: Truncated pest datasheets will be developed for secondary pests.

• Pests found on the AHP Prioritized Pest List (for the fiscal year of interest) that are not identified as major pests of the commodity in the literature.

C. PPQ Program and Line Item Pests: Plant Protection and Quarantine Program pests and pests with their own line item funding should be listed by scientific name and common name only. These pests will not receive pest datasheets, unless specifically requested by the National CAPS Committee. If a PPQ website exists for the pest, a link should be provided to that site. CPHST Ft. Collins can assist in determining which program pests and line item pests are relevant to the commodity.

D. Other Pests Determined by the National CAPS Committee or requested by the CAPS Community: Full pest datasheets will be developed for specific pests requested by the CAPS community.

8 Introduction to Reference

3. Finally, organisms are arranged alphabetically by their scientific names. Common names are provided as well. Previous manuals have included pests from the Eastern and Western Region pest lists. The restructuring of the CAPS program and shift from regional guidelines to a single set of national guidelines has made these lists obsolete. Therefore, pests from these lists were not included in this CSR. States now have more flexibility to survey for pests of state concern, and most regional pests were captured in one or more state CAPS pest lists.

To help provide a rationale for the inclusion of each pest in the reference, the authors have included a section titled, “Reason for Inclusion in Manual”. Pests are either considered to be a CAPS target and are listed in the CAPS prioritized pest list or a national threat. The pests considered as national threats are not known to be present in the United States; however, they are not associated with the CAPS prioritized pest lists but are found on another list or identified through the literature. An additional category, requested by the CAPS community, is present in some manuals if a pest is suggested that is a primary pest, exotic to the United States, or is of regulatory significance.

Appendix M1 The survey methodology presented in Appendix M1 in the 2011 CAPS National Survey Guidelines (http://caps.ceris.purdue.edu/guidelines/2011/apdx_m1; http://pest.ceris.purdue.edu/services/napisquery/query.php?code=cam) lists the most up-to-date, CAPS-approved methods for survey and identification/diagnostics of CAPS target pests from the Priority Pest List, consisting of pests from the 1) commodity- and taxonomic-based surveys and 2) AHP Prioritized Pest List. The information in this table supersedes any survey and identification/ diagnostic information found in any other CAPS document (i.e., Commodity-based Survey References and Guidelines, EWB/BB National Survey Manual, etc.). All other CAPS documents will be revised to include the information contained in this table; however, this table should always be the authoritative source for the most up-to-date, CAPS-approved methods.

9 Introduction to Corn

Introduction to Corn

Four crops, corn, soybeans, and hay, occupy 80% of the planted crop acreage in the United States. Corn (Zea mays L.) is an important grain crop in the United States. In 1998, corn was harvested on 73 million acres with a production of about 9.7 billion bushels. In 1995, approximately 18% of the crop was used for food, seed, and industrial uses; 59% was used for feed; and 23% was exported.

Corn is widely adaptable to different environments (length of growing season, rainfall amounts, etc.). According to the United States Department of Agriculture (USDA), 86,977,000 acres of corn were planted in 2008 and 93,600,000 acres were planted in 2007. Corn production occurs across the United States with the greatest concentration in the central U.S. “Corn Belt”. The Corn Belt is a region of the Midwestern United States where corn has traditionally been the predominant crop. Geographic definitions of the region vary. Typically it is defined to include , , , and — approximately 50% of all corn grown in the United States is from these four states. The Corn Belt also includes parts of South Dakota, North Dakota, Nebraska, Kansas, , , , , and Kentucky.

Figure of the United States showing corn grain harvested by county in 2012. Map courtesy of USDA-APHIS-PPQ-CPHST. 10 Introduction to Corn

Corn growth and development can be assessed using the Iowa State University growth staging system (Ritchie et al., 1992). Stages are numbered separately for vegetative and reproductive stages.

Vegetative stages:

VE: emergence

V1 to V18: V1, one leaf emerged with leaf collar visible to V18, eighteenth leaf emerged

VT: tasseling, last branch of tassel is completely visible

Reproductive stages:

R1: silking, silks visible outside the husks

R2: blister, kernels are white and resemble a blister in shape (10-14 days after silking)

R3: milk, kernels are yellow on the outside with a milky inner fluid (18-22 days after silking)

R4: dough, milky inner fluid thickens to a pasty consistency (24-28 days after silking)

R5: dent, nearly all kernels are denting (35-42 days after silking)

R6: physiological maturity, the black abscission layer has formed (55-65 days after silking)

References cited: USDA-NASS. http://www.nass.usda.gov/.

Ritchie, R.W., Hanway, J.J., and Benson, G.O. 1992. How a corn plant develops. http://www.extension.iastate.edu/hancock/info/corn.htm. Iowa State University Special Report 48. Ames, IA.

11 Autographa gamma Primary Pest of Corn Arthropods Silver-Y Moth

Arthropods

Primary Pests of Corn (Full Pest Datasheet)

Autographa gamma

Scientific Name Autographa gamma L.

Synonyms: Phytometra gamma and Plusia gamma

Common Name(s) Silver-Y moth, beet worm A Type of Pest Moth

Taxonomic Position Class: Insecta, Order: , Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List - 2009 B Pest Description : Semi-spherical, 0.57 mm in Figure 1. Eggs (A) and larvae (B) of A. diameter. Eggs are yellowish-white (Fig. gamma. Photos courtesy of Jurgen 1A), later turning yellowish-orange to Rodeland (www.invasive.org), and brown. The number of ribs varies from 28 Horticulture Research International to 29 (Paulian et al., 1975). The eggs are Wellesbourne (CABI, 2007). deposited in bunches or singly on the underside of leaves.

Larvae: The larva is a semi-looper with three pairs of prolegs. It occurs in varying shades of green (Fig. 1B), with a dark green dorsal line and a paler line of whitish-green on each side. The spiracular line is yellowish, edged above with green. There are several white transverse lines between the yellow spiracular line and the dorsal black line. Some larval forms have a number of white spots. The head may have a dark patch below the ocelli or be entirely black. Maximum length is 20 to 40 mm (Emmett, 1980; Hill, 1983; Jones and Jones, 1984; USDA, 1986).

12 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

Pupae: Pupation takes place within a translucent, whitish cocoon spun amongst plant foliage (Fig. 2A). The leaves may sometimes be folded over. The is brown to black, greenish or even whitish-green on its ventral side, 16 to 21 mm long, and 4.5 to 6.0 mm broad. Cremaster globular, with four pairs of hooklets (Paulian et al., 1975; Carter and Hargreaves, 1986).

Adults: The adults are gray-colored and the forewings are marbled in A appearance, their color being silvery-gray to reddish-gray to black with a velvety sheen. Wing expanse is 36 to 40 mm. The ‘Y’ mark on the forewing is distinct and silvery (Fig. 2B). The hindwings are brownish with a darker border (USDA, 1958; Hill, 1983; Jones and Jones, 1984).

Biology and Ecology A. gamma is a migratory species and adults undertake seasonal migrations to areas where they are B able to breed. The silver-Y moth can be found in many habitats including agricultural land, waste land, and gardens. In areas where A. gamma is unable to overwinter, severe infestations occur sporadically.

Female take nectar from flowers and can often be seen feeding during the day or early evening. Females lay from 500 to more than 1000 whitish eggs (Hill, 1983) (Fig. 1A), singly or in small Figure 2. Cocoon (A) and adult (B) A. gamma. batches, on the underside of leaves Photos courtesy of Alain Fraval and Jeremy of low-growing . In temperate Lee, respectively. regions, hatching may take 10 to 12 days (Hill, 1983). The incubation period lasts for 3 days at 25°C (77°F) (Ugur, 1995).

The young larvae feed on the foliage of their host plants and tend to occur singly, rather than in groups. When larvae are young, they skeletonize the leaves, but older caterpillars eat the whole leaf (Hill, 1983). Larval development takes from 51 days at 13°C (55°F) to 15 to 16 days at 25°C and the pupal stage from 32 days at 13°C to 6 to 8 days at 25°C (Hill and Gatehouse, 1992; Ugur, 1995). When the larvae are disturbed,

13 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

they drop off the plant.

Local distribution, reproductive potential, and migration are determined to a considerable extent by the availability of suitable wild plants in a given area, and good weed control reduces the threat of outbreaks. Mortality in the stage and the first larval instar is lowest at high humidity levels; mass outbreaks are known to have occurred mainly during periods of very wet weather (Maceljski and Balarin, 1974).

In areas where A. gamma is able to survive the winter, it overwinters as third to fourth larval instars (Tarabrina, 1970; Kaneko, 1993; Saito, 1988) or in the pupal stage (Dochkova, 1972). There is no true diapause (Tyshchenko and Gasanov, 1983).

Symptoms/Signs Leaves may be skeletonized by young larval feeding, leaving plants with a brownish appearance. Older leaves are preferred by larvae. The petioles or leaf stalks may be cut by the larvae. Older larvae eat from the edge of the leaf towards the midrib, consuming the leaves completely or at times leaving pieces of the midrib.

Eggs (singly or in small clusters) may be visible on leaves of low growing plants. Larvae are active at night. During the day, they remain pressed against the underside of the leaf; when disturbed, they tend to drop off the plant. Frass may or may not be visible. Pupae are found in the folds of the lower leaves of the host plant. Webbing may be present. Adult moths feed in flowers (nectar) and can often be seen feeding during the day or early evening.

Pest Importance From CABI (2007): Outbreaks of A. gamma occur periodically over wide areas of Europe, , and North . The outbreak of 1928, which occurred in most of central Europe, caused widespread defoliation of peas in Poland. Damage from this and Pieris rapae (cabbage white ) in areas of the Netherlands was valued at as much as 320,000 guilders (roughly $178,591) during some years in the 1800s. It is also very destructive in England and Denmark. Damage to globe artichokes was severe near Bari, Italy in 1982 to 1985, with about 55% of plants being damaged. A. gamma was one of the major pests.

Studies in Czechoslovakia (Novak, 1975) indicated that damage became of economic significance when 25% of the leaf area of a plant was destroyed. The critical density of larvae was, therefore, the number of larvae/unit area required to do this, which varied according to both the larval instar concerned and the development stage of the plant. The numbers of larvae per plant causing 25% leaf loss varied from 0.07 when the plant had only two leaves to 20 when it had 30 leaves.

Known Hosts This polyphagous pest is found on cereals, grasses, fiber crops, Brassica spp., and other vegetables, including legumes. Corn is considered a primary host (Carter, 1984).

14 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

A. gamma can feed on at least 224 plant species, including 100 weeds, from 51 families (Maceljski and Balarin, 1972).

Major hosts Beta vulgaris (beet), Beta vulgaris var. saccharifera (sugarbeet), Borago officinalis (borage), Brassica oleracea var. capitata (cabbage), Brassica oleracea var. gemmifera (Brussels sprouts), Brassica rapa subsp. chinensis (Chinese cabbage), Brassica rapa subsp. pekinensis (Pe-tsai), Cannabis sativa (hemp), Capsicum spp. (peppers), indicum (chrysanthemum), arietinum (), Cichorium intybus (chicory), Cynara scolymus (artichoke), Daucus carota (carrot), Glycine max (soybean), Gossypium spp. (cotton), Helianthus annuus (sunflower), Hyssopus officinalis (hyssop), Lactuca sativa (lettuce), Linum usitatissimum (flax), Medicago sativa (), Nicotiana tabacum (), (geranium) hybrids, Petroselinum crispum (parsley), Pisum sativum (pea), tuberosum (potato), Spinacia oleracea (spinach), Trifolium pratense (purple ), Triticum aestivum (wheat), Vitis vinifera (grape), Zea mays (corn), and Zinnia elegans (zinnia).

Known Vectors (or associated organisms) A. gamma is not a known vector and does not have any associated organisms.

Known Distribution A. gamma is widely distributed throughout all of Europe and eastward through Asia to and ; it also occurs in North Africa (USDA, 1958).

Asia: Azerbaijan, China, India, Iran, Iraq, Israel, , Kazakhstan, Korea, , Syria, Turkey, and Uzbekistan. Europe: Austria, Belgium, Bulgaria, Former Czechoslovakia, Denmark, Finland, Former USSR, France, Germany, Greece, Hungary, Iceland, Italy, Latvia, Lithuania, Moldova, Netherlands, Poland, , Romania, Russian Federation, Serbia and Montenegro, Slovakia, Spain, Sweden, Switzerland, Ukraine, and United Kingdom. Africa: Algeria, Egypt, Libya, and Morocco.

Potential Distribution within the United States The likelihood and consequences of establishment by A. gamma have been evaluated in a pathway-initiated risk assessment. Autographa gamma was considered highly likely of becoming established in the United States if introduced. The consequences of its establishment for U.S. agricultural and natural ecosystems were also rated high (i.e., severe) (Lightfield, 1997). The risk assessment indicates that California and the southern United States have the greatest risk for A. gamma establishment based on host availability and climate within the continental United States. Establishment is precluded in many areas of the northwestern, north central, and northeastern United States.

Survey CAPS-Approved Method: Trap with lure. A plastic bucket trap (unitrap) (green canopy, yellow funnel, white bucket) with dry kill strip is the approved trap for Autographa gamma. The lure information is provided below:

15 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

Lure Dispenser Dispenser Lure Length of Compound Load Type Abbreviation Effectiveness a) 'Z,7-12:AC a) 0.99 mg gray rubber AG 4 weeks b) 'Z,7-12:OH b) 0.01 mg septum

For instructions on using the trap, see Appendix G “Plastic Bucket Trap Protocol”.

IMPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters.

Literature-Based Methods: Due to the migratory nature of this species, adult A. gamma can be observed every month from April to November, usually peaking in late summer (CABI, 2007).

Trapping: The sex , (Z)-7-dodecenyl acetate and (Z)-7-dodecenol in ratios from 100:1 to 95:5 (19:1) has been used to attract and monitor male flight of A. gamma. In field applications, the pheromone may be dispensed from rubber septa at a loading rate of 1 mg. Lures should be replaced every 30 days. Newly-emerged adult males of A. gamma are not attracted to the pheromone; 3-day old males are most responsive to the lure. The pheromone of A. gamma may also attract other Lepidoptera in the United States including: Anagrapha ampla, Anagrapha falcifera, Autographa ampla, Autographa biloba, Autographa californica, Caenurgia spp., Epismus argutanus, Geina periscelidactyla, Helvibotys helvialis, Lacinipolia lutura, Lacinipolia renigera, Ostrinia nubilalis, Pieris rapae, Polia spp., Pseudoplusia includens, Rachiplusia ou, Spodoptera ornithogalli, Syngrapha falcifera, and Trichoplusia ni.

Trapping is suggested in major truck farming areas. Traps should be placed within or on the edge of fields of the host crops. Traps should be suspended from stakes and placed at crop height and raised as the crop matures.

Visual survey: The USDA (1986) provides some considerations for visual inspections of host plants for the presence of eggs, larvae, or pupae. In general, eggs may be found on the lower and upper surfaces of leaves. Larvae are likely to be found, if left undisturbed, on leaves that have been skeletonized or that have holes in the interior. Pupae may be found on the lower leaf surface (USDA, 1986).

Not Recommended: Adult males and females have also been collected using Robinson black-light traps, but these traps attract moths non-discriminately. Such traps, placed 3 meters above the ground, have been used to successfully monitor the dynamics of A. gamma and other Noctuid moths. Sticky traps have been used, but are not recommended as pheromone traps are much more effective.

16 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of A. gamma is by morphological identification.

Literature-Based Methods: Species are most reliably identified by close examination of the genitalia (Nazmi et al., 1980; USDA, 1986).

Easily Confused Pests Several life stages of Noctuid pests can be confused with A. gamma. Of these, the most important species include: Trichoplusia ni (cabbage looper) (Fig. 3), Syngrapha celsa (plain silver-Y or western conifer looper) (Fig. 4), A. pseudogamma (delicate silver-Y) (Fig. 5), and A. californica (alfalfa looper) (Fig. 6). All are already present in the continental United States. The other easily confused species are Cornutiplusia circumflexa (Essex Y), which is distributed in Europe, Asia, and Africa, and Syngrapha interrogationis (scarce silver Y), which is established in the United Kingdom (Venette et al., 2003). Adults of A. gamma are gray to grayish brown in color with a “Y mark or gamma [γ] on the forewing”. See Nazmi et al. (1980) for a comparison of similarities and differences between closely related species. A screening aid is available at: http://caps.ceris.purdue.edu/webfm_send/548, http://caps.ceris.purdue.edu/webfm_send/633.

Figure 3. Adult and larva of Trichoplusia ni. Photos courtesy of Keith Naylor and Extension Entomology, A&M University.

17 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

Figure 4. Adult and larva of Syngrapha celsa. Photos courtesy of John Cooper and Natural Resources Canada.

Figure 5. Adult of Autographa pseudogamma. Photo courtesy of Natural Resources Canada.

18 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

Figure 6. Adult and larva of Autographa californicum. Photos courtesy Franklin Dlott, U.C. Cooperative Extension, Monterey County.

References CABI. 2007. Crop Protection Compendium. Wallingford, UK: CAB International. http://www.cabi.org/compendia/cpc/.

Carter, D.J. 1984. Pest Lepidoptera of Europe with special reference to the British Isles. Dr. W. Junk (ed.). Kluwer Academic Publications, Boston, MA. 307-309, 353-354.

Carter, D.J. and Hargreaves, B. (Illustrator). 1986. A field guide to caterpillars of and moths in Britain and Europe. London, UK: Collins, 296 pp.

Dochkova, B. 1972. Some biological and ecological studies of the gamma moth. Rasteniev"dni Nauki 9(10): 141-149.

Emmett, B.J. 1980. Key for the identification of lepidopterous larvae infesting Brassica crops. Plant Pathology 29(3): 122-123.

Hill, D.S. 1983. Agricultural insect pests of the tropics and their control. 2nd edition. Cambridge, UK: Cambridge University Press.

Hill, J.K., and Gatehouse, A.G. 1992. Effects of temperature and photoperiod on development and pre- reproductive period of the silver Y moth Autographa gamma (Lepidoptera: Noctuidae). Bulletin of Entomological Research 82: 335-341.

Jones, F.G.W. and Jones, M.G. 1984. Pests of field crops. London, UK: Edward Arnold, 127-127.

Kaneko, J. 1993. Existence ratio of the silver Y moth, Autographa gamma (L.) after overwintering in a cabbage field at Sapporo, Japan. Annual Report of the Society of Plant Protection of North Japan No. 44: 124-126.

Lightfield, J. 1997. Importation of leaves and stems of parsley, Petroselinum crispum, from Israel into the United States: qualitative, pathway-initiated pest risk assessment. Animal and Plant Health Inspection Service, US Department of Agriculture, Riverdale, MD.

19 Autographa gamma Primary Pest of Corn Arthropods Silver-Y moth Moth

Maceljski, M. and Balarin, I. 1972. On knowledge of polyphagy and its importance for the silver-Y moth (Autographa gamma L.). Acta Entomologica Jugoslavica 8: 39-54.

Maceljski, M. and Balarin, I. 1974. Factors influencing the population density of the looper-Autographa gamma L. in Yugoslavia. Acta Entomologica Jugoslavica 10: 63-76.

Nazmi, N., El-Kady, E., Amin, A., and Ahmed, A. 1980. Redescription and classification of subfamily in Egypt. Bulletin de la Societe Entomologique d'Egypte 63: 141-162.

Novak, I. 1975. Critical number of Autographa gamma L. caterpillars (Lep., Noctuidae) on sugar-beet. Sbornik UVTI - Ochrana Rostlin 11: 295-299.

Paulian, F., Mateias, M.C., Sapunaru, T., and Sandru, I. 1975. Results obtained in the study and control of the pests of lucerne crops in the Socialist Republic of Romania. Probleme de Protectia Plantelor 3: 359-385.

Saito, O. 1988. Notes on larval hibernation of the alfalfa looper, Autographa gamma (Linnaeus) on cabbage plant in Sapporo, Hokkaido. Annual Report of the Society of Plant Protection of North Japan No. 39: 217.

Tarabrina, A.M. 1970. The silver Y moth in the Voronezh region. Zashchita Rastenii 15: 19.

Terytze, K., Adam, H., and Kovaljev, B. 1987. The use of pheromone traps for monitoring harmful Lepidoptera species on cabbage. Archiv für Phytopathologie und Pflanzenschutz 23: 465-473.

Tyshchenko, V.P., and Gasanov, O.G. 1983. Comparative study of photoperiodic regulation of diapause and weight of pupae in several species of Lepidoptera. Zoologicheskii Zhurnal 62: 63-68.

Ugur, A. 1995. Investigations on some biological aspects of Autographa gamma (L.) (Lepidoptera, Noctuidae). Türkiye Entomoloji Dergisi 19: 215-219.

USDA. 1958. Coop. Economic Insect Report 8 (23). USDA.

USDA. 1986. Pests not known to occur in the United States or of limited distribution No. 75: Silver Y Moth, pp. 1-16. APHIS-PPQ, Hyattsville, MD.

Venette, R.C., Davis, E.E., Heisler, H., and Larson, M. 2003. Mini Risk Assessment Silver Y Moth, Autographa gamma (L.)[Lepidoptera: Noctuidae]. Cooperative Agricultural Pest Survey, Animal and Plant Health Inspection Service, US Department of Agriculture. Available on line at: http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/agammapra.pdf.

20 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic borer Moth

Chilo suppressalis

Scientific name Chilo suppressalis Walker

Synonyms Jartheza simplex, Chilo oryzae, Chilo simplex, and Crambus suppressalis

Common names Asiatic rice borer, striped rice stem borer, striped rice stalk borer, rice stem borer, rice chilo, purple-lined borer, rice borer, moth borer, pale-headed striped borer, and rice stalk borer.

Type of pest Moth

Taxonomic position Class: Insecta, Order: Lepidoptera, Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010 Figure 1. Chilo suppressalis egg masses. Pest Description Image courtesy of Eggs: Eggs (Fig. 1) are fish scale-like, about 0.9 x 0.5 International Rice mm, turning from translucent-white to dark-yellow as Research Institute Archive. they mature. They are laid in flat, overlapping rows www.bugwood.org containing up to 70 eggs. Eggs of other Chilo spp. are

quite similar and can not be easily distinguished (USDA, 1988).

Larvae: First-instar larvae are grayish-white with a black head capsule and are about 1.5 mm long (CABI, 2007). The head capsule of later instars becomes lighter in color, changing to brown. Last instar larvae (Fig. 2) are 20-26 mm long, taper slightly toward each end, and are dirty- white, with five longitudinal purple to brown stripes running down the dorsal surface of the body (Hill, 1983).

Pupae: Pupae are reddish-brown, 11-13 mm Figure 2. Chilo suppressalis larva. long, 2.5 mm wide (Hill, 1983) and have two Image courtesy of Probodelt, SL. ribbed crests on the pronotal margins and two short horns on the head. The cremaster (the terminal spine of the abdomen) bears several small spines (Hattori and Siwi, 1986).

21 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

Adults: In general, C. suppressalis forewings are 11-15 mm long with a wingspan of 20-30 mm (Hill, 1983) with a color varying from dirty-white to yellow-brown (Fig. 3), sprinkled with gray-brown scales. The hindwings are white to yellowish-brown (Hattori and Siwi, 1986).

Adult male: Forewing straw colored, variably or often uniformly suffused with light brown, with brown to dark- brown specks scattered irregularly, sometimes forming small patches; sometimes with broken, diffuse, oblique brown median band between middle of wing and apex; outer margin with row of small dark spots; fringe not metallic, of a lighter shade distally. Hindwing whitish, faintly shaded with brown; fringe uniformly whitish. Front of head conical, strongly protruding forward beyond eye, with very distinct corneous point and a protruding ridge along lower margin; front between point and ventral margin appears concave in profile. These features made Figure 3. Chilo visible by brushing away scales from face between the suppressalis adult. eyes. Labial palp 3 times as long as diameter of eye. Photos courtesy of Wing expanse ranges from 20-30 mm. Male genitalia International Rice bifurcate, juxta symmetrical, the arms bowed, equally Research Institute long, distinctively swollen, and without subapical teeth; Archive aedeagus with long, thin, ventral arm (USDA, 1988). www.bugwood.org.

Adult female: Larger than male, with paler forewing and fewer dark flecks. Hindwing nearly white. Ridge along lower margin of front of head only about half as prominent as that of male, sometimes barely apparent, but still useful for recognizing this species, as so many others lack it entirely. Labial palp 3.5 times as long as diameter of eye. Wing expanse ranges from 24-30 mm. Female genitalia have heavily sclerotized ostial pouch, slightly demarcated from ductus bursae; the latter distinctly swollen posterior to ostial pouch, with heavily sclerotized band; Signum distinct, elongate, with median ridge (USDA, 1988).

Biology and Ecology C. suppressalis is mainly a pest of rice and most of its phenology reflects observations taken on rice.

C. suppressalis is adapted to temperate climatic conditions; larvae survive low winter temperatures in Japan, China, and other northern areas. This is in marked contrast to most other species of Chilo, which are restricted to tropical or sub-tropical regions (CABI, 2007). In favorable (tropical) conditions, up to six generations develop in a year, often overlapping where rice cropping is continuous. In colder climates, final instars remain dormant during the winter. It seems that photoperiod is more important than temperature for diapause; a facultative diapause has been observed when the photoperiod drops below 14 hours (Cho et al., 2005).

22 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

Adults are nocturnal and survive up to a week under field conditions, with females generally living longer than males. Although there is minor temporal isolation in the development of C. suppressalis developing on rice and other hosts, significant overlap and mating exists among these groups (Ueno et al., 2006). Eggs are laid in batches of 60-70, mainly on the basal halves of leaves and occasionally on leaf sheaths (the part of the blade that wraps around the stem). The optimum temperature for hatching is Figure 4. Dead heart symptom. Photo 21-33°C (71-91°F). courtesy of the Purdue Extension Entomology

(J. Obermeyer). After hatching, larvae cluster

between the leaf sheaths and stems, then burrow into the stems to feed. Some early larval instars may move to other plants by wind-aided dispersal. Several larvae may feed together within a single internode, living in a moist pulp of chewed plant debris and frass. They pupate within stems, having first prepared an exit hole from which the adult will emerge. In the tropics and on rice, normal development times are: egg (5 to 6 days), larva (30 days), and pupa (6 days). The life cycle is completed in 35-60 days.

Degree-day models have been calculated for C. suppressalis. At 25°C (77°F) with a 14L:10D photoperiod, the average degree days for eggs, larvae, and male and female pupae were 124, 521, 111, and 103 day-degrees, respectively (Tsumuki et al., 1994).

Symptoms/Signs Infested leaf sheaths first show transparent patches, later turning yellow brown and drying. Stems weaken and easily break as a result of larval feeding inside the stem around the nodes. Seedlings attacked at the base show "dead hearts" (Fig. 4) or drying of the central shoot produced when stem borer larvae kill the growing points of young shoots. Infested plants bear "whiteheads” (empty panicles or with a few filled grains). There are other possible causes of these symptoms (other , fungi, and etc.) and samples of stems should be dissected to establish that C. suppressalis is responsible for the damage.

Pest Importance C. suppressalis is considered one of the most serious pests of rice in the Far East (Grist and Lever, 1969). For example, up to 100% loss have been reported from individual fields in Japan, but normally 4-7% yearly losses have been attributed to this pest overall in Asia (Cho et al., 2005).

23 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

Historically, organophosphates have been used to control this pest in rice and other cropping systems. Due to increasing regulation and resistance in certain populations, pyrethroids are now being studied for their effectiveness (He et al., 2007).

A pheromone dispenser (Selibate CSTM) has been developed for both the monitoring and control of C. suppressalis in Spain. Studies to control the cost of using pheromone disruption by streamlining the density and placement within and around rice fields are in their fourth year (Alfaro et al., 2009). These studies have shown that treatments provide effective control even with reduced pheromone dispenser densities.

Transgenic lines of rice expressing toxin genes from Bacillus thuringensis have been developed with great success against stem borers (ex., cryIA coding sequences, Cheng et al., 1998). Corn has been similarly transformed with equivalent coding sequences; although there are no published reports quantifying control of C. suppressalis with these transgenic lines of , the protection these genes provide rice yields would also have a high likelihood of being successful in corn.

Known Hosts Major hosts Oryza sativa (rice), spp. (grasses), bicolor (sorghum), and Zea mays (corn).

Wild/Minor hosts Andropogon sorghum (broomcorn), Brachiaria mutica (para grass), Brassica oleracea (broccoli), Chaetochloa verticillata (bristlegrass), Coix lacryma-jobi (job's tears), Cymbopogon citratus (lemongrass), Echinochloa spp., Eleusine spp., Oryza spp. (rice), miliaceum (millet), (sour grass), Paspalum distichum (knotgrass), Pennisetum americanum (), Pennisetum glaucum (pearl millet), (common reed), Phragmites communis (common reed), Saccharum fuscum, Saccharum officinarum (sugarcane), Sclerostachya fusca, Solanum lycopersicon (), Setaria verticillata (bristly foxtail), Solanum melongena (), (cattail), Triticum spp. (wheat), Zizania aquatica (annual wildrice), and .

Known Vectors (or associated organisms) Chilo suppressalis is not a known vector and does not have any associated organisms.

Known Distribution Asia: Bangladesh, Brunei, Burma, Cambodia, China, India, , Iran, Japan, Korea, Laos, , Nepal, Pakistan, Philippines, , Thailand, and Vietnam. Europe: France, Portugal, Russian Federation, and Spain. North America: Mexico and United States (). Oceania: and Papua New Guinea.

24 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

Potential Distribution within the United States Chilo suppressalis is present in Hawaii and was first found in 1927. Rice production was already starting to decline in Hawaii prior to this discovery. This discovery, however, is thought to have hastened the decline of the rice industry in Hawaii (USDA, 1957). Yasumatsu et al. (1968) stated that the Asiatic rice borer became extinct in Hawaii sometime between 1939 and 1962. On October 31, 1968, Chilo suppressalis was identified on rice at Wailalua, Kauai. It was not known whether the infestation resulted from progeny of borers from those discovered in 1929 or from the progeny of a new borer introduction (addendum to Yasumatsu et al. (1968).

A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that areas of Texas and have the highest risk from C. suppressalis within the continental United States. Most other states have a low to moderate risk for C. suppressalis establishment based on host availability and climate within the continental United States.

Survey CAPS-Approved Method: Trap with lure. The large plastic delta trap is the approved trap for Chilo suppressalis. The lure information is provided below:

Lure Dispenser Dispenser Lure Length of Compound Load Type Abbreviation Effectiveness a) Z,11-16:AL a) 0.224 Rubber septa CS 4 weeks b) Z,13-18:AL b) 0.0305 c) Z,9-16:AL c) 0.0254 d) BHT d) 0.028

IMPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters.

Notes: The BHT (butylated hydroxytoluene) in the lure dispenser is an added as an anti-oxidant only.

Literature-Based Methods: Trapping: Male moths may be monitored with the female sex pheromone traps. The sex pheromone of female C. suppressalis was initially identified in 1975 as a binary mixture of (Z)-11-hexadecenal (Z11-16:AL) and (Z)-13-octodecenal (Z13-18:AL). The pheromone was present in the female ovipositor extract (Nesbitt et al., 1975; Ohta et al., 1975, 1976). In 1983, another active component of the sex pheromone from C. suppressalis, (Z)- 9-hexadecenal (Z9-16:AL), was discovered (Tatsuki et al, 1983). The three-component blend with aldehyde stabilizing agents has been the primary compound for commercial development of controlled release formulations.

25 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

Goh et al. (1983) compared trap catches when using synthetic [4.5:1 mixture of (Z)-11-hexadecenal and (Z)-13-octodecenal] to those using virgin females. Circular water traps located 100 cm above the ground level were used in paddy fields (three pheromone traps per 30 acres or three pheromone traps plus one virgin female trap per 20 acres). In this study, virgin female traps attracted a higher number of males than synthetic pheromone traps, but according the authors the use of synthetic pheromones was warranted for monitoring C. suppressalis and for mating disruption (Goh et al., 1983).

Ishida et al. (2000) used sticky traps (24 x 25 cm) baited with a 0.6 mg synthetic sex pheromone mixture of the three component sex pheromone [(Z)-11-hexadecenal, (Z)- 13-octodecenal, and (Z)-9-hexadecenal] in a 48:6:5 ratio and 0.06 mg butylated hydroxytoluene soaked into a rubber septa. Each septa was changed monthly and the traps were placed 0.8 meters above the ground with four traps per location. Trapped moths were detached with xylene. Mochida et al. (1984) showed that the three component pheromone trapped males at a much greater rate than traps with virgin females. Kanno et al. (1985) compared traps baited with the three component sex pheromone to light traps. The pheromone trap catch was very high in both the first and second flight seasons. The number of moths caught in the pheromone traps was about three to five times greater than that of light traps.

Visual survey: Visually look or of “deadhearts” in younger plants and "whiteheads" and broken stems on older plants. Examine stalks for borer entry and exit holes. Cut open suspect stems and look for larvae or pupae near the middle of the basal internode. In corn, feeding damage in whorl stage corn should be examined for the presence of larvae. Proper identification of these larvae is important to discriminate between stem borer species.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of C. suppressalis is by morphological identification. Examination of male and female genitalia is needed to distinguish between other Chilo species. Use Passoa (2009) as a key reference to confirm C. suppressalis.

Literature-Based Methods: Hattori and Siwi (1986) published an account of the morphology together with field keys for the identification of adults, larvae, and pupae. One key diagnostic of larvae are the five longitudinal stripes, which can be purple to brown in color. The head of larvae also appear to have two short horns. A key to commonly intercepted larvae of Pyraloidea, which includes Chilo suppressalis, is available (Solis, 2006).

Chilo suppressalis resembles many other species of Chilo in external adult and larval characters, but can be distinguished by examination of the male and female genitalia. Bleszynski (1970) is the best reference for genitalic identification, because his work gives illustrations of the genitalia of both sexes for all known species of Chilo. Peng

26 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

(1971) compares morphological characteristics of C. suppressalis with those of the paddy borer (Tyrporyza incertulas), another important stem borer in .

The most useful external adult feature for recognition of C. suppressalis males is the profile of the head, or face, showing the conical front and the ridge along the lower edge of the front between the eyes. Although a conical front is common in this group of moths, few other species have the ridge on the lower margin, and the only other Old World Chilo spp. known to have such a ridge is C. phragmitellus, which is a stem borer in Phragmites australis and Glyceria aquatica across Eurasia from western Europe to China and Japan. Although the frontal ridge is obvious and works well for identification of males of C. suppressalis, it is less developed in females and may sometimes leave one guessing as to whether it is actually present (USDA, 1988).

Easily Confused Pests Larva of the indigenous North American species, Chilo plejadellus (rice stalk borer), appears almost indistinguishable, insofar as can be determined from specimens preserved in alcohol. C. plejadellus is a widespread species of the eastern United States from the Gulf Coast to Canada and is a stem borer in various coarse grasses, including rice in the south. The pupa of C. plejadellus, however, differs from C. suppressalis in that it lacks a separate pair of small points or ridges on the dorsal side of the caudal segment immediately anterior to the cremaster that always seem to be present in C. suppressalis. The adult is about the same size, shape, and color as C. suppressalis, but it has a scattering of gold metallic scales on the forewing, including the fringe, lacks the ridge along the lower margin of the front between the eyes, and differs in the genitalia of both sexes. The male genitalia of C. plejadellus have the arms of the juxta similarly bowed but less swollen, slightly unequal in length, and each with a short, sharp, subapical tooth (USDA, 1988).

Eoreuma loftini (Mexican rice borer) is a somewhat similar corn and sugarcane borer of Mexico and adjacent states of the United States. The adult is smaller than that of C. suppressalis, often grayer, and lacks the frontal ridge; the larva has one rather than two subventral setae on the meso- and metathorax; and the pupa has numerous enlarged, sharp, thorn-like setae on the abdominal segments (USDA, 1988).

In North American corn, there are other stem borers that tend to be polyphagous and could potentially be confused with C. suppressalis. Papaipema nebris (Lepidoptera: Noctuidae) can be found, and this larva has 5 white stripes that gradually fade during maturity. Fall armyworm may also be found in the whorl of corn plants, and has 3 yellow longitudinal lines on larval bodies. crambidoides, sometimes found in the whorls before entering the rootstock, is yellowish in color with spots (no longitudinal stripes). The European corn borer (Ostrinia nubilalis) larva is without longitudinal stripes. immanis, the hop vine borer can also be found in corn stalks, but also does not have longitudinal stripes.

27 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

References Alfaro, C., Navarro-Llopis, V., and Primo, J. 2009. Optimization of pheromone dispenser density for managing the rice striped stem borer, Chilo suppressalis (Walker), by mating disruption. Crop protection 28(7): 567-572.

Bleszynski, S. 1970. A revision of the world species of Chilo Zincken (Lepidoptera: ). Bulletin of the British Museum (Natural History) Entomology 25: 101-195.

CABI. 2007. Crop Protection Compendium. Commonwealth Agricultural Bureau. http://www.cabicompendium.org.

Cheng, X., Sardana, R., Kaplan, H., and Altosaar, I. 1998. Agrobacterium-transformed rice plants expressing synthetic cryIA(b) and cryIA(c) genes are highly toxic to striped stem borer and yellow stem borer. Proceedings of the National Academy of Sciences 95: 2767-2772.

Cho, J.R., Lee, J.S., Kim, J.J., Lee, M., Kim, H.S., and Boo, K.S. 2005. Cold hardiness of diapausing rice stem borer, Chilo suppressalis Walker (Lepidoptera: Pyralidae). Journal of Asia-Pacific Entomology 8(2): 161-166.

Goh, H.G., Lee, J.O., and Kim, Y.H. 1983. Mass trapping of the striped rice borer (Chilo suppressalis W.) by sex pheromone trap. Res. Rep.Off. Rural Dev. Sci. Fert. Crop Prot. Mycology Farm Prod. Util. 25(10): 136-139.

Grist, D.H. and Lever, R.J.A.W. 1969. Pests of Rice. London, UK: Longman

Hattori, I. and Siwi, S.S. 1986. Rice stemborers in Indonesia. JARQ (Japan Agricultural Research Quarterly) 20(1): 25-30

He, Y.P., Chen, W.M., Shen, J.L., Gao, C.F., Huang, L.Q., Zhou, W.Z., Liu, X.G., and Zhu, Y.C. 2007. Differential susceptibilities to pyrethroids in field populations of Chilo suppressalis (Lepidoptera: Pyralidae). Pesticide Biochemistry and Physiology 89: 12-19.

Hill, D. 1983. Agricultural Insect Pests of the Tropics and their Control. Cambridge University Press, pg. 316.

Ishida, S., Kikui, H., and Tsuchida, K. 2000. Seasonal prevalence of the rice stem borer (Chilo suppressalis (Lepidoptera: Pyralidae) feeding on water-oats (Zizania latifolia) and the influence of its two egg parasites. Res. Bull. Fac. Agr. Gifu. Univ. 65: 21-27.

Kanno, H., Abe, N., Mizusawa, M., Saeki, Y., Koike, K., Kobayashi, S., Tatsuki, S., and Usui, K. 1985. Comparison of trap efficiency and the fluctuation pattern of moth catches between the synthetic sex pheromone and the light-trap in the rice stem borer moth, Chilo suppressalis Walker (Lepidoptera: Pyralidae). Jpn. J. Appl. Ent. Zool. 29: 137-139. (English Abstr.)

Mochida, O., Arida, G.S., Tatsuki, S., and Fukami, J. 1984. A field test on a third component of the female sex pheromone of the rice striped stem borer, Chilo suppressalis, in the Philippines. Entomol. Exp. Appl. 36: 295-296.

Nesbitt, B.F., Beevor, P.S., Hall, D.R., Lester, R., and Dyck, V.A. 1975. Identification of female sex pheromones of moth, Chilo suppressalis. J. Insect Physiol. 21: 1883-1886.

Ohta, K., Tatsuki, S., Uchiumi, K., Kurihara, M., and Fukami, J. 1975. Sex-pheromone of rice stem borer – purification and chemical properties. Agric. Biol. Chem. 39: 2437-2438.

28 Chilo suppressalis Primary Pest of Corn Arthropods Asiatic rice borer Moth

Ohta, K., Tatsuki, S., Uchiumi, K., Kurihara, M., Fukami, J. 1976. Structures of sex-pheromones of rice stem borer. Agric. Biol. Chem. 40: 1897-1899.

Passoa, S. 2009. Screening Key for CAPS Target Pyraloidea in the Eastern and Midwestern United States (males). Lab Manual for the Lepidoptera Identification Workshop. University of . http://caps.ceris.purdue.edu/webfm_send/95

Peng, W. 1971. Morphological studies on rice stem borer (Chilo suppressalis (Walker) and paddy borer (Tryporyza incertulas (Walker). Taipei Nat. Taiwan Univ. Coll. Agr. Mem. 12(1): 150-172.

Solis, M.A. 2006. Key to selected Pyraloidea (Lepidoptera) larvae intercepted at U.S. ports of entry: revision of Pyraloidea in “Keys to some frequently intercepted larvae” by Weisman 1986. http://www.sel.barc.usda.gov/lep/selected_pyraloid_larval_key.pdf

Tatsuki, S., Kurihara, M., Usui, K., Ohguchi, Y., Uchiumi, K., Fukami, J., Arai, K., Yabuki, S., and Tanaka, F. 1983. Sex-pheromone of the rice stem borer Chilo suppressalis (Walker) (Lepidoptera: Pyralidae), the 3rd component Z-9-hex-adecenal. Appl. Entomol. Zool. 18: 443-446.

Tsumuki, H., Take, T., Kanehisa, K., Saito, T., and Ccu, Y.I. 1994. Effect of temperature and voltinism of the rice stem borer, Chilo suppressalis (Lepidoptera: Pyralidae) in Taiwan. European Journal of Entomology 91: 477-479.

Ueno, H., Furukawa, S., and Tsuchida, K. 2006. Difference in the time of mating activity between host- associated populations of the rice stem borer, Chilo suppressalis (Walker). Entomological Science 9: 255-259.

USDA. 1957. Insects not known to occur in the United States. Asiatic rice borer (Chilo suppressalis (Walker)). U.S. Department of Agriculture, Coop. Econ. Insect Rep. 7(44): 855-856.

USDA. 1988. Insects not known to occur in the United States. No. 97. Asiatic rice borer.

Yasumatsu, K., Nishida, T., and Bess, H.A. 1968. On the extinction of the Asiatic rice borer Chilo suppressalis in Hawaii. Proceedings, Hawaiian Entomological Society XX(1): 239- 245.

29 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit Beetle

Diabrotica speciosa

Scientific name Diabrotica speciosa Germar

Synonyms: Diabrotica amabilis, Diabrotica hexaspilota, Diabrotica simoni, Diabrotica simulans, Diabrotica vigens, and Galeruca speciosa

Common names Cucurbit beetle, chrysanthemum beetle, San Antonio beetle, and South American corn rootworm

Type of pest Beetle

Taxonomic position Class: Insecta, Order: Coleoptera, Family: Chrysomelidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List - 2010

Pest Description Diabrotica speciosa was first described by Germar in 1824, as Galeruca speciosa. Two subspecies have been described, D. speciosa vigens (Bolivia, Peru and Ecuador), and D. speciosa amabilis (Bolivia, Colombia, Venezuela and Panama). These two subspecies differ mainly in the coloring of the head and elytra (Araujo Marques, 1941; Bechyne and Bechyne, 1962).

Eggs: Eggs are ovoid, about 0.74 x 0.36 mm, clear white to pale yellow. They exhibit fine reticulation that under the microscope appears like a pattern of polygonal ridges that enclose a variable number of pits (12 to 30) (Krysan, 1986). Eggs are laid in the soil near the base of a host plant in clusters, lightly agglutinated by a colorless secretion. The mandibles and anal plate of the developing larvae can be seen in mature eggs.

Larvae: Defago (1991) published a detailed description of the third instar of D. speciosa. First instars are about 1.2 mm long, and mature third instars are about 8.5 mm long. They are subcylindrical; chalky white; head capsule dirty yellow to light brown, epicraneal and frontal sutures lighter, with long light-brown setae; mandibles reddish dark brown; antennae and palpi pale yellow. Body covered by sparse, short, dark setae; light brown irregular prothoracic plate; dark brown anal plate on the ninth segment, with a pair of small urogomphi. A pygopod is formed by the tenth segment, which serves as a locomotion and adherence organ.

30 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

Pupae: Pupae are 5.8 to 7.1 mm long and white. Females with a pair of tubercles near the apex. Mature third instars build an 8 x 4 mm oval cell in the soil in which they pupate, and tenerals remain for about 3 days.

Adults: Full descriptions of D. speciosa are given by Baly (1886), Araujo Marques (1941), and Christensen (1943). Adults are 5.5 to 7.3 mm long; antennae 4 to 5 mm (Fig. 1). General color grass-green (USDA, 1957); antennae filiform and dark (reddish-brown to black) and nearly equal to the body in length, first three basal segments lighter; head ranging from reddish brown to black; labrum, scutellum, metathorax, tibiae and tarsi black; elytra each with three large oval transverse spots, basal spots larger Figure 1. Adult Diabrotica speciosa. Photo and usually reddish toward the courtesy of Hernan Tolosa. humeral callus, the rest yellow. Ventrally, head and metathorax dark brown, prothorax green, mesothorax and abdomen light brown or yellow-green. Pronotum bi-foveate, convex, smooth, shiny, ¼ wider than long. Male antennae proportionally longer than female antennae. Males with an extra sclerite on the apex of the abdomen that makes it look blunt, compared with the rather pointed female apex.

Biology and Ecology Eggs are laid on the soil near a larval host plant. An approximately 92% success rate at 27°C is takes place after about 8 days. Diabrotica speciosa undergoes three larval instars, which are easily differentiated by the size of the head capsule (see larval description above). In laboratory tests, maize was included in the grouping of most suitable hosts (along with wheat and peanuts), in terms of survival from egg to adult (Cabrera Walsh, 2003). First instars are normally scattered throughout the host's system, but mature larvae tend to congregate in the upper 10 cm of the root under the crown. The larval stage lasts 23 to 25 days (~12 days in laboratory conditions at 25°C), including an inactive prepupal period of 2 to 3 days. At 25°C, the pupal stage lasts 6 days, and is followed by a period of 3 to 5 days during which the recently molted adults remain in the pupal cell, presumably for the cuticle to tan (USDA, 1957).

Young have a yellowish or pale brown color, which turns green with bright yellow spots in 3 days if fresh food is provided. Under laboratory conditions, mating has been observed between 4 and 6 days after emergence, and some females were observed mating again at day 35. Each female laid an average of 1164 eggs during her lifetime, starting on day 8 and extending for a maximum of 77 days. Peak oviposition was observed on days 16 through 56. In a laboratory environment, oviposition on

31 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

maize was preferred over pumpkin, potato and bean seedlings, and maize was as attractive as peanuts in choice tests (Cabrera Walsh, 2003). The number of overlapping generations is conditioned by latitude and climate, being continuous in tropical areas. In Buenos Aires, Argentina, observations indicate there are about three generations per year; the number and timing depends on latitude and climate. Overwintering occurs as an adult (USDA, 1957). These adults can be found concealed in the rosette and crown of winter-growing plants, and they are fairly cold-tolerant (EPPO, 2005).

Pest Importance Diabrotica speciosa is considered to be an important pest throughout southern South America (except Chile), but, being highly polyphagous, qualitative reports of its impact on different crops vary in different regions. It is considered an important pest of maize, cucurbits, and orchard crops throughout its distribution (CABI, 2007). Although it migrates as an adult, no information on observed distances has been found. Redistributing soil via farm machinery that is contaminated with eggs and-or pupae is also a concern.

Adults of this chrysomelid feed on foliage, pollen, flowers and fruits of many plants. The larvae are pests of , especially maize. It is the most harmful species of Diabrotica in Argentina, mainly affecting peanuts in the center of the country. It causes considerable damage to watermelon, squash and tomatoes in Brazil, and potatoes and wheat in southeast Brazil. Young squash plantings and immature tomato fruits are severely damaged in Brazil. Populations are so heavy in some years in Paraguay that vegetable crops are almost completely destroyed. Severe injury also occurs on flowers of various ornamentals such as and (USDA, 1957). Economic thresholds of two insects per plant for vulgaris were determined by Pereira et al. (1997).

IPM programs to combat D. speciosa in South America recommend no-till cultural practices, insecticides when reaching economically damaging levels and a rotation of maize, wheat, and soybeans. In South America, insecticides (carbamates, organophosphates and, more recently, tefluthrin and chlorethoxyfos) to control larvae and baits (along with broad-spectrum insecticides) to control adults are widely used. These baits are sliced roots of several different wild cucurbits laced with insecticides.

Although there is research into using parasitoids (brachonids and tachinids) and pathogens (Beauveria spp. and Metarhizium anisopliae) to combat this pest, no successful biological control programs have been mentioned.

Symptoms/Signs The larval damage resulting from root feeding can cause host death when the host is small, but the larvae will usually only induce stunted growth in larger host plants, due to a reduction in uptake. In corn, attack on young plants by larvae produces a typical condition known as 'goose neck', in which the plant exhibits stunted growth, reduced vigor, and the first few internodes of the plant grow bent, sometimes to such an extent that the plant actually lies on the ground (Figure 2). In the case of peanuts and

32 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle potatoes, the larvae cause external damage or short bores, similar to those of several other pests such as wireworms and other chrysomelids.

On corn, the most economically important stage is the adult, which feeds on the tassels, preventing pollination and kernel number. Adults also cause defoliation and general feeding damage to leaves, flowers and fruit (EPPO, 2005). Like other Diabrotica spp., they ‘Gooseneck’ growth form of corn. Photo are especially associated with Figure 2. courtesy of The Ohio State University. Cucurbitaceae and are tolerant of cucubitacins and generally feed on pollen-rich plant structures of over 70 plant species. When flowers are scarce, beetles may feed on the tender green parts of other hosts, such as alfalfa, potatoes, corn, bean, soybean, lettuce, and cabbage (EPPO, 2005).

In grape, adult beetles eat young leaf edges during budding, which usually does not seriously damage the host (Roberto et al., 2001). During the blooming period, however, beetles have been observed on flowers eating the style, stigma, and eventually the ovary. Beetle stigma feeding determines flower aborting and, as a consequence, clusters show low numbers of flowers and fruits (Fig. 3). Weedy hosts need to be controlled as beetles can also be observed feeding on and moving into grape from surrounding weeds.

B A

Figure 3. Grape cluster after a severe outbreak of D. speciosa during the bloom period (A) and normal cluster (B). Photos courtesy of Roberto et al. (2001).

33 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

Known Hosts Root-feeding larvae of D. speciosa are polyphagous, but the known host range includes corn, wheat, peanut, soybean, and potato. Cabrera Walsh (2003) found that larvae developed well on corn, peanut, and soybean roots, but not so well on pumpkin, beans, and potato. Oviposition preferences roughly parallel larval suitability, but there was a clear preference for cucurbits as adult food, when available; pigweed, sunflower, and alfalfa are secondary hosts. As an adult, D. speciosa has been reported feeding on more than 70 host species (Christensen, 1943; Heineck-Leon and Salles, 1997).

Major hosts Arachis spp. (peanut), Capsicum spp. (pepper), Cucurbita maxima (winter squash), Cucurbita pepo (ornamental gourd), Glycine max (soybean), Solanum tuberosum (potato), Triticum spp. (wheat), Vitis vinifera (grape), and Zea spp. (corn).

Minor hosts Allium spp. (onion, leek), Alternanthera philoxeroides (alligatorweed), Amaranthus spp. (pigweeds), Apium graveolens (celery), Artemisia spp. (absinthium, tarragon), Asparagus spp. (asparagus), Avena spp. (oats), Baccharis articulata, Beta vulgaris (beet), Brassicaceae (mustards), Bromus catharticus (prairie grass), Carica papaya (papaya), Cayaponia spp., Chenopodium spp., Chrysanthemum spp., Cichorium spp. (chicory, endive), Citrullus vulgaris (watermelon), Citrus spp., Coriandrum sativum (coriander), Coronopus didymus (twin cress), Cucumis spp. (melons, cucumbers, gerkins), Cucurbitaceae (cucurbits), Cucurbitella asperata, Cynara spp. (artichoke), Cynodon dactylon (Bahama crass), Cyphomandra betacea (tree tomato), pinnata (pinnate dahlia), spp., Daucus carota (carrot), Fragaria vesca (wild strawberry), Gossypium spp. (cotton), Helianthus annuus (sunflower), Helianthus tuberosus (Jerusalem artichoke), Hibiscus spp., Ilex paraguayensis (Paraguay tea), Ipomoea spp. (sweet potato, morning glory), Lactuca sativa (lettuce), Lagenaria siceraria (bottle gourd), Lavandula officinalis (English lavender), Lilium maculatum (sukash-yuri), Linum usitatissimum (flax), Lolium perenne (rye grass), Luffa spp. (loofah), Lycopersicon esculentum (tomato), Malus spp. (apple), Malva spp. (mallow), Matricaria chamomilla (chamomile), Medicago sativa (alfalfa), Melilotus albus (yellow sweet clover), Mentha spp. (mint), Morrenia odorata (latex plant), Musa spp. (banana), Nasturtium officinale (watercress), Nicotiana tabacum (tobacco), Ocimum basilicum (basil), Orginanum vulgare (oregano), Oryza sativa (rice), Passiflora coerulea (passion flower), Petroselinum crispum (parsley), Pharbitis purpurea (common morning glory), Phaseolus spp. (beans), Physalis viscose (starhair groundcherry), Pimpinella anisum (anise), Pisum sativum (pea), Prunus spp. (stone fruit), Raphanus sativus (radish), Rosa spp. (), Sechium edule (chayote), Sicyos polycanthus, Solanum spp., Solidago chilensis (goldenrod), Sorghum spp., Spinacia oleracea (spinach), Taraxicum officinale (dandelion), Thea sinensis (tea), Trifolium spp. (clover), Triticum spp., Tropaeolum majus (Nasturtium), and Zingiber officinale (ginger).

Known Vectors (or associated organisms) There is evidence that D. speciosa is a viral vector for comoviruses, southern bean mosaic virus, mimosa mosaic virus, tymoviruses (such as passionfruit yellow mosaic

34 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

virus), carmoviruses, and purple granadilla mosaic virus (Ribeiro et al., 1996; Germain, 2000). Lin et al. (1984) showed that D. speciosa transmitted severe mosaic virus (CPSMV – comovirus) to bean. Ribeiro et al. (1996) showed that eggplant mosaic virus (EMV – tymovirus) was transmitted to tobacco by D. speciosa.

Known Distribution Central America: Costa Rica and Panama. South America: Argentina, Bolivia, Brazil, Columbia, Ecuador, French Guiana, Paraguay, Peru, Uruguay, and Venezuela. There is a record of D. speciosa from Mexico, but according to Krysan (1986), it is almost certainly an error.

Potential Distribution within the United States As of 2004, Diabrotica speciosa has been intercepted over 300 times at ports of entry in the United States, but little is known on its potential distribution within the United States. According to a recent analysis by USDA-APHIS-PPQ-CPHST, the greatest risk for establishment of D. speciosa based solely on the presence of hosts occurs in portions of , Colorado, Illinois, Indiana, Iowa, Kansas, Kentucky, , Maryland, Michigan, Minnesota, Mississippi, Missouri, Nebraska, , North Dakota, Ohio, Oklahoma, Texas, and Wisconsin. The pest occurs from temperate Argentina to tropical Brazil. The polyphagous nature of D. speciosa increases the likelihood of finding hosts and suitable environment if it were introduced into the United States, and is thought to be able to adapt to more temperate climates.

Survey CAPS-Approved Method: Visual.

Literature-Based Methods: Visual survey: Visual detection of adults is easy, as their feeding period spans from dawn until dusk. Detection of larval damage, on the other hand, is more difficult. First instars are very difficult to sample, and even large infestations can go undetected until the damage caused to the host is extensive. Larger larvae can sometimes be observed feeding on the roots of plants immediately after pulling out of the soil, but methodical sampling and counting methods have not been developed, as they have been for the North American pest species (Fisher and Bergman, 1986).

Trapping: Adults D. speciosa appear to be universally attracted to aromatic compounds from squash blossoms, Figure 4. Adult banded cucumber beetle, though the specific compound(s) that Diabrotica balteata. Photo courtesy of John attract the beetles varies from species L. Capinera, University of Florida

35 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

to species. Often, simple blends of two or three compounds are much more potent attractants than any single compound. In addition, female-produced sex attractant pheromones are used for mate location in this . In a preliminary trapping test in Brazil, a number of squash volatiles were screened for potential attraction, and 1,4- dimethoxybenzene showed promise as an attractant for D. speciosa (Ventura et al., 2000). Traps baited with 1,4-dimethoxybenzene, a volatile substance of Cucurbita maxima blossoms captured 29.4 times and 9.4 times more beetles than controls in soybean and common bean fields, respectively (Ventura et al., 2000).

The USDA-CPHST laboratory in Otis, MA has applied for funding to manufacture and test potential lures for D. speciosa, but has yet to begin work toward this goal.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of D. speciosa is by morphological identification. Diabrotica speciosa is almost identical to D. balteata (Fig. 4), which is widely present in the southern United States. Confirmation by a chrysomelid specialist is required. D. speciosa can also be confused with Diabrotica viridula (not present in the United States) and other pestiferous Diabrotica species in South America.

Literature-Based Methods: Diabrotica speciosa somewhat Figure 5. Western corn rootworm, Diabrotica resembles the other main pestiferous virgifera. Courtesy of USDA-ARS. Diabrotica in South America, D. viridula, in coloring, size, biology and host range; but D. viridula has dark brown areas toward the cephalic edge of the elytral spots, and distinct humeral plicae. Also, the larvae of D. viridula lack urogomphi on the anal plate.

Easily Confused Pests Survey and detection based on visual detection of symptoms is quite difficult and many other pests can be easily confused. Symptoms, such as dead heart in wheat, goose neck in maize, or stunted growth in most of the larval Figure 6. Southern corn rootworm, hosts of D. speciosa, could be attributed Diabrotica undecimpunctata. Courtesy of to several other root feeders, such as Clemson University - USDA Cooperative Extension Slide Series, www.bugwood.org.

36 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

wireworms (Conoderus spp.; Elateridae), white grubs, (Phytalus spp., Cyclocephala spp., Diloboderus abderus; Melolonthidae), Pantomorus spp. and (), and several chrysomelids (Caeporis spp., Colaspis spp., Maecolaspis spp., Diphaulaca spp. and Cerotoma arcuata) (Gassen, 1984, 1989).

Other rootworms (western corn rootworm, southern corn rootworm) are easily distinguished from D. speciosa as adults by the markings on elytra (compare Figs. 1, 5 and 6).

References Araujo Marques, M. 1941. Contributio ao estudo dos crisomeledeos do gunero Diabrotica. Bol. Escola Nac. Agron., 2:61-143. (In Portuguese)

Baly, J.S. 1886. The Colombian species of the genus Diabrotica, with descriptions of those hitherto uncharacterized. Part I. Zoological Journal of the Linnean Society, 19:213-229.

Bechyne, J. and Bechyne, B. 1962. Liste der bisher in Rio Grande do Sul gefundenen Galeruciden. Pesquisas (Zool.), 6:1-63.

CABI. 2007. Crop Protection Compendium. Wallingford, UK: CAB International. www.cabicompendium.org/cpc

Cabrera Walsh, G. 2003. Host range and reproductive traits of Diabrotica speciosa (Germar) and Diabrotica viridula (F.) (Coleoptera: Chrysomelidae) two species of South American pest rootworms, with notes on other species of Diabroticina. Environmental Entomology 32(2): 276-285.

Christensen, J.R. 1943. Estudio sobre el g'nero Diabrotica Chev. en la Argentina. Rev. Facultad de Agronomia y Veterinaria, 10:464-516. (In Spanish)

Defago, M.T. 1991. Caracterizacion del tercer estadio larval de Diabrotica speciosa. Rev. Peruana de Ent., 33:102-104. (In Spanish)

EPPO. 2005. Diabrotica speciosa. OEPP/EPPO Bulletin 35: 374-376.

Fisher, J.R. and Bergman, M.K. 1986. In: [Krysan JL, Miller TA, eds.] Methods for the Study of Pest Diabrotica. New York, USA: Springer.

Gassen, D.N. 1984. Insetos associados a cultura do trigo no Brasil. Circular Tecnica, 3. Passo Fundo, RS, Brazil: EMRAPA-CPNT.

Gassen, D.N. 1989. Insetos subterraneos prejudiciais as culturas no sul do Brasil. EMBRAPA-CNPT. Documentos, 13. Passo Fundo, RS, Brazil: EMBRAPA-CNPT.

Germain, J.F. 2000. Diabrotica speciosa (Germar), PRA Area: Europe (01/8504). 1-3. European and Mediterranean Plant Protection Organization.

Heineck-Leonel, M.A. and Salles, L.A.B. 1997. Incidence of parasitoids and pathogens in adults of Diabrotica speciosa (Germ.) (Coleoptera: Chysomelidae) in Pelotas, RS. Anais da Sociedade Entomológica do Brasil, 26(1):81-85.

Krysan, J.L. 1986. Introduction: biology, distribution, and identification of pest Diabrotica. In: [Krysan JL, Miller TA, eds.] Methods for the Study of Pest Diabrotica. New York, USA: Springer.

37 Diabrotica speciosa Primary Pest of Corn Arthropods Cucurbit beetle Beetle

Lin, M.T., Hill, J.H., Kitajima, E.W., and Costa, CL. 1984. Two new serotypes of cowpea severe mosaic virus. Phytopathology 74: 581-585.

Pereira, M.F.A., Delfini, L.G., Antoniacomi, M.R., and Calafiori, M.H. 1997. Damage caused by the , Diabrotica speciosa (Germar, 1824), on beans (Phaseolus vulgaris L.), with integrated management. Ecossistema, 22:17-20.

Ribeiro, S.G., Kitajima, E.W., Oliveira, C.R.B., and Koenig, R. 1996. A strain of eggplant mosaic virus isolated from naturally infected tobacco plants in Brazil. Plant Dis. 80: 446-449.

Roberto, S.R., Genta, W., and Ventura, M.U. 2001. Diabrotica speciosa (Ger.) (Coleoptera: Chrysomelida): New Pest in Table Grape Orchards. Neotropical Entomology 30(4): 721-722.

USDA, 1957. Cooperative Economic Insect Report, 7(2):5-6.

Ventura, M.U., Martins, M.C., and Pasini, A. 2000. Response of Diabrotica speciosa and Cerotoma acuata tingomariana (Coleoptera: Chrysomelidae) to volatile attractants. Florida Entomologist 83(4): 403- 410.

38 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

Helicoverpa armigera

Scientific Name Helicoverpa armigera Hübner

Synonyms: Bombyx obsoleta, Chloridea armigera, Chloridea obsoleta, Helicoverpa commoni, Helicoverpa obsoleta, Heliothis armigera, Heliothis conferta, Heliothis fusca, Heliothis obsoleta, Heliothis pulverosa, Heliothis rama, Heliothis uniformis, Noctua armigera, and Noctua barbara

Common Name(s) Old world bollworm, scarce bordered straw worm, corn earworm, cotton bollworm, African cotton bollworm, tobacco budworm, tomato grub, tomato worm, and gram pod borer.

Type of Pest Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Family: Noctuidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description For more information, see Common (1953), Dominguez Garcia-Tejero (1957), Kirkpatrick (1961), Hardwick (1965, 1970), Cayrol (1972), Delattre (1973), and King (1994).

Eggs: Yellowish-white when first laid (Fig. 1), later changing to dark brown just before Figure 1. Eggs of Helicoverpa armigera hatching. Eggs are gum drop-shaped and Photo courtesy of BASF Corp. 0.4-0.6 mm in diameter. The top is smooth, otherwise the surface contains approximately 24 longitudinal ribs. The eggs then change to dark or gray black a day before hatching (Bhatt and Patel, 2001; CABI, 2007).

Larvae: Larval color darkens with successive molts for the six instars typically observed for H. armigera. Coloration can vary considerably due to diet content (Fig. 2 A, B). Coloration ranges from bluish green to brownish red (Fowler and Lakin, 2001). Freshly emerged first instar larvae are translucent and yellowish-white in color. The head, prothoracic shield, supra-anal shield and prothoracic legs are dark-brown to black, as

39 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth are also the spiracles and raised base of the setae, which give the larvae a spotted appearance (Fig. 2A, B) due to sclerotized setae, tubercle bases and spiracles (King, 1994; Bhatt and Patel, 2001). Second instar larvae are yellowish green in color with black thoracic legs. Five abdominal prolegs are present on the third to sixth, and tenth abdominal segments. The full grown larvae are brownish or pale green with brown lateral stripes and distinct dorsal stripe; long and ventrally flattened but convex dorsally. Larval size in the final instar ranges from 3.5 to 4.2 cm in length (King, 1994).

Pupae: Dark tan to brown (Fig. 2C), 14 to 22 mm long and 4.5 to 6.5 mm in width. Body is rounded both anteriorly and posteriorly, with two tapering parallel spines at posterior tip. Pupae typically are found in soil.

A B

C D

Figure 2. Life stages of Helicoverpa armigera (images not to scale): (A, B) larva, (C) pupa, and (D) adult. Photos courtesy of Central Science Laboratory, Harpenden Archive, British Crown and Paolo Mazzei www.bugwood.org.

Adults: A stout-bodied moth with typical noctuid appearance, with 3.5-4 cm wing span; body, 14-19 mm long. Color is variable, but male usually greenish-gray and female orange-brown (Fig. 2D). Forewings have a line of seven to eight blackish spots on the margin and a broad, irregular, transverse brown band. Hind wings are pale-straw color with a broad dark-brown border that contains a paler patch; they have yellowish margins and strongly marked veins and a dark, comma-shaped marking in the middle.

40 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

Biology and Ecology H. armigera overwinters in the soil in the pupal stage. Moths emerge in May to June depending on latitude, and lay eggs singly on a variety of host plants on or near floral structures. Plants in flower are preferred to those that are not in flower (Firempong and Zalucki, 1990b). Depending on the quality of the host, H. armigera may also lay eggs on leaf surfaces. Female moths tend to choose pubescent (hairy) surfaces for oviposition rather than smooth leaf surfaces (King, 1994). ‘Tall’ plants also tend to attract heavier oviposition than shorter plants (Firempong and Zaluski, 1990b). The number of larval instars varies from five to seven, with six being most common (Hardwick, 1965). Larvae drop off the host plant and pupate in the soil, then emerge as adults to start the next generation.

Because H. armigera exhibits overlapping generations, it can be difficult to determine the number of completed generations. Typically 2-5 generations are achieved in subtropical and temperate regions and up to 11 generations can occur under optimal conditions, particularly in tropical areas (Tripathi and Singh, 1991; King, 1994; Fowler and Lakin, 2001). Temperature and availability of suitable host plants are the most important factors influencing the seasonality, number of generations, and the size of H. armigera populations (King, 1994).

The duration of the different life stages decreased as temperature increased from 13.3 to 32.5°C (56 to 91°F). A thermal constant of 51 degree days above the threshold of 10.5°C (51°F) was required for the development of eggs. The larval stage required 215.1 degree days and the pupal stage 151.8 degree days above 11.3 and 13.8°C (52 and 57°F) developmental thresholds, respectively (Jallow and Matsumura, 2001). In a laboratory study, 475 degree days above an 11°C (52°F) threshold were needed to complete development from larvae to adult (Twine, 1978).

H. armigera has a facultative pupal diapause, which is induced by short day lengths (11- 14 hours per day) and low temperatures (15-23°C; 59-73°F) experienced as a larva (CABI, 2007). A summer diapause, in which pupae enter a state of arrested development during prolonged hot, dry conditions, has been recorded in the Sudan (Hackett and Gatehouse, 1982) and Burkina Faso (Nibouche, 1998).

Under adverse conditions, moths can migrate long distances (King, 1994; Zhou et al. 2000; Casimero et al., 2001; Shimizu and Fujisaki, 2002; CABI, 2007). Adults can disperse distances of 10 km during “non-migratory flights” and hundreds of kilometers (up to 250 km) when making “migratory flights”, which occur when host quality or availability declines (Saito, 1999; Zhou et al., 2000; Casimero et al., 2001; Fowler and Lakin, 2001).

For further information, see Dominguez Garcia-Tejero (1957), Pearson (1958), Hardwick (1965), Cayrol (1972), Delattre (1973), Hackett and Gatehouse (1982), King (1994), and CABI (2007).

41 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

Symptoms/Signs H. armigera larvae prefer to feed on reproductive parts of hosts (flowers and fruits), but may also feed on foliage. Feeding damage results in holes bored into reproductive structures, and feeding within the plant. It may be necessary to cut open the plant organs to detect the pest. Secondary pathogens (fungi, bacteria) may develop due to the wounding of the plant. Frass may occur alongside the feeding hole from larval feeding within. Figure 3. Larva feeding on corn cob. On corn: Eggs are laid on the silks, larvae Photo courtesy of Antoine Gyonnet, invade the cobs (Fig. 3,) and developing grain Lépidoptères Poitou-Charentes, is consumed. Secondary bacterial and fungal www.bugwood.org. infections are common. Pest Importance Heliothine moths of the genus Helicoverpa are considered to be among the most damaging insect pests in Australian agriculture, costing approximately $225.2 million per year to control (Clearly et al., 2006). Helicoverpa armigera is a major insect pest of both field and horticultural crops in many parts of the world (Fitt, 1989). The pest status of H. armigera is due in part to the broad host range of its larvae, its feeding preference for reproductive stages of plants, its high fecundity, its high mobility, and its ability to enter facultative diapause and thus adapt to different climates (Cleary et al., 2006). These characteristics make H. armigera particularly well adapted to exploit transient habitats, such as man-made ecosystems.

H. armigera has been reported causing serious losses throughout its range, in particular to cotton, tomatoes and corn. For example, on cotton, two to three larvae on a plant can destroy all the bolls within 15 days; on corn, they consume grains; and on tomatoes, they invade fruits, preventing development and causing falling (CABI, 2007). Young larvae (second and third instar) can cause up to 65% loss to cotton yields (Ting, 1986). Worldwide, H. armigera has been reported on over 180 cultivated hosts and wild species in at least 45 plant families (Venette et al., 2003). The larvae feed mainly on the flowers and fruit of high value crops, and thus high economic damage can be caused at low population densities (Cameron, 1989; CABI, 2007). In pigeon pea, an important grain legume in south Asia, east Africa, and Latin America, this single pest causes yield losses of up to 100% in some years and locations, and worldwide losses to pigeon pea of more than $300 million per year (Thomas et al., 1997).

Helicoverpa armigera is capable of long-distance migratory flights (King, 1994; Zhou et al., 2000; Casimero et al., 2001; Shimizu and Fujisaki, 2002; CABI, 2007).

Management of Helicoverpa spp. in the past has relied heavily on the use of insecticides, and this has led to resistance problems in cotton (Fitt, 1994). Resistance to

42 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

pyrethroids amongst H. armigera is a serious problem (McCaffrey et al., 1989; Trowell et al., 1993).

Known Hosts Note: Not all host plants are equally preferred for oviposition but can be utilized in the absence of a preferred host. There have been several studies within the laboratory setting on host preference. Jallow and Zalucki (1996) found that most females ranked corn, sorghum, and tobacco highest, followed by cotton varieties. The least preferred were cowpea and alfalfa. Cotton and corn were more suitable for development and reproduction of the cotton bollworm than peanut (Hou and Sheng, 2000). Pigeon pea and corn are considered to be the most suitable host for this insect, when compared to sorghum, red ambadi (Hibiscus subdariffa), marigold, and artificial diet (Bantewad and Sarode, 2000). Tobacco, corn, and sunflower were categorized as the most preferred hosts; soybean, cotton, and alfalfa were categorized as intermediate hosts; and cabbage, pigweed, and linseed were the least preferred in an additional study (Firempong and Zalucki, 1990a).

Major hosts Abelmoschus esculentus (okra), Allium spp. (onions, garlic, leek, etc.), Arachis hypogaea (peanut), Avena sativa (oats), Brassicaeae (cruciferous crops), Cajanus cajan (pigeon pea), Capsicum annuum (bell pepper), Carthamus tinctorius (), Cicer arietinum (chickpea, gram), Citrus spp., Cucurbitaceae (cucurbits), caryophyllus (carnation), (finger millet), Glycine max (soybean), Gossypium spp. (cotton), Helianthus annuus (common sunflower), Hordeum vulgare (barley), Lablab purpureus (hyacinth bean), Linum usitatissimum (flax), Malus spp. (apple), Mangifera indica (mango), Medicago sativa (alfalfa), Nicotiana tabacum (tobacco), Pelargonium spp. (geranium), Pennisetum glaucum (pearl millet), Phaseolus spp. (beans), Phaseolus vulgaris (common bean), Pinus spp. (pines), Pisum sativum (pea), Prunus spp. (stone fruit), Solanum esculentum (tomato), Solanum melongena (eggplant), Solanum tuberosum (potato), Sorghum bicolor (sorghum), Triticum spp. (wheat), Triticum aestivum (wheat), Vigna unguiculata (cowpea), and Zea mays (corn) (CABI, 2007).

Poor hosts Vitis vinifera (grape) (Voros, 1996).

Wild hosts Acalypha spp. (copperleaf), Amaranthus spp. (pigweed, ), Datura spp., Datura metel (datura), Gomphrena, niger (black henbane), and Sonchus oleraceus (annual sowthislte) (Gu and Walter, 1999; CABI, 2007).

For a complete listing of hosts see Venette et al. (2003).

Known Vectors (or associated organisms) Helicoverpa armigera is not a known vector and does not have any associated organisms.

43 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

Known Distribution Asia: Afghanistan, Armenia, Azerbaijan, Bangladesh, Bhutan, Brunei Darussalam, Cambodia, China, Cocos Islands, Republic of Georgia, India, Indonesia, Iran, Iraq, Israel, Japan, Jordan, Kazakhstan, Korea, Kuwait, Kyrgyzstan, Laos, Lebanon, Malaysia, Myanmar, Nepal, Philippines, Saudi Arabia, Singapore, Sri Lanka, Syria, Tajikistan, Thailand, Turkey, Turkmenistan, United Arab Emirates, Uzbekistan, Vietnam, and Yemen. Europe: Albania, Bulgaria, Cyprus, Finland, France, Germany, Greece, Hungary, Italy, Macedonia, Malta, Portugal, Romania, Russian Federation, Serbia and Montenegro, Slovenia, Spain, Switzerland, and Ukraine. Africa: Algeria, Angola, Benin, Botswana, Burkina Faso, Burundi, Cameroon, Cape Verde, Central African Republic, Chad, Congo, Cote d’Ivoire, Egypt, Eritrea, Ethiopia, Gabon, Gambia, , Guinea, Kenya, Lesotho, Libya, Madagascar, Malawi, Mauritania, Mauritius, Mayotte, Morocco, , Namibia, Niger, Nigeria, Reunion, Rwanda, Saint Helena, Senegal, Seychelles, Sierra Leone, Somalia, , Sudan, Swaziland, Tanzania, Togo, Tunisia, Uganda, Zambia, and . Oceania: American Samoa, Australia, Belau, Federated States of Micronesia, Fiji, Guam, Kiribati, Marshall Islands, New Caledonia, New Zealand, Norfolk Island, Northern Mariana Islands, Papua New Guinea, Samoa, Solomon Islands, Tonga, Tuvalu, and Vanuatu (CABI, 2007).

Potential Distribution within the United States According to Fowler and Lakin (2001), it is probable that H. armigera could establish in every state in the continental United States based on habitat and host suitability and would probably pose the greatest economic threat to the following states: , , Arkansas, California, Georgia, Illinois, Iowa, Kansas, Louisiana, Michigan, Minnesota, Mississippi, Nebraska, New Mexico, North Carolina, Ohio, Pennsylvania, South Carolina, South Dakota, Tennessee, Texas, Virginia, and Wisconsin. A recent risk analysis by USDA-APHIS-PPQ-CPHST, however, indicates that areas of Alabama, Arkansas, Arizona, California, Florida, Georgia, Louisiana, Mississippi, Missouri, North Carolina, Oklahoma, South Carolina, Tennessee, and Texas have the greatest risk for H. armigera establishment based on host availability and climate within the continental United States. Areas of most states, however, have moderate risk for H. armigera establishment.

Survey CAPS-Approved Method: Trap with lure. Use one of the following traps for H. armigera: 1) Plastic bucket trap [Unitrap], 2) Heliothis trap (plastic mesh cone trap), 3) Texas (Hartstack) trap. For instructions on using the bucket trap, see Appendix G “Plastic Bucket Trap Protocol”. The Texas (Hartstack) trap is not available commercially. See Hartstack et al. (1979) or Johnson and McNeil (n.d) for images and trap design. Trap comparison trials are currently being conducted to evaluate the efficacy of the three traps.

The lure information is provided below:

Lure Dispenser Dispenser Lure Length of Compound Load Type Abbreviation Effectiveness

44 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

a) 'Z,11-16:AL a) 2 mg gray rubber HA 4 weeks b) 'Z,9-16:AL b) 0.08 mg septum c) 'BHT c) 0.208 mg

IMPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters.

Literature-Based Methods: Trapping: (From Venette et al., 2003). Pheromone traps using (Z)-11-hexadecenal and (Z)-9-hexadecenal in a 97:3 ratio have been used to monitor populations of H. armigera (Pawar et al., 1988; Loganathan and Uthamasamy, 1998; Loganathan et al., 1999; Visalakshmi et al., 2000; Zhou et al., 2000). Of three pheromone doses tested in the field (0.75, 1.0, and 1.25 mg/septum), 1 mg attracted the most males (Loganathan and Uthamasamy, 1998); the trap type was not specified. Rubber septa impregnated with these sex pheromone components (1 mg/septum) were equally effective in capturing males for 11 days in the laboratory (Loganathan et al., 1999). Captures of H. armigera in the field were significantly lower with 15-day-old lures than with fresh lures, and the authors recommend replacing lures every 13 days (Loganathan et al., 1999). Similar observations were reported by Pawar et al. (1988). Males responded to the pheromone during dark hours only, commencing at 6:00 PM and terminating at 6:00 AM. The highest response was between 11:00 PM and 4:00 AM (Kant et al., 1999).

Trap design has a significant impact on the number of male H. armigera moths that will be captured with pheromone lures. Funnel traps and Texas traps are substantially more effective than sticky traps (Kant et al., 1999). Hartstack (i.e., hollow cone) traps have also been used to effectively monitor densities of adults (Walker and Cameron, 1990). Cone traps are significantly more effective than water-pan traps (Sheng et al., 2002). Traps have been placed approximately 6 feet (1.8 meter) above the ground (Kant et al., 1999; Zhou et al., 2000), and have been separated by a distance of at least 160 feet (50 meters) (Kant et al., 1999). Aheer et al. (2009), however, installed traps at a height of 4.9 feet (1.5 meters) and were separated by a distance of about 33 feet (10 meters). For routine monitoring of pests, pheromone traps are deployed at a density of 5 traps per hectare (Sidde Gowda et al., 2002).

Adults of both sexes can be captured in black light traps.

Visual survey: Visual inspections of plants for eggs and/or larvae are frequently used to monitor and assess population sizes for H. armigera. Females lay several hundred eggs on the leaves (top 20 cm), flowers and fruits (Duffield and Chapple, 2000). The lower leaf surface is a preferred oviposition site. Eggs may hatch in less than 3 days at an optimum temperature of 27 to 28°C (81 to 82°F). The feeding larvae can be seen on the surface of plants but they are often hidden within plant organs (flowers, fruits, etc.). Bore holes and heaps of frass (excrement) may be visible, but otherwise it is necessary to cut

45 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

open the plant organs, especially damaged fruit, to detect the pest (Bouchard et al., 1992). In temperate regions, H. armigera overwinters as a pupa buried several cm in the soil. Adults appear in April to May and can be observed until October, because of the long migration period.

In vegetative Australian cotton and irrigated soybean, a minimum of 60 whole plants per 100 hectare commercial field are examined for the presence of H. armigera eggs or larvae; when plants begin to produce squares, only the upper terminal (approximately 20 cm) of a plant is inspected (Brown, 1984; Dillon and Fitt, 1995; Duffield and Chapple, 2000). In experimental plots, visual inspections for H. armigera in pigeon pea were restricted to the upper third of whole plants (4 sets of five plants in a 30 x 30 meter plot) (Sigsgaard and Ersbøll, 1999).

Leaves of tomato plants are more attractive than flowers or fruits as H. armigera oviposition sites, but use of a single-leaf sample unit (with a sample size of 30 plants per field) has proven ineffective in detecting low densities of H. armigera (Cameron et al., 2001). On some tomato cultivars, leaves in the upper half of the plant are preferentially selected for oviposition (Saour and Causse, 1993).

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of Helicoverpa armigera is by morphological identification. H. armigera can be visually screened to some degree, but definitive screening and identification requires dissection. Helicoverpa armigera and the native, abundant species, are very similar looking. Final identification is by dissection of (adult) male genitalic structures.

Literature-Based Methods: Adults H. armigera may be identified by distinct differences in genitalia (Common, 1953; Kirkpatrick, 1961; Hardwick, 1965). Differentiation between H. armigera and H. zea is very difficult; identification is by dissection of internal structures of adult males (Pogue, 2004). A morphological study of H. assulta, H. punctigera, and Heliothis virescens (formerly H. rubrescens) compares similarities and differences between species; a key is provided for identifying adults (Kirkpatrick, 1961). Immunological tests are available to differentiate H. punctigera and Heliothis virescens in egg or larval stages (Ng et al., 1998).

The LepTon test, an Linked Immunosorbent Assay (ELISA) based approach, has been developed to distinguish between H. armigera and H. punctigera in the egg and larval stages (Trowell et al., 1993). Cahill et al. (1984) provide morphological information to distinguish third/fourth and sixth instar larvae of H. armigera and H. punctigera.

Agusti et al., (1999) developed sequence amplified characterized region (SCAR) markers to detect H. armigera eggs in the gut of predators. It may be possible to adapt this procedure to detect H. armigera in planta.

46 Helicoverpa armigera Primary Pest of Corn Arthropods Old world bollworm Moth

A screening aid is available for H. armigera at: http://caps.ceris.purdue.edu/webfm_send/552.

Instructions for dissecting H. armigera are available at: http://caps.ceris.purdue.edu/webfm_send/551, and http://caps.ceris.purdue.edu/webfm_send/550.

Easily Confused Pests Several noctuid pests can be confused easily with H. armigera, including H. assulta (not known in the United States), H. punctigera (not known in the United States), H. zea (present in the United States), and Heliothis virescens (present in the United States) (Kirkpatrick, 1961; CABI, 2007).

References Agusti, N., De Vicente, M.C., and Gabarra, R. 1999. Development of sequence amplified characterized region (SCAR) markers of Helicoverpa armigera: a new polymerase chain reaction-based technique for predator gut analysis. Molecular Ecology 8: 1467-1474.

Aheer, G.M., Ali, A., and Akram, M. 2009. Effect of weather factors on populations of Helicoverpa armigera moths at cotton-based agro-ecological sites. Entomological Research 29: 36-42.

Bantewad, S.D., and Sarode, S.V. 2000. Influence of different hosts on the biology of Helicoverpa armigera (Hübner). Shashpa 7(2): 133-136.

Bhatt, N.J., and Patel, R.K. 2001. Biology of chickpea pod borer, Helicoverpa armigera. Indian Journal of Entomology 63(3): 255-259.

Bouchard, D., Oudraogo, A., and Boivins, G. 1992. Vertical distribution, spatial dispersion and sequential sampling plan for fruit damage by Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae) on tomato crop in Burkina Faso. Tropical Pest Management 38(3): 250-253.

Brown, G. 1984. Field experience in cotton pest management in north western New South Wales, pp. 128-134. In P. Bailey and D. Swincer [eds.], Proceedings of the fourth Australian Applied Entomological Research Conference, Adelaide, Australia.

CABI. 2007. Crop Protection Compendium. Commonweath Agricultural Bureau, International. http://www.cabicompendium.org/.

Cahill, M., Easton, C., Forreserm N., and Goodyer, G. 1984. Larval identification of Heliothis punctigera and Heliothis armigera. Aust. Cotton Growers Res. Conv. Toowoomba. Pg. 216-221.

Cameron, P.J. 1989. Helicoverpa armigera (Hübner), a tomato fruitworm (Lepidoptera: Noctuidae). Tech. Commun. Commw. Inst. Biol. Control 10: 87-91.

Cameron, P., Walker, G., Herman, T., and Wallace, A. 2001. Development of economic thresholds and monitoring systems for Helicoverpa armigera (Lepidoptera: Noctuidae) in tomatoes. Journal of Economic Entomology 94: 1104-1112.

Casimero, V., Nakasuji, F., and Fujisaki, K. 2001. The influence of larval and adult food quality on the calling rate and pre-calling period of females of the cotton bollworm, Helicoverpa armigera Hübner (Lepidoptera : Noctuidae). Appl. Entomol. Zool. 36(1): 33-40.

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Cayrol, R.A. 1972. Famille des Noctuidae. Sous-famille des Melicleptriinae. Helicoverpa armigera Hb. In: Balachowsky AS, ed. Entomologie appliquée à l'agriculture, Vol. 2, Paris, France: Masson et Cie, 1431- 1444.

Cleary, A.J., Cribb, B.W., and Murray, D.A.H. 2006. Helicoverpa armigera (Hübner): can wheat stubble protect cotton from attack. Australian Journal of Entomology 45: 10-15.

Common, I.F.B. 1953. The Australian species of Heliothis (Lepidoptera: Noctuidae) and their pest status. Australian Journal of Zoology 1: 319-344.

Delattre, R. 1973. Pests and diseases in cotton growing. Phytosanitary handbook. Parasites et maladies en culture cotonniere. Manuel phytosanitaire. Paris, Institut de Recherches du Coton et des Textiles Exotiques., France.

Dillon, G. and Fitt, G. 1995. Reassessment of sampling relationships for Helicoverpa spp. (Lepidoptera: Noctuidae) in Australian cotton. Bulletin of Entomological Research 85: 321-329.

Dominguez Garcia-Tejero, F. 1957. Bollworm of tomato, Heliothis armigera Hb. (= absoleta F). In: Dossat SA, ed. Plagas y Enfermedades de las Plantas Cultivadas, 403-407. Madrid, Spain.

Duffield, S.J. and Chapple, D.G. 2000. Within-plant distribution of Helicoverpa armigera (Hubner) and Helicoverpa punctigera (Wallengren) (Lepidoptera: Noctuidae) eggs on irrigated soybean. Australian Journal of Entomology 40: 151-157.

Firempong, S. and Zalucki, M. 1990a. Host plant preferences of populations of Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae) from different geographic locations. Australian Journal of Zoology 37: 665-673.

Firempong, S. and Zalucki, M. 1990b. Host plant selection by Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae); role of certain plant attributes. Australian Journal of Zoology 37: 675-683.

Fitt, G.P. 1989. The ecology of Heliothis spp. in relation to agroecosystems. 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.

Fowler, G.A. and Lakin, K.R. 2001. Risk Assessment: The old world bollworm, Helicoverpa armigera (Hübner), (Lepidoptera: Noctuidae). USDA-APHIS-PPQ-CPHST-PERAL.

Gu, H. and Walter, G.H. 1999. Is the common sowthistle (Sonchus oleraceus) a primary host plant of the cotton bollworm, Helicoverpa armigera (Lep. Noctuidae)? Oviposition and larval performance. J. Appl. Ent. 123: 99-105.

Hackett D.S. and Gatehouse A.G. 1982. Diapause in Heliothis armigera (Hubner) and H. fletcheri (Hardwick) (Lepidoptera: Noctuidae) in the Sudan Gezira. Bulletin of Entomological Research 72(3): 409- 422.

Hardwick, D.F. 1965. The corn earworm complex. Memoirs of the Entomological Society of Canada, 40: 1-247.

Hardwick, D.F. 1970. A generic revision of the North American Heliothidinae (Lepidoptera: Noctuidae). Memoirs of the Entomological Society of Canada, 73: 1-59.

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Hartstack, A.W., Witz, J.A, and Buck, D.R. 1979. Moth traps for the tobacco budworm. J. Econom. Entomol. 72: 519-522.

Hou, M. and Sheng, C. 2000. Effects of different foods on growth, development, and reproduction of cotton bollworm, Helicoverpa armigera (Hubner) (Lepidoptera: Noctuidae). Acta Entomologica Sinica 43(2): 168-175. English summary pg. 174-175.

Jallow, M.F.A. and Zalucki, M.P. 1996. Within- and between- population variation in host-plant preference and specificity in Australian Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae). Australian Journal of Zoology 44: 503-519.

Jallow, M.F.A. and Matsuura, M. 2001. Influence of temperature on the rate of development of Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae). Appl. Entomol. Zool. 36: 427-430.

Johnson, D. and McNeil, S. n.d. Plans and Parts List: "Texas" Style Cone Trap for Monitoring Certain Insect Pests. ENTFACT-010.University of Kentucky College of Agriculture.

Kant, K., Kanaujia, K.R., and Kanaujia, S. 1999. Rhythmicity and orientation of Helicoverpa armigera (Hübner) to pheromone and influence of trap design and distance on moth trapping. Journal of Insect Science 12: 6-8.

King, A.B.S. 1994. Heliothis/Helicoverpa (Lepidoptera: Noctuidae) In: Matthews G.A. & J. P. Tunstall (eds), Insect Pests of Cotton. Wallingford, UK: CAB International, Wallingford, 39-106.

Kirkpatrick, T. H. 1961. Comparative morphological studies of Heliothis species (Lepidoptera: Noctuidae) in Queensland. Queensland Journal of Agricultural Science 18: 179-194.

Loganathan, M. and Uthamasamy, S. 1998. Efficacy of a sex pheromone formulation for monitoring Heliothis armigera Hübner moths on cotton. Journal of Entomological Research 22: 35-38.

Loganathan, M., Sasikumar, M., and Uthamasamy, S. 1999. Assessment of duration of pheromone dispersion for monitoring Heliothis armigera (Hüb.) on cotton. Journal of Entomological Research 23: 61- 64.

McCaffery, A.R., King, A.B.S., Walker, A.J., and El-Nayir, H. 1989. Resistance to synthetic pyrethroids in the bollworm, Heliothis armigera from Andhra Pradesh, India. Pesticide Science 27: 65-76.

Ng, S., Cibulsky, R., and Trowell, S. 1998. LepTon HTK - a heliothine diagnostic test kit: an update. In: Dugger P, Richter D, Proceedings of the Beltwide Cotton Conferences, Volume 2, pp. 1040-1043., pp. 1040-1043, Beltwide Cotton Conferences. National Cotton Council, Memphis, USA.

Nibouche, S. 1998. High temperature induced diapause in the cotton bollworm Helicoverpa armigera. Entomologia Experimentalis et Applicata 87: 271-274.

Pawar, C., Sithanantham, S., Bhatnagar, V., Srivastava, C., and Reed, W. 1988. The development of sex pheromone trapping of Heliothis armigera at ICRISAT, India. Tropical Pest Management 34: 39-43.

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Saito, O. 1999. Flight activity changes of the cotton bollworm, Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae), by aging and copulation as measured by flight actograph. Applied Entomology and Zoology 35: 53-61.

Saour, G. and Causse, R. 1993. Oviposition behaviour of Helicoverpa armigera Hübner (Lepidoptera: Noctuidae) on tomato. Journal of Applied Entomology/Zeitschrift für Angewandte Entomologie, 115: 203- 209. (English Abstract)

Sheng, C.F., Su, J.W., Wang, H.T., Fan, W.M., and Xuan, W.J. 2002. An efficiency comparison of cone and water tray traps baited with pheromone for capturing male moths of Helicoverpa armigera. Acta Entomologica Sinica, 45: 271-274. (English abstract).

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51 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

Heteronychus arator

Scientific Name Heteronychus arator (Fabricius)

Synonyms: Heteronychus sanctaehelenae, Heteronychus transvaalensis, and Scarabaeus arator

Common Names African black beetle, black maize beetle, black lawn beetle, and black beetle

Type of Pest Beetle

Taxonomic Position Class: Insecta, Order: Coleoptera, Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description Life stages are shown in Figures 1 and 2.

Eggs: White, oval, and measuring approximately 1.8 mm long at time of oviposition. Eggs grow larger through development, and become more round in shape. Eggs are laid singly at a soil depth of . Illustration of each stage of 1 to 5 cm. Females each lay between 12 to Figure 1 the life cycle of the African black 20 eggs total. In the field, eggs hatch after beetle, showing a close up view of approximately 20 days (Matthiessen and each stage and a background view Learmonth, 2005; CABI, 2007). showing that the eggs, larvae, and

pupae are all underground stages Larvae: There are three larval instars. Larvae with the adult beetles as the only can be seen clearly with the naked eye. stage appearing above ground. Larvae are creamy-white except for the brown Illustration courtesy of NSW head capsule and hind segments, which Agriculture. appear dark where the contents of the gut http://www.ricecrc.org/Hort/ascu/zecl/ show through the body wall. The head zeck113.htm capsule is smooth textured, measuring 1.5 mm, 2.4 mm, and 4.0 mm at each respective instar. The third-instar larva is approximately 25 mm long when fully developed. African black beetle larvae are soil-dwelling and resemble white 'curl grubs.’ They have three

52 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

pairs of legs on the thorax, a prominent brown head with black jaws, and are up to 25 mm long. The abdomen is swollen, baggy, and gray/blue-green due to the food and soil they have eaten. Larvae eat plant roots, potentially causing significant damage to turf, horticultural crops, and ornamentals. Turf is the preferred host of the larvae (Matthiessen and Learmonth, 2005; CABI, 2007).

Figure 2. Eggs, larvae, and adult African black beetle. Photo courtesy of Yates Ltd. http://www.yates.com/au/ProblemSolver/BlackBeetle.asp

Pupae: The larvae, when fully grown, enter a short-lived pupal stage, which measures approximately 15 mm long and is typically coleopteran in form (cylindrical shape), initially pale yellow, but becoming reddish-brown nearer to the time of emergence (Matthiessen and Learmonth, 2005).

Adults: The adult beetle when newly emerged is a rich chestnut color, but soon changes to a characteristic glossy black. Beetles are 12 to 15 mm long; shiny black dorsally and reddish-brown ventrally. The females are slightly larger than males. Males and females are readily differentiated by the shape of the foreleg tarsus. The tarsus of the male is much thicker, shorter, and somewhat hooked compared with that of the female, which is longer and filamentous. A less obvious sexual difference is in the form of the pygidium at the end of the abdomen. In the male, it is broadly rounded, and in the female, it is apically pointed. The adult beetle is the main pest stage (Matthiessen and Learmonth, 2005; CABI, 2007).

53 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

Biology and Ecology H. arator is a polyphagous, univoltine pest of pasturelands, turf, and agricultural crops in Australia, New Zealand, and Africa. These scarab beetles spend their entire lifecycle belowground, with the exception of the adult stage (Matthiessen and Learmonth, 1998) (Fig. 1). In spring, the majority of mating occurs, although some may ensue in fall. During this time, adults crawl on the soil surface at night, and flying is limited. Larvae mature in midsummer. Adults emerge after about two weeks in summer to late autumn. The adults are usually found on or under the soil surface, to a depth of about 150 mm. They are a shiny black and cylindrical cockchafer that is slow moving and is approximately 15 mm long. The adult is capable of flying, which serves to disperse the beetle to new sites (Matthiessen and Learmonth, 2005). This insect is particularly noted for its massed swarms that occur at high population levels (Watson, 1980). The beetles are nocturnal. Wet conditions during the egg and first instar larval stages are fatal, but as the larvae grow, their ability to cope with high moisture levels increases (Matthiessen and Learmonth, 2005).

In Australia, development from egg laying to adult emergence takes about three months. Temperatures above 15°C (59°F) are most favorable for development and survival of H. arator, with optimum larval development occurring at 20-25°C (68-77°F) (King et al., 1981a).

Symptoms/Signs Stems experience external feeding, and the whole plant may be toppled or uprooted. Adult damage to plants typically involves chewing of the cortex of stems just below the surface of the ground.

In corn, the beetles eat into the stems of the growing plants just below the soil surface, causing rapid wilt of the growing center leaves (‘deadheart’) and death of the plant. The damaged area of the stem has a frayed (shredded) appearance, which distinguishes it from the damage caused by cutworms. The fraying is caused by the beetles consuming the soft tissues but leaving the fibrous material. Both young and mature plants may be attacked, but damage usually occurs in the early stages of growth up to 7 weeks after planting. The peak period is 3-5 weeks after planting. If young plants are infested, however, a reduction in plant stand is often observed. Replanting is the only solution for this plant stand reduction. Older plants are weakened and prone to lodging. Beetles may attack the ears of lodged plants. The grubs (larvae) prefer to feed on organic matter in the soil, but may cause some root damage. Corn plants are often stunted and have multiple tillers. Toit et al. (1997) indicated that natural grass habitats may be preferred for oviposition and that adults migrate from natural vegetation to corn for feeding.

In potato, the adult beetles burrow into the tubers and makes holes about 10 mm in diameter with frayed edges. The depth of the holes vary depending on the length of time the beetles feed on the tubers. They can be as large as holes made by millipedes (Venter and Louw, 1978). At times, the larvae can feed in the cavities made by the adult beetles.

54 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

In sugarcane, damage is similar to that in corn but results in ‘dead hearts’ in young cane or attack of the underground buds and stem bases in older cane.

In woody vines (e.g., grape) and eucalyptus, this type of damage occurs most frequently, causes greater growth distortion, and is potentially fatal to newly planted cuttings or seedlings. African black beetles eat the cuttings and rootlings at or just below ground level, ring barking of the vine, and causing wilting and collapse. The chewing is more likely to be sufficiently deep or to extend more fully around the circumference of the thinner stems at early stages of plant growth. The problem is greatest where vines have been planted onto old pasture land, especially if kikuyu (Pennisetum clandestinum) is present. High densities of H. arator in pastures lead to clover (a non- host) becoming dominant over grasses (King et al., 1982; Matthiessen and Learmonth, 2005).

Crops planted in heavy, moist in low-lying areas seem to be at the greatest risk of attack in Africa.

Pest Importance The adult is the main pest stage. The adult is the only aboveground stage and is capable of flight. The beetles are of considerable economic importance because they attack a wide range of plants. The beetle damages pastures, particularly newly-sown ryegrass and perennial grasses, millet, corn, turf, barley, triticale, wheat crops (not oats), a wide range of vegetable crops, grape vines, ornamental plants and newly- planted trees. Larvae damage turf and underground crops, notably potato tubers (Matthiessen and Learmonth, 2005). According to Drinkwater (1979, 1982), H. arator is a soil insect of great importance on maize in South Africa,

Impact on newly planted grapevine and eucalyptus seedlings can be severe in patches within a vineyard or plantation, leading to areas of total loss amongst the plant stand. Heavy damage to perennial pasture can be caused by H. arator build-up in years with a drier than average spring and early summer, causing greater than usual survival of first- instar larvae (King et al., 1981b). These climate-driven outbreaks are characteristic of regions that typically have a wet summer, such as the North Island of New Zealand and eastern Australia. Across the regions infested by African black beetle, this insect can cause significant economic damage to horticultural crops such as young vines (newly planted cuttings and young, rooted vines), olives, and potatoes. In grape, damage primarily occurs in the first two years after planting because after this time the vines become too woody to be damaged by the beetle. However, older vines may still be damaged, especially if they have been stressed. The impact of losing young vines is twofold, including replanting costs (especially if grafted vines are involved), and loss of yield through delayed grape production. The unevenness in vine maturity in the block presents management problems, for example, in terms of weed control and vine training. Partial damage to vines by African black beetle can result in retarded growth and add to the cost of vine training, because of the prolonged time that such vines require individual attention (Matthiessen and Learmonth, 2005; CABI, 2007).

55 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

Known Hosts Major hosts Eucalyptus spp. (Eucalyptus tree), Lolium perenne (perennial ryegrass), pastures, Solanum tuberosum (potato), Vitis vinifera (grape), Saccharum officinarum (sugarcane), and Zea mays (corn) (CABI, 2007).

Minor hosts Ananas comosus (pineapple), Begonia spp. (begonia), Brassica napus (turnip), Brassica oleracea var. capitata (cabbage), Bromus catharticus (prairie grass), Calendula spp. (marigold), Cucurbita spp. (squash), Daucus carota (carrot), Elymus repens (couch grass), Eucalyptus saligna (blue gum), Fragaria x ananassa (strawberry), Lactuca sativa (lettuce), Lycopersicon esculentum (tomato), Olea spp. (olives), Paspalum nicorae (Brunswick grass), Pennisetum clandestinum (kikuyu grass), Petunia spp. (petunia), Phaseolus vulgaris (bean), Phlox spp. (phlox), Pisum sativum (pea), Protea spp. (protea), Rheum rhabarbarum (rhubarb), Secale montanum (perennial rye), Sorghum spp. (sorghum), and Triticum aestivum (wheat) (CABI, 2007).

Known Vectors (or associated insects) H. arator is not a known vector and does not have any associated organisms.

Known Distribution Africa: Angola, Botswana, Comoros, Congo Democratic Republic, Ethiopia, Kenya, Lesotho, Madagascar, Malawi, Mozambique, Namibia, Saint Helena, South Africa, Tanzania, Zaire, Zambia, and Zimbabwe; Oceania: Australia, New Zealand, Norfolk Island, and Papua New Guinea (CABI, 2007). There are a few records for South America and Central America, but they are not considered reliable.

Potential Distribution within the United States The current distribution in Australia and New Zealand of H. arator indicates that many regions in the United States may be climatically suitable for the beetle. It is found throughout coastal mainland Australia (north to Brisbane and south to Melbourne) and found in coastal South and Western Australia. H. arator is also a pest on the north island of New Zealand. Computer projections for Australia indicate a potential distribution from northern Queensland to southern Tasmania. These areas would correspond to plant hardiness zones 7 through 11 in the United States.

A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that areas of the southeastern United States, Louisiana, Mississippi, and Texas, have the greatest risk for H. arator establishment based on host availability and climate within the continental United States. Areas of most states, however, have areas of moderate risk for H. arator establishment. Establishment is unlikely in portions of Colorado, Idaho, Kentucky, Minnesota, Montana, Nebraska, New Mexico, Nevada, Oregon, Virginia, Washington, , Wisconsin, Wyoming, and Utah.

56 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

Survey CAPS-Approved Method: Visual survey is the method to survey for H. arator.

Literature-Based Methods: Visual survey: Areas that are rotated with or replace pasture lands are most at risk of damage from the African black beetle. Most damage by the African black beetle occurs during the spring to early summer when the adults are most active crawling on the soil surface and again after new adults emerge in mid summer to fall. In corn, the beetles eat into the stems of the growing plants just below the soil surface, causing rapid wilt of the growing center leaves and death of the plant. The damaged area of the stem has a frayed appearance.

In grass and turf, heavy infestations can be detected by lifting up tufts of grass and inspecting for abundant frass or distinct channeling of soil with embedded larvae. Less dense infestations will be evident if sections of grass are dug and examined for presence of larvae or adults.

Sampling: Mathiessen and Learmonth (1993) devised a method for sampling H. arator in potato crops where the pest was known to be present. A modified version of their approach may be useful for surveys in other crops. In their survey, 50 cm-long portions of hilled-up rows comprised the sample unit. A 70 x 30 cm piece of sheet steel is pressed into soil across the row with soil on one side of the metal sheet being excavated. The steel sheet is then removed to expose an undisturbed soil face of the potato hill. Presence of the beetle or other pests and associated plant damage in the top, center, or bottom of soil cross sections is recorded. Fifty samples were examined at each sampling time in a uniform grid across a 0.2 ha crop area.

Soil sampling is also used to monitor populations of H. arator in Australia. Adults were counted from shovels full of soil, and six beetles per square meter represented a potentially damaging population (Matthiessen and Learmonth, 1998). To estimate the density of H. arator in pastures 100 soil core samples, 10 cm in diameter x 15 cm deep were used. The soil cores were broken up at the time they were taken and searched only for easily seen large life stages of H. arator that occur in the summer and autumn (Matthiessen and Ridsdill-Smith, 1991). King et al. (1981c) collected cores and extracted the large insect stages by dry sorting of the sample Figure 3. Example of a cores; eggs and small larvae were separated from the pitfall trap. Photo courtesy soil using the flotation and wet-sieving technique of Kain of Mnolf. and Atkinson (1976). http://commons.wikimedia. org/wiki/File:Barber_pitfall_ Trapping: Matthiessen and Learmonth (1998) used pitfall trap.jpg

57 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

(Fig. 3), light, and window traps to monitor H. arator in Australia. Light traps are often used in Australia to monitor adult flight activity during the summer and fall prior to planting on old pasture or potato land. Light traps were similar to a Pennsylvania trap (Southwood, 1978). The light was a vertically-oriented 60 cm-long 20 watt fluorescent black light, the center of which was 1.5 meters above the ground. Four vertically- oriented 17.5-cm wide panels equi-radial from the light served as baffles to arrest the flight of insects attracted to the light, causing them to fall through a 21 cm diameter funnel into a collecting container holding an insecticidal vapor strip. A timer kept the light on daily from sunset to sunrise. Light traps were cleared weekly.

Because the beetles are clumsy walkers, they can be collected by pitfall traps or sharp sided plough lines. Matthiessen and Learmonth (1998) made pitfall traps from a 21 cm diameter funnel fitted at ground level into a buried PVC cylinder. Insects fell into a 21 cm plastic jar containing 500 ml of 1:1 ethylene glycol and water. Mesh panels on the upper sides of the collecting jar allowed rainfall to drain away. These traps were spaced at 10 meter intervals at one location. Subsequent traps were made from a 10 cm diameter plastic funnel glued into the screw-top lid of a 250 ml plastic jar. These smaller traps were more easily placed in pasture by creating a hold with a 10 cm diameter corer, and inserting the whole trap assembly. No preservative was used for these smaller traps due to the small size and absence of large predators capable of consuming adult H. arator. The narrow neck of the funnel fastened into the lid prevented escape of beetles, and holes at the base of the collecting jar allowed drainage. Typically, ten traps were deployed, at 5 meter intervals in each of two lines 10 meters apart. Captures in all pitfall traps were assessed weekly.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of H. arator is by morphological identification. Larvae can be identified with the naked eye. Larvae may be confused with the exotic species Australaphodius frenchi.

Literature-Based Methods: African beetle larvae can be identified with the naked eye, since their anal opening is horizontal, compared with a vertical opening in other species. Smith et al. (1995) provided detailed illustrated descriptions and a laboratory and field key to third star larvae. Cumpston (1940) also described the features of the larvae that allow H. arator to be distinguished from other species. Keys to identify adults from related species are given by Enrodi (1985).

Diagnostic images of H. arator are available at http://www.padil.gov.au/viewPestDiagnosticImages.aspx?id=754.

Easily Confused Pests Larvae can be confused with grass grubs. The overall size of H. arator larvae is much greater than that of the grass grub, reaching about 2.5 cm when fully grown. The head of H. arator is light brown and the body grayish or creamy white except for the hind end. Spiracles (breathing pores) are more prominent in the black beetle than in grass grubs and show clearly as orange spots down the sides of the larvae. The anal cleft in the

58 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

black beetle is distinctly half-moon-shaped but in the grass grub is prominently Y- shaped.

The larvae can be confused with lesser pasture cockchafer (Australaphodius frenchi) in Australia. However, larvae of the lesser pasture beetle are never larger than first instar African black beetle and are much shorter (only up to 3 to 4 mm). To avoid confusion between H. arator and native cockchafers, close examination is necessary (Matthiessen and Learmonth, 2005). The adults of H. arator may be confused with the red headed pasture cockchafer (Adoryphorus coulonii). In dorsal view, the H. arator body shape is almost parallel compared to distinctly oval in A. coulonii. The elytra of H. arator is weakly impressed with longitudinal striae and indistinctly punctuate between striae compared to A. coulonii elytra, which has deeply impressed striae and distinctly punctuate between striae. After reviewing the literature, it appears that Australaphodius frenchi and Adoryphorus coulonii are not currently present in the United States. References CABI. 2007. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Cumpston, D.M. 1940. On the external morphology and biology of Heteronychus sanctae-Helenae Blanch and Metanastes vulgivagus Olliff (Col., Scarabaeidae, Dynastinae). Proceedings of the Linnaean Society of New South Wales 65: 289-300.

Drinkwater, T.W. 1979. The black maize beetle. Fmg. S. Afr. Maize Ser. D. Leaflet D4.

Drinkwater, T.W. 1982. The control of the black maize beetle, Heteronychus arator (Coleoptera: Scarabaeidae) in maize in South Africa. Phytophylactica 14(4): 165-167.

Endrodi, S. 1985. The Dynastinae of the world. Dordrecht, Netherlands: Dr. W. Junk.

Kain, W.M. and Atkinson, D.S. 1976. Population studies of (White). II. A rapid mechanical extraction method suitable for intensive sampling of Costelytra zealandica and other scarabeids. New Zealand Journal of Experimental Agriculture 4: 391-397.

King, P.D., Meekings, J.S., and Mercer, C.F. 1981a. Effects of whitefringed weevil (Graphognathus leucoloma) and black beetle (Heteronychus arator) populations on pasture species. New Zealand Journal of Agricultural Research 25: 405-414.

King, P.D., Mercer, C.F., and Meekings, J.S. 1981b. Ecology of black beetle, Heteronychus arator (Coleoptera Scarabaeidae)- population modeling. New Zealand Journal of Agricultural Research 24(1): 99-105.

King, P.D., Mercer, C.F., and Meekings, J.S. 1981c. Ecology of black beetle, Heteronychus arator (Coleoptera Scarabaeidae)- population sampling. New Zealand Journal of Agricultural Research, 24(1): 79-86.

King, P.D., Meekings, J.S., and Mercer, C.F. 1982. Effects of whitefringed weevil (Graphognathus leucoloma) and black beetle (Heteronychus arator) populations on pasture species. New Zealand Journal of Agricultural Research 25: 405-414.

Matthiessen, J. N. and Learmonth, S.E. 1993. Spatial sampling of insects, plant parts and insect attacks in the soil of potato crops. Bulletin of Entomological Research 83 (4): 607-612.

Matthiessen, J.N. and Ridsdill-Smith, T.J. 1991. Populations of African black beetle, Heteronychus

59 Heteronychus arator Primary Pest of Corn Arthropods African black beetle Beetle

arator Coleoptera: Scarabaeidae) in a Mediterranean-climate area of Australia. Bulletin of Entomological Research 81: 85-91.

Matthiessen, J.N. and Learmonth, S.E. 1998. Seasonally contrasting activity of African black beetle, Heteronychus arator (Coleoptera: Scarabaeidae): Implications for populations, pest status and management. Bulletin of Entomological Research 88 (4): 443-450.

Matthiessen, J., and Learmonth, S. 2005. African black beetle. http://www.agric.wa.gov.au/pls/portal30/docs/Folder/IKMP/PW/INS/PP/FN017_1991.htm. (Last Assessed May 8, 2009).

Smith, T.J., Petty, G.J., and Villet, M.H. 1995. Description and identification of white grubs (Coleoptera: Scarabaeidae) that attack pineapple crops in South Africa. African Entomology 3: 153-166.

Southwood, T.R.E. 1978. Ecological methods with particular reference to the study of insect populations. 524 pp. London, Chapman and Hall.

Toit, H.D., Allsopp, P.G., Rogers, D.J., and Robertson, L.N. 1997. Habitat preference of African black beetle and other soil insect pests of maize in South Africa. Soil invertebrates in 1997. Proceedings of the 3rd Brisbane Workshop on Soil Invertebrates. Pp. 44-47. Brisbane, AU.

Venter, R.J.H. and Louw, M. 1978. Heteronychus arator (Fabricius), a potential dangerous pest on potatoes (Coleoptera: Scarabeidae). Phytophylactica 10: 99.

Watson, R.N. 1980. Dispersal and distribution of Heteronychus arator in New Zealand (Coleoptera: Scarabaeidae). Proc. 2nd Australas. Conf. Grassl. Invertebr. Ecol. 149-152.

60 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

Ostrinia furnacalis

Scientific name Ostrinia furnacalis Guenée

Synonyms Botys damoalis, Botys furnacalis, Botys salentialis, Hapalia damoalis, Micractis nubilalis, Micractis varialis, Ostrinia damoalis, Ostrinia salentialis, Pyrausta damoalis, Pyrausta furnacalis, Pyrausta nubilalis, Pyrausta nubilalis salentialis, Pyrausta polygoni, Pyrausta salentialis, Pyrausta vastatrix, and Spilodes kodzukalis.

Originally, the Asian corn borer was described as Botys furnacalis by Guenée in 1854. Subsequently, it was re-described by a series of authors or repeatedly misidentified as the European corn borer, Ostrinia nubilalis. Much of the literature prior to 1966 treats O. furnacalis as that species. Muturra and Munroe (1970) revised the genus Ostrinia, confirming the status of O. furnacalis as a valid species separate from the European corn borer and synonymizing it with Botys (Pyrausta) damoalis, Botys (Pyrausta) salentialis, Pyrausta polygoni, P. vastatrix, and Spilodes kodzukalis. Many other variants and combinations of these names are present in the literature.

Common names Asian corn borer, Asian maize borer, Asiatic corn borer, China corn borer, and Oriental corn borer

Type of pest Moth

Taxonomic position Class: Insecta, Order: Lepidoptera, Family: Crambidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List - 2009 Figure 1. Pest Description O. furnacalis egg Eggs: Individually measuring about 0.6 to 1 mm in mass. Photo diameter, eggs are laid in white masses with eggs courtesy of James overlapping one another in a fish-scale like pattern. These Litsinger (CABI, masses usually contain 20-40 eggs (Fig.1) (Nafus and 2007). Schreiner, 1991).

Larvae: Newly hatched larvae are about 1-2 mm long, with a dark brown head and white body. As larvae develop, the head capsule gets progressively lighter in color (Fig. 2). Mature larvae are about 19-25 mm long, with a medium to dark brown head, and creamy white to gray body. Raised, darkened spots are evident on the body.

61 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

st Figure 2. 1 instar larvae (left) and late instar larvae of O. furnacalis within a corn stem (right). Notice the darkened lateral spots. Photos courtesy of James Litsinger and www.maizedoctor.cimmyt.org, respectively.

Pupae: Pupae are 12-17 mm long (Lee at al., 1980), medium to dark brown, rounded at the head end, and with a tiny hook at the tail end of the abdomen (Fig. 3).

Adults: Wing span of male moths is 20-26 mm; female moth wings are normally 26-30 mm. The forewings are straw-colored to brown, and the hindwings are lighter yellow to white in color (Fig. 4). Females are lighter in color than males. There is some geographical variation in wing marking. Male specimens from the tropics have more reddish-brown scales intermixed with the ground color, which makes the line markings more obvious. In female specimens, the lines and markings are more fuscous.

Biology and Ecology The biology of O. furnacalis is reviewed by Nafus and Schreiner (1991).

Ostrinia furnacalis is a common pest of corn in the Asian-Western Pacific region, distributed as far south as Australia, as far west as Afghanistan, and as far east and north as the Far East of Russia (CABI, 2007). Infestations in Africa have also been reported. O. furnacalis has one to a few distinct generations per year in temperate zones; these populations undergo diapause. Populations in the tropics without distinct wet and dry seasons can see as many as 12 continuous and overlapping generations. Rainfall increases moth activity and oviposition, and moisture is the likely trigger for the start and end of diapause (Hussein et al., 1983; Hussein and Kameldeer, 1988; Sapin et al., 2006).

Females tend to oviposit on taller (2.5 to 5 feet (75-150 cm) corn plants, mainly on the underside of fully expanded middle to upper leaves (Hussein et al., 1983; Legacion and Gabriel, 1988) and occasionally on leaves’ upper surfaces and on husks. Oviposition usually begins two to four weeks after planting (Nafus and Schreiner, 1987) and is

62 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

highest during the mid-whorl to silk stages, peaking at the tassel stage (Meksongsee et al., 1979; Hussein et al., 1983). Individual females can lay more than 1000 eggs in their lifetime. Eggs fully develop in three to four days (Nafus and Schreiner, 1991).

Depending on host availability, diet, season, and weather, full larval development requires from 5-7 instars and 16-46 days (Nafus and Schreiner, 1991; Liu and Hou, 2004). The pupal stage lasts 4-9 days; adults emerge in the late afternoon to early evening and begin to mate 19-24 hours later. Oviposition begins 1 or 2 days after mating (Camarao, 1976). Adults are active throughout the night, and can live up to 14 days (females tend to live longer than males).

Larvae are able to move from plant to plant via silk, on self-created threads via wind and on strands already attached to multiple plants. If a population Figure 3. Top: pupa of O. furnacalis. undergoes diapause, it does so in the Below pupa is an adult (Euborellia larval stage, usually in corn stubble and stali) a predator of these pupae. Bottom: stalks (Lee et al., 1980; Nafus and Adult O. furnacalis. Photos courtesy of Schreiner, 1991; Li et al., 1999). Long James Litsinger and K.V.N. Maes, distance dispersal and immigration into respectively (CABI, 2007) corn fields is also common for adult moths, especially when they first emerge.

Almost any part of the plant can be consumed by O. furnacalis larvae, but feeding on the whorl, tassels, ears and stalk are the areas that significantly affect crop yield; O. furnacalis rarely, if ever, feeds on the roots. First and second instar larvae feed within the whorl and, if present, the tassel and ears. They feed on the anthers, silk, and leaves. Later instars feed on these structures, but also bore into the stem and the shank/cob of the ear. Damage to corn by O. furnacalis can start as early as two to four weeks after germination. As the growing season progresses, feeding moves up the plant until the corn has shed its pollen; after this, most of the tunneling into ears and stalks occurs (Nafus and Schreiner 1987; Schreiner and Nafus, 1987; Li et al., 1999).

63 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

Symptoms/Signs Symptoms are similar to that of the European corn borer, Ostrinia nubilalis. Larvae most often feed on and/or in the tassels, whorl, leaf sheath (Fig. 4), stalk and ear. Older larvae tend to burrow into the stalk, or into the ear cob or kernels. Feeding by later larvae instars is usually considered to be most damaging, but tunneling by even young larvae can result in broken tassels (Fig. 4) and, less often, lodging. Yield losses are greatest when damage occurs at the reproductive stages.

Pest Importance Ostrinia furnacalis is an important pest in much of Asia on maize and sweet corn. It is a consistent problem in China, Japan, Korea, the Philippines and several other Pacific Islands, as well as East Africa. Up to 100% loss has been reported, but reported yield losses of up to 40% are much more common. Because of its broad ecological range in both climate and host, O. furnacalis can have 1-12 generations per year; different predictive models and economic threshold values exist due to this variation. The number of egg masses per plant (Meksongsee et al., 1979; Hussein and Kameldeer, 1988; Morallo-Rejesus et al., Figure 4. Shotholing (top) 1990), the number of cavities bored into the and broken tassels (bottom) plant (Hsu et al., 1988), the number of larvae are common symptoms of O. per plant (Morallo-Rejesus et al., 1990) and furnacalis infestation. Photos sea surface temperature (Liu et al., 1999) courtesy of James Litsinger. have all been used to predict when action is needed to minimize yield loss from O. furnacalis. Cou nting egg masses and cavities are easiest for field workers. In the Philippines, 35% of plants with egg masses signaled the immediate need to treat with insecticide, whereas in Thailand, the threshold was 15%. In both cases, a minimum of 100 plants were sampled. In some regions, up to 5 cavities per plant did not produce significant yield reductions in field corn; in others, as little as 1 cavity reduced yield (in field and sweet corn).

Historically, insecticides (including carbamates, trichlorfon, granular carbaryl, phoxim and Bt) have been the most often used control for this pest. However, in many tropical regions the larvae are a continuous presence (due to year round, overlapping generations), and oftentimes feed inside plant structures where contact insecticides are

64 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth not effective (Nafus and Schreiner, 1989). Therefore, successful insecticidal treatments need to be frequently applied, well-timed and accurate, even in regions where distinct generations occur. When used alone, insecticides are rarely the most economical solution. Therefore, a combination of chemical, biological and cultural control (including transgenic and/or resistant corn varieties) has been researched and implemented to control O. furnacalis.

Many field studies and long-term control efforts using biological control organisms have been recorded and studied. In Asia, egg parasitoids have been widely used (Tran et al., 1988; Zhang, 1988; J. Zhang et al. 1990a), including mass production and inundative releases (Tseng, 1990; J. Zhang et al., 1990b) and in combination with cultural controls and insecticides (Tandan and Nillama, 1987; Lu et al., 1989). Nematodes (Hu et al., 2004), fungi (A.W. Zhang et al., 1990) and (Fig. 3) (Camarao, 1976) have also been investigated and found to be successful against this pest, although the nematodes and fungi in a true production environment have not been reported.

Detasselling, which removes a significant food source of larvae, has also been widely used to reduce the damage by this borer (Schreiner and Nafus 1987, 1988; Felkl, 1988). Manipulating the planting dates to later in the growing season has also been successful. Transgenic corn varieties (expressing Cry1A and the Cry1Ab gene) have seen significant success in decreasing damage by O. furnacalis and increasing corn yield (Fernandez et al., 1999; Kanglai et al., 2003) without having a detrimental effect on beneficial insects due to the decline in O. furnacalis populations (Reyes and Mostoles, 2005) or pollen feeding (Wang et al., 2007). Inbred lines of corn are also available that are resistant to leaf and tassel feeding (Nafus and Schreiner, 1989; Kuo et al., 1990).

Pheromone traps have also been developed in hopes of widespread mating disruption, but no published reports have shown they can be successful in the control of this moth.

Known Hosts Due to confusion about the of O. furnacalis, there are issues with the reported host range. Some host records may actually be from other species of Ostrinia. Studies are ongoing to sort out the host range. Schreiner et al. (1990) report that the growth rate and survival of the O. furnacalis larvae were greater on sweet corn ears than on any other potential host tested. The second best host was Johnson grass (Sorghum halepense). Only two larvae completed development on pepper and Brachiaria mutica (para grass); while only one developed on wildcane (Saccharum spontaneum). In this study, although reported as minor hosts, (Indian goosegrass), Pennisetum purpureum (elephant grass), and Phragmites karka (tall reed) did not support any Asian corn borers through their complete development.

The following are the reported hosts according to the literature at this time.

Major hosts Panicum miliaceum (millet), Sorghum bicolor (sorghum), Sorghum halepense (Johnson

65 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

grass), Zea mays (maize/corn), Zingiber officinale (ginger) (Lewvanich, 1973; Young, 1979; Schreiner et al., 1990; Nafus and Schreiner, 1991; Talekar et al., 1991).

Minor hosts Amaranthus spp., Artemisia spp. (wormwoods), Apocynum cannabium (Indian hemp), Artemesia spp., Blumea lacera, Brachiaria mutica (buffalo grass), Cannabis sativa (hemp), Capsicum annuum (bell pepper), Coix lacryma-jobi (Job's-tears), Eleusine indica (finger millet), Emex spp. (Emex), Gossypium hirsutum (cotton), (hops), Oryza sativa (rice), Panicum virid, Pennisetum glaucum (pearl millet), Pennisetum spp. (feather grass), Phaseolus spp. (bean), Phragmites karka (tall reed), Phytolacca spp. (pokeweed), Polygonum spp. (knotweed), Polytoca macrophylla, Populus spp. (poplars), Rheum rhabarbarum (rhubarb), Rumex dentatus, Saccharum officinarum (sugarcane), Saccharum robustum, Saccharum spontaneum (wild sugarcane), Setaria italica (foxtail millet), Setaria viridis (barbed bristlegrass), Solanum melongena (eggplant), Themeda intermedia, Urochloa mutica (tall panicum), and Vigna sinensis (cowpea).

Known Vectors (or associated organisms) Ostrinia furnacalis is not a known vector and does not have any associated organisms. However, the injuries produced by this borer may increase fungal and bacterial infections (Dalmacio et al., 2007).

Known Distribution Asia: Afghanistan, Brunei Darussalam, Cambodia, China, India, Indonesia, Japan, Korea, Laos, Malaysia, Myanmar, Pakistan, Philippines, Singapore, Sri Lanka, Thailand and Vietnam. Europe: Russian Federation. Oceania: Australia, Micronesia, Guam, Northern Mariana Islands, Papa New Guinea, and Solomon Islands.

Potential Distribution within the United States A recent risk map analysis by USDA-APHIS-PPQ-CPHST indicates that areas of Alabama, Arkansas, California, Colorado, Florida, Georgia, Illinois, Indiana, Iowa, Kansas, Kentucky, Louisiana, , Maryland, Michigan, Minnesota, Mississippi, Missouri, Nebraska, New York, North Carolina, North Dakota, Ohio, Pennsylvania, South Carolina, South Dakota, Tennessee, Texas, Utah, and Wisconsin have the greatest risk for O. furnacalis establishment based on host availability and climate within the United States. In general, other than those areas with very high elevations, the entire contiguous United States is at least a moderate risk for O. furnacalis.

Survey CAPS-Approved Method: Trap with lure. A wing trap is the approved trap for Ostrinia furnacalis. The lure information is provided below:

Lure Compound Dispenser Load Dispenser Type Lure Abbreviation a) E12-14:Ac a) 0.06 mg gray rubber septum ACB b) Z12-14:Ac b) 0.1 mg

66 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

MPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters.

Literature-Based Methods: Ostrinia furnacalis can be identified by egg masses, larvae, damage and with pheromone traps that attract adult males. Because each of its life stages are very similar to the European corn borer (Ostrinia nubilalis) and traps that are used for O. furnacalis also trap an occasional O. nubilalis male, proper identification (via adult genitalia, if possible) is still necessary in areas that have no known establishment of O. furnacalis.

Trapping: The components of the sex pheromone of O. furnacalis have been analyzed (Cheng et al., 1981; Ruo et al., 1983; Du et al., 1986; Zhu et al., 1987; Yeh et al., 1989; Zhao et al.,1990). While all populations of O. furnacalis produce a pheromone with a mixture of (E) - and (Z)-12-tetradecanyl acetates, in many instances different populations produce and are attracted to significantly different proportions of these two stereoisomers. Reports from Japan (Huang et al., 1998) and China (Cheng et al., 1981) report that populations prefer pheromones containing 36-39%, 44%, and 54% of the (E) stereoisomer. Boo and Park (1998) summarize the ratios of the two stereoisomers used in six studies for O. furnacalis.

In addition, a third compound, tetradecanyl acetate was discovered and used in the pheromone mixture to attract males in Taiwan (Kou et al., 1992). The addition of tetradecanyl acetate did not enhance or suppress male response in the study. The addition of the compound in its natural ratio, however, resulted in a decrease in trap catches in China (Cheng et al., 1981; Chen et al., 1982).

Kou et al. (1992) used plastic tubes containing (E) - and (Z)-12-tetradecanyl acetates and tetradecanyl acetate in a 48:37:15 ratio inside wing-shaped sticky traps. Traps were placed 32.8 feet (10 m apart) and 2.6-3.3 feet (0.8-1 m) above the ground. Catches were compared with blank traps and traps baited with 2- to 3-day old virgin females every two days. Cheng et al. (1981) used simple water traps (20-cm diameter vessel filled with water and with detergent added to reduce surface tension). A paper roll was impregnated with the pheromonal solution and supported 1-1.5 cm above the water surface. Trap height ranged from 2.6-3.3 feet (0.8-1 m), depending on the height of the plants, and lures were changed each night. Jackman et al. (1983) tested four types of traps: metal basin trap, plastic basin trap, Pherocon® 1C sticky trap, and Biotrap® sticky traps. No significant differences were observed between traps, although differences did exist between bait used (virgin female versus synthetic pheromone). Jackman et al. (1984) also used water traps. A circular plastic pan 30 cm in diameter and 9 cm deep was placed in a wood frame at 1.2 meters high and supplied with a 1 cm thick wooden cover 8 cm above the pan top. Basins were filled to 5 cm with water and about 1 teaspoon of commercial laundry detergent was added.

67 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

Russell IPM (United Kingdom) manufactures a pheromone lure for O. furnacalis. They suggest that using a Delta trap is the most sensitive trap to use for monitoring this insect. A bucket trap, however, may be used in dusty conditions or where there is a high moth population density. Two traps per hectare are recommended for small fields or fields with uneven topography; while one trap per two hectares is recommended for large scale field and homogenous land. The trap should be placed near the highest point of the plant using support posts approximately 1 meter high or higher if the crop is higher. See http://www.russellipm-agriculture.com/insect.php?insect_id=211&lang=en for additional information.

For the European corn borer (ECB) (Ostrinia nubilalis), traps used also include various 'cone' style traps, including the Scentry (cloth) design and the Harstack (metal screen) trap. These are currently used mostly for scouting (determining population levels in a field as a prelude to deciding whether or not to apply a treatment). Black light (UV) traps have also been used for ECB monitoring, but these traps catch many insect pests including moths.

Visual survey: O. furnacalis are detected in the field by surveying standing crops for egg batches or damage by larvae (cavities).

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of O. furnacalis is by morphological identification.

Literature-Based Methods: Because of their similarities as larvae and adults, adult genitalia must be examined to differentiate between Ostrinia species (Mutuura and Munroe, 1970).

Easily Confused Pests O. furnacalis is closely related to O. nubilalis, the European corn borer (Fig. 6). It is thought to have similar biology and ecology, and a number of parasites in common. When pheromone traps for O. furnacalis are placed in the field, adult males of the European corn borer are often caught.

Figure 6. Larvae (left) and adults (right) of Ostrinia nubilalis, the European corn borer (Frank Peairs, Colorado State University, www.bugwood.org).

68 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

References Boo, K.S. and Park, J.W. 1998. Sex pheromone composition of the Asian corn borer moth, Ostrinia furnacalis (Guenee) (Lepidoptera: Pyralidae) in South Korea. Journal of Asia-Pacific Entomology 1: 77- 84.

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Camarao, G.C. 1976. Population dynamics of the cornborer, Ostrinia furnacalis (Guenée), I. Life cycle, behavior, and generation cycles. Philippine Entomologist 3(3/4): 179-200.

Chen, Z.Q., Xiao, J.C., Huang, X.T., Huang, L.G., Yang, Z.X., and Zhang, H.L. 1982. Attraction of synthetic sex pheromone components to male corn borer, Ostrinia furnacalis Guenee in the field. Acta Entomologica Sinica 25(2): 164-171. English Abstr.

Cheng, Z.Q., Xiao, J.C., Huang, X.T., Chen, D.L., Li, J.Q., He Y.S., Huang, S.R., Luo, Q.C., Yang, C.M. and Yang, T.H. 1981. Sex pheromone components isolated from China corn borer, Ostrinia furnacalis Guenee (Lepidoptera: Pyralidae), (E)- and (Z)-12-tetradecenyl acetates. Journal of Chemical Ecology 7(5):841-851.

Dalmacio, S.C., Lugod, T.R., Serrano, E.M. and Munkvold, G.P. 2007. Reduced incidence of bacterial rot on transgenic insect-resistant maize in the Philippines. Plant Disease 91(4): 346-351.

Du, J.W., Zhu, Y.X., Zang, T.P., Xu, S.F. and Dai, X.L. 1986. Studies on the precise blending of the pheromone components of the Asian corn borer, Ostrinia furnacalis Guenée (Lep. Pyralidae). Contributions from the Shanghai Institute of Entomology 6: 17-22.

Felkl, G. 1988. Economic aspects of detasselling corn plants and insecticide use to control Asian corn borer, Ostrinia furnacalis Guenée. Journal of Applied Entomology 105(4): 379-386.

Fernandez, E.C., Logrono, M.L., Lapiz, R.V., Maligalig, E.R., Deleus, E. and Ancheta, R.T. 1999. Resistance of tropical corn hybrids with Bt gene against the Asiatic corn borer, Ostrinia furnacalis Guenée. Philippine Journal of Crop Science 24 (supplement 1): 25.

Hsu, S. L., Peng, W.K. and Hsieh, F.K. 1988. Loss assessment of corn infested with Asian corn borer, Ostrinia furnacalis (Guenée). Memoirs of the College of Agriculture, National Taiwan University 28(2): 27- 31.

Hu, X., Liang, G. and Ling, Z. 2004. Studies on the control efficacy of Steinernema feltiae against Ostrinia furnacalis. Journal of South China Agricultural University 25(2): 52-55.

Huang, Y., Takanashi, T., Hoshizaki, S., Tatsuki, S., Honda, H., Yoshiyasu, Y. and Ishikawa, Y. 1998. Geographic variation in sex pheromone of Asian corn borer, Ostrinia furnacalis, in Japan. Journal of Chemical Ecology 24(12): 2079-2088.

Hussein, M.Y., Kamaldeer, A.K., and Ahmad, N.M. 1983. Some aspects of the ecology of Ostrinia furnacalis (Guenée) on corn. MAPPS Newsletter 7(2): 11-12.

69 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

Hussein, M.Y. and Kameldeer, A.K. 1988. A field study on the oviposition of Ostrinia furnacalis Guenée (Lepidoptera; Pyralidae) on maize in Selangor, Malaysia. Tropical Pest Management 34(1): 44-47, 115- 120.

Jackman, J.A., Benigno, E.A., Klun, J.A., and Schwarz, M. 1983. Field tests of (Z) and (E)-12- tetradecen-1-ol acetate as a sex attractant for Asian corn borer, Ostrinia furnacalis (Guenee). Philippine Entomologist 5&6: 519-530.

Jackman, J.A., Acree, T.A., and Benigno, E.A. 1984. Evidence of additional sex pheromone components in the Asian corn borer, Ostrinia furnacalis (Guenee). Philippine Entomologist 6(1): 39-45.

Kanglai, H.E., Zhenying, W., Darong, Z., Liping, W., Yanying, S. and Yao, Z. 2003. Evaluation of transgenic Bt corn for resistance to the Asian corn borer (Lepidoptera: Pyralidae). Journal of Economic Entomology 96(3): 935-940;

Kou, R., Ho, H.Y., Yang, H.T., Chow, Y.S., and Wu, H.J. 1992. Investigation of sex pheromone components of female Asian corn borer, Ostrinia furnacalis (Hübner) (Lepidoptera: Pyralidae) in Taiwan. Journal of Chemical Ecology 18(6): 833-840.

Kuo, N.C, Chen, C., and Tseng, C.T. 1990. The evaluation, comparison and stability analysis of corn- leaf resistance to the Asian corn borer (Ostrinia furnacalis Guenée). Memoirs of the College of Agriculture, National Taiwan University, 30(1): 114-122.

Lee, Y.B., Hwang, C.Y., Choi, K.M., and Shim, J.Y. 1980. Studies on the bionomics of the Oriental corn borer Ostrinia furnacalis (Guenee). Korean Journal of Plant Protection 19(4): 187-192.

Legacion, D.M. and Gabriel, B.P. 1988. Oviposition of Asiatic corn borer moths on corn plants (note). Philippine Agriculturalist 71(3): 375-378.

Lewvanich, A. 1973. A study of the identity of the corn stem borer in Thailand. Thai Journal of Agricultural Science 7: 103-109.

Li, Q., Gao, Z., Wang, W., and Cui, L. 1999. Sequential sampling and control of corn borers in cotton fields. Journal of Henan Agricultural Sciences 12:13-14.

Liu, L., Huang, S., and Kong, F. 1999. A SST based forecasting model for G3 maize borer. Journal of Nanjing Institute of Meteorology 22(2): 264-268.

Liu, S. and Hou, Z. 2004. Observation on bionomics of Ostrinia furnacalis. Entomological Knowledge 41(5): 461-464.

Lu, X., Wang, G.X., and Lei, P. 1989. Contact toxicity of eight insecticides to Macrocentrus linearis (Hym.: Brachonidae), a parasite of the corn borer, Ostrinia furnacalis (Lep.:Pyralidae). Chinese Journal of Biological Control 5(3): 123-124.

Meksongsee, B., Poonyathavorn, P., Prachaubmoh, O., Kongkanjana, A., Chawanapong, M., Wongkobrat, A., Wongkamhaeng, W., and Weerawut, T. 1979. Corn and sorghum Insect. 11th Annual Thai National Corn and Sorghum Program Annual Report. pg 252-274.

Morallo-Rejesus, B., Buctuanon, E.M., and Rejesus, R.S. 1990. Defining the economic threshold determinants for the Asian corn borer, Ostrinia furnacalis (Guenée) in the Philippines. Tropical Pest Management 36(2): 114-121.

70 Ostrinia furnacalis Primary Pest of Corn Arthropods Asian corn borer Moth

Mutuura, A. and Munroe, E. 1970. Taxonomy and distribution of the European corn borer and allied species: genus Ostrinia (Lepidoptera: Pyralidae). Memoirs of the Entomological Society of Canada, No. 71, 112 pp.

Nafus, D.M. and Schreiner, I. H. 1987. Location of Ostrinia furnacalis (Lepidoptera: Pyralidae) eggs and larvae on sweet corn in relation to plant growth stage. Journal of Economic Entomology 80: 411-416.

Nafus, D.M. and Schreiner, I.H. 1989. Management of Ostrinia furnacalis on Corn in Micronesia. Tropical and Subtropical Agricultural Research Under PL 89-106, Special Research Grants. Progress and Achievement, the Pacific Basin Group. pp 20-21.

Nafus, D.M. and Schreiner, I. H. 1991. Review of the biology and control of the Asian corn borer, Ostrinia furnacalis (Lep.: Pyralidae). Tropical Pest Management 37(1):41-56.

Reyes, S.G and Mostoles, M.D.A. 2005. Diversity, community structure and wet season population abundance of insect on Bt-corn agroecosystem in two sites on Luzon Island, Philippines. The Asian International Journal of Life Sciences 14(1): 55-73.

Ruo, H.C., Tian, C., and Hao, D.Q. 1983. The discrimination of the predominant species of corn borers in Ningxia by sex pheromones. Ningxia Agricultural Science and Technology (Ningxia Nongye Keji) 5:13-16.

Sapin, G.B., Adalla, C.B., and Alzona, F.D. 2006. Life history and feed ovipositional preference of Asian corn borer for Bt and non-Bt corn. Philippine Entomologist 20(2): 179-180.

Schreiner, I.H. and Nafus, D.M. 1987. Detasselling and insecticides for control of Ostrinia furnacalis (Lepidoptera: Pyralidae) on sweet corn. Journal of Economic Entomology 80(1): 263-267.

Schreiner, I.H. and Nafus, D.M. 1988. No-tillage and detasselling: effect on the Asian corn borer Ostrinia furnacalis and ants. Philippine Entomologist 7(4): 435-442.

Schreiner, I.H., Nafus, D.M., and Dumaliang, N. 1990. Growth and survival of the Asian corn borer Ostrinia furnacalis Guenee (Lep: Pyralidae) on alternative hosts in Guam. Tropical Pest Management 36(2): 93-96.

Talekar, N.S., Lin, C.P., Yin, Y.F., Ling, M.Y., Wang, Y.D., and Chang, D.C.Y. 1991. Characteristics of infestation by Ostrinia furnacalis (Lepidoptera: Pyralidae) in mungbean. Journal of Economic Entomology 84(5): 1499-1502.

Tandan, J.S. and Nillama, N.C. 1987. Biological control of Asiatic corn borer (Ostrinia furnacalis Guenée) and corn earworm (Helicoverpa armigera Hubner). CMU Journal of Agriculture, Food and Nutrition 9(1): 33-48.

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Tseng, C.T. 1990. Use of Trichogramma ostriniae (Hym., Trichogrammatidae) for controlling the oriental corn borer, Ostrinia furnacalis (Lep., Pyralidae) in Taiwan, China. FFTC-NARC International Seminar in ‘The use of parasitoids and predators to control agricultural pests,’ Tukuha Science CityIbaraki-ken, 305 Japan, October 2-7, 15pp.

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72 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

Spodoptera littoralis

Scientific Name Spodoptera littoralis Boisduval

Synonyms: Hadena littoralis, Noctua gossypii, Prodenia littoralis, Prodenia litura, Prodenia retina, Spodoptera retina, and Spodoptera testaceoides

The two Old World cotton leafworm species S. littoralis and S. litura are allopatric, their ranges covering Africa and Asia, respectively. Many authors have regarded them as the same species.

Common Name(s) Cotton leafworm, Egyptian cotton leafworm, Mediterranean climbing cutworm, tobacco caterpillar, tomato caterpillar, Egyptian cotton worm, Mediterranean brocade moth, and Mediterranean climbing cutworm

Type of Pest Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Family: Noctuidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description Eggs: Spherical, somewhat flattened, 0.6 mm in diameter, laid in clusters arranged in more or less regular rows in one to three layers, with hair scales derived from the tip of the abdomen of the female moth (Fig. 1). The hair scales give the eggs a “felt-like appearance”. Usually Figure 1. Spodoptera egg masses (top); whitish-yellow in color, changing to black just eggs and neonates (bottom). Eggs are prior to hatching, due to the big head of the laid in batches covered with orange- larva showing through the transparent shell brown hair scales. Photo courtesy of (Pinhey, 1975). Bernard Fransen, www.invasive.org, and Larvae: Upon hatching, larvae are 2-3 mm long http://www.defra.gov.uk/planth/pestnote/ with white bodies and black heads and are very spod.htm, respectively.

73 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth difficult to detect visually. Larvae grow to 40 to 45 mm and are hairless, cylindrical, tapering towards the posterior and variable in color (blackish- gray to dark green, becoming reddish- brown or whitish-yellow) (Fig. 2). The sides of the body have dark and light longitudinal bands; dorsal side with two dark semilunar spots laterally on each segment, except for the prothorax; and spots on the first and eighth abdominal segments larger than the others, interrupting the lateral lines on the first Figure 2. Larva of S. littoralis. Photos segment. The larva of S. littoralis is courtesy of Biologische Bundesanstalt figured by Bishari (1934) and Brown and für Land-und Forstwirtschaft Archive, Dewhurst (1975). Larvae are nocturnal Biologische Bundesanstalt für Land-und and during the day can be found at the Forstwirtschaft, www.bugwood.org. base of the plants or under pots.

Pupae: When newly formed, pupae are green with a reddish color on the abdomen, turning dark reddish-brown after a few hours (Fig. 3). The general shape is cylindrical, 14-20 x 5 mm, tapering towards the posterior segments of the abdomen. The last segment ends in two strong straight hooks (Pinhey, 1975).

Figure 3. Pupae of S. littoralis (A) and pupa with adult on soil (B). Photos courtesy of Esmat M. Hegazi, University of Alexandria, www.bugwood.org and CABI, 2007.

Adults: Moth with gray-brown body (Fig. 4), 15 to 20 mm long; wingspan 30 to 38 mm; forewings gray to reddish brown with paler lines along the veins (in males, bluish areas occur on the wing base and tip); the ocellus is marked by two or three oblique whitish stripes. Hindwings are grayish white, iridescent with gray margins, and usually lack darker veins (EPPO, 1997).

A B

74 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

Figure 4. Adult moths of S. littoralis. Photos courtesy of Entopix and Bernard Fransen, www.invasive.org, respectively.

Biology and Ecology S. littoralis is a multivoltine species that does not enter a diapause stage. Female moths lay most of their egg masses (20-1,000 eggs) on the lower leaf surface of younger leaves or upper parts of the plant. Anderson and Alborn (1999) showed that S. littoralis preferred to oviposit on small plants (3-4 leaves) that had been fed upon by 3rd or 4th instar larvae (72%) over non-damaged control plants. When using larger plants (8 to 10 true leaves), however, the preference was reversed with only 30% of eggs deposited on induced (previously fed upon) plants. Eggs begin to hatch after 28.6 degree days (DD) at a base temperature of 14.8°C (59°F). The optimal temperature for egg hatch is 28- 30°C (82-86°F).

As the insect develops, it completes six instars. Early instars remain on the underside of leaves and feed throughout the day. On cotton, the first three larval instars feed mainly on the lower surface of the leaves, whereas later instars feed on both surfaces. Third and fourth instars remain on a plant, but do not feed during the daylight; later instars migrate off the plant and rest in the soil during the day and return to the plant at night.

Upon pupation, the fully grown larva pushes downward on the loose surface of the soil until it reaches more solid ground 3-5 cm deep. It then creates a clay ‘cell’ or cocoon in which it usually pupates within 5-6 hours. Emergence of adult moths occurs at night, and they have a life span of 5-10 days. Adults fly at night, mostly between the hours of 8 pm and midnight.

Symptoms/Signs On most crops, damage arises from extensive feeding by larvae, leading to complete stripping of the plants. Damage of S. littoralis consists of feeding scars and skeletonizing caused by feeding on the undersides of the leaves. Initially there are numerous small feeding points, which finally spread over the entire leaf. Later holes and bare sections are found on leaves, young stalks, bolls, and buds resulting from feeding activities of the pest. In certain cases, the shoot tips above a hole wilt and eventually die.

75 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

Corn stems are often mined by S. littoralis and young grains in the ear may also be damaged.

Pest Importance S. littoralis is one of the most destructive agricultural lepidopterous pests within its subtropical and tropical range. The pest causes a variety of damage as a leaf feeder and sometimes as a cut worm on seedlings. It can attack numerous economically important crops throughout the year (EPPO, 1997). On cotton, the pest may cause considerable damage by feeding on the leaves, fruiting points, flower buds, and occasionally on bolls. When peanuts are infested, larvae first select young folded leaves for feeding, but in severe attacks, leaves of any age are stripped off. Sometimes, even the ripening kernels in the pods in the soil may be attacked. Pods of and the seeds they contain are also often badly damaged. In tomatoes, larvae bore into the fruit, rendering them unsuitable for consumption. Numerous other crops are attacked, mainly on their leaves.

In Europe, damage caused by S. littoralis was minimal until about 1937. In 1949, there was a catastrophic population explosion in southern Spain, which affected alfalfa, potatoes, and other vegetable crops. At present, this noctuid pest is of great economic importance in Cyprus, Israel, Malta, Morocco, and Spain (except the north). In Italy, it is especially important on protected crops of ornamentals and vegetables (Inserra and Calabretta, 1985; Nucifora, 1985). In Greece, S. littoralis causes slight damage in Crete on alfalfa and clover only. In North Africa, tomato, Capsicum spp., cotton, corn, and other vegetables are affected. In Egypt, it is one of the most serious cotton pests.

Many populations of S. littoralis are extremely resistant to pesticides, and if they become well established, can be exceptionally difficult to control (USDA, 1982).

Known Hosts The host range of S. littoralis covers over 40 families, containing at least 87 species of economic importance (Salama et al., 1970).

Major Hosts Abelmoschus esculentus (okra), Allium spp. (onion), Amaranthus spp., Apios spp. (groundnut), Arachis hypogea (peanut), Beta vulgaris (beet), Brassica oleracea (cabbage, broccoli), Brassica rapa (turnip), Brassica spp. (mustards), Camellia sinensis (tea), Capsicum annuum (pepper), Chrysanthemum spp., Citrullus lanatus (watermelon), Citrus spp., Coffea arabica (coffee), Colocasia esculenta (taro), Corchorus spp. (jute), Cucumis spp. (squash, pumpkin), Cynara scolymus (artichoke), Daucus carota (carrot), Dianthus caryophyllus (carnation), Ficus spp. (fig), Glycine max (soybean), Gossypium spp. (cotton), Helianthus annus (sunflower), Ipomoea batatas (sweet potato), Lactuca sativa (lettuce), Linum spp. (flax), Medicago sativa (alfalfa), Morus spp. (mulberry), Musa spp. (banana, plantain), Nicotiana tabacum (tobacco), Oryza sativa (rice), Pennisetum glaucum (pearl millet), Persea americana (avocado), Phaseolus spp. (bean), Pisum sativum (pea), Prunus domestica (plum), Psidium guajava (guava), Punica granatum (pomegranate), Raphanus sativus (radish), Rosa

76 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth spp. (rose), Saccharum officinarum (sugarcane), Solanum esculentum (tomato), Solanum melongena (eggplant), Solanum tuberosum (potato), Sorghum bicolor (sorghum), Spinacia spp. (spinach), Theobroma cacao (cacao), Trifolium spp. (clover), Triticum aestivum (wheat), Vicia faba (broad bean), Vigna spp. (cowpea, black-eyed pea), Vitis vinifera (grape), and Zea mays (corn).

Minor Hosts Acacia spp. (wattles), Actinidia arguta (tara vine), Alcea rosea (hollyhock), Anacardium occidentale (), Anemone spp. (anemone), Antirrhinum spp., Apium graveolens (celery), Asparagus officinalis (asparagus), Caladium spp. (caladium), Canna spp. (canna), Casuarina equisetifolia (she-oak), Convolvulus spp. (morning glory, bindweeds), Cryptomeria spp. (Japanese cedar), Cupressus spp. (cypress), Datura spp. (jimsonweed), Eichhornia spp. (water hyacinth), Eucalyptus spp. (eucalyptus), Geranium spp. (geranium), Gladiolus spp. (gladiolus), Malus domesticus (apple), Mentha spp. (mint), Phoenix dacylifera (date palm), Pinus spp. (pine), and Zinia spp. (zinnia).

Known Vectors (or associated organisms) S. littoralis is not a known vector and does not have any associated organisms.

Known Distribution The northerly distribution limit of S. littoralis in Europe corresponds to the climatic zone in which winter frosts are infrequent. It occurs throughout Africa and extends eastwards into Turkey and north into eastern Spain, southern France and northern Italy. However, this boundary is probably the extent of migrant activity only; although the pest overwinters in southern Spain, it does not do so in northern Italy or France. In southern Greece, pupae have been observed in the soil after November and the species overwinters in this stage in Crete. Low winter temperatures are, therefore, an important limiting factor affecting the northerly distribution, especially in a species with no known diapause (Miller, 1976; Sidibe and Lauge, 1977).

Africa: Algeria, Angola, Benin, Botswana, Burkina Faso, Burundi, Cameroon, Cape Verde, Central African Republic, Chad, Comoros, Congo, Cote d’Ivoire, Egypt, Equatorial Guinea, Eritrea, Ethiopia, Gabon, Gambia, Ghana, Guinea, Kenya, Liberia, Libya, Madagascar, Malawi, Mali, Mauritania, Mauritius, Morocco, Mozambique, Namibia, Nigeria, Reunion, Rwanda, Senegal, Siera Leone, Somalia, South Africa, Sudan, Swaziland, Tanzania, Togo, Tunisia, Uganda, Zaire, Zambia, and Zimbabwe. Asia: Afghanistan, Bangladesh, Brunei, and India. Europe: France, Germany, Greece, Italy, Malta, Portugal, Spain, and United Kingdom. Middle East: Bahrein, Cyprus, Iran, Iraq, Israel, Jordan, Lebanon, Oman, Saudi Arabia, Syria, United Arab Emirates, and Yemen. Oceania: American Samoa.

Potential Distribution in the United States The pest has been intercepted at U.S. ports on plant parts, leaves, and flowers. The potential U.S. range of most S. littoralis may be limited to the west coast through the lower southwestern and southeastern United States, reaching only as far north as

77 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

Maryland (USDA, 1982). Migratory moths may be capable of periodic spread into northern states and even Canada by late summer or early fall. Venette et al. (2003) suggest that approximately 49% of the continental United States would be suitable for S. littoralis. A recent risk map developed by USDA-APHIS-PPQ-CPHST (Fig. 5) shows that portions of Alabama, Arkansas, Arizona, California, Florida, Georgia, Louisiana, Mississippi, Missouri, North Carolina, Oklahoma, South Carolina, and Texas are at the greatest risk from S. littoralis. Portions of most states within the continental United States have moderate risk of S. littoralis establishment based on climate and host range.

Survey CAPS-Approved Method: Trap with lure. A plastic bucket trap [unitrap] with dry kill strip is the approved trap for Spodoptera littoralis. The lure information is provided below:

Lure Dispenser Dispenser Lure Length of Compound Load Type Abbreviation Effectiveness a) 'Z,E,9,11- a) 1.99 mg laminate ECL 12 weeks 14:AC b) 0.01 mg b) 'Z,E,9,12- 14:AC

The plastic bucket trap (also known as the Universal moth trap or unitrap) should have a green canopy, yellow funnel, and white bucket and should be used with dry kill strip. For instructions on using the trap (see Appendix G “Plastic Bucket Trap Protocol”).

IMPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet). As of June 2010, S. litura and S. littoralis lures should be placed in different traps and separated by at least 20 meters (65 feet).

Literature-Based Methods: (From Venette et al., 2003; CABI, 2007) Trapping: Pheromone traps can be used to monitor the incidence of S. littoralis (Rizk et al., 1990). The synthetic sex pheromone (Z,E)-(9,11)-tetradecadienyl acetate has proven highly effective at trapping male moths of S. littoralis (Salem and Salama, 1985). Kehat and Dunkelblum (1993) found that the minor sex pheromone component, (9Z,12Z)-9,12-tetradecadienyl acetate in addition to the major component (9Z,11Z)- 9,11-tetradecadienyl acetate was required to attract males.

Sex-pheromone baited delta traps remained attractive for approximately 2 weeks, but effectiveness declined after 3 to 4 weeks of use (Ahmad, 1988). To monitor male flight activity in vegetable production areas, delta traps were placed 1.7 m above the ground at a rate of 2 traps/ha (approximately 1 trap/acre) (Ahmad, 1988). Pheromone lures

78 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

impregnated with 2 mg of the pheromone blend (blend not specified) were replaced after 4 weeks of use (Ahmad, 1988). Traps are deployed at a similar height (1.5 m) to monitor male flight in cotton (Salem and Salama, 1985). Catches in pheromone traps did not correlate as well with densities of egg-masses in cotton fields as did catches in a black-light trap (Rizk et al., 1990). The attractiveness of traps baited with (Z,E)-(9,11)- tetradecadienyl acetate is governed primarily by minimum air temperature, relative humidity, adult abundance, and wind velocity. Densities of female S. littoralis also affect the number of males that are captured at different times of the year (Rizk et al., 1990). Lures for S. littoralis may also attract Erastria spp. (established in the United States) (PPQ, 1993).

Visual survey: Visual surveys for this pest can take place any time during the growing season while plants are actively growing (usually spring through fall in temperate areas). Early instars (<3rd) are likely to be on lower leaf surfaces during the day. The larvae will skeletonize leaves by feeding on this surface and such damage to the leaf provides evidence of the presence of larvae. Sweep net sampling may be effective at dawn or dusk. Specimen identification should be confirmed by a trained taxonomist (USDA, 1982). However, not all sampling methods are equally effective for all life- stages of the insect. Eggs are only likely to be found by visual inspection of leaves. First through third instars may be detected by sweep net sampling; nearly all instars can be detected by visual inspection of plants; and, later instars (4th-6th) and pupae may be found by sieving soil samples (Abul-Nasr and Naguib, 1968; Abul-Nasr et al.,1971).

Not recommended: Light traps using a 125 W mercury-vapor bulb have been used to nondiscriminately capture multiple Spodoptera spp. (Blair, 1974) and most assuredly other insects as well. A modified light trap using six 20-W fluorescent lights also proved effective for monitoring flight activity of S. littoralis (El-Mezayyen et al., 1997).

For additional survey information see: http://www.aphis.usda.gov/import_export/plants/manuals/emergency/downloads/nprg_s podoptera.pdf.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of S. littoralis is by morphological identification. S. littoralis is difficult to distinguish from S. litura without close examination of the genitalia. S. littoralis is also confused with S. dolichos, S. ornithogalli, S. latifascia and other Spodoptera species (present in the United States).

Literature-Based Methods: Observation of adult genitalia is often the only certain method to separate species. Screening aids to help identify S. littoralis in the field and by using wing diagnostics are available http://caps.ceris.purdue.edu/webfm_send/553 and http://caps.ceris.purdue.edu/webfm_send/554.

Easily Confused Pests

79 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

S. littoralis is often confused with S. litura, and the variability and similarity of the two species makes correct identification difficult; examination of adult genitalia is often the only certain method to separate the two species. For more information on morphological discrimination between the adult, pupal, and larval stages of the two species, refer to Schmutterer (1969), Cayrol (1972), Mochida (1973), and Brown and Dewhurst (1975). Although markings on larvae are variable, a bright-yellow stripe along the length of the dorsal surface is characteristic of S. litura. On dissection of the genitalia, the ductus and ostium Figure 5. Larva of S. exigua. Photo bursae are the same length in female S. courtesy of Oklahoma State littoralis, whereas they are different University. lengths in S. litura. The shape of the juxta in males in both species is very characteristic, and the ornamentation of the aedeagus vesica is also diagnostic. The genitalia must be removed, cleaned in alkali, and examined microscopically. S. litura is not established in the continental United States, but has been reported in Hawaii. Larvae of S. littoralis can be confused with S. exigua, the beet armyworm, (established in the United States) (Fig. 5), but S. littoralis larvae are light or dark brown, while S. exigua are brown or green. S. littoralis is also larger than S. exigua (Venette et al., 2003).

Adults of S. littoralis are almost nearly identical in appearance to S. ornithogalli (Fig. 6), the yellow striped armyworm, a common pest in the United States. The hind wings of female S. littoralis are darker than those of S. ornithogalli Figure 6. Adult of S. ornithogalli. (USDA, 1982). Photo courtesy of Mississippi Entomological Museum. http://mothphotographersgroup.mss tate.edu/Files/JV/JV50.7.shtml

References Abul-Nasr, S. and Naguib, M.A. 1968. The population density of larvae and pupae of Spodoptera littoralis (Boisd.) in clover fields in Egypt (Lepid.: Agrotidae). Bulletin De La Societe Entomologique D'Egypte 52: 297-312.

Abul-Nasr, S., El-Sherif, S.I., and Naguib, M.A. 1971. Relative efficiency of certain sampling methods for the assessment of the larval and pupal populations of the cotton leafworm Spodoptera littoralis (Boisd.) (Lepid.: Agrotidae) in clover fields. Journal of Applied Entomology/Zeitschrift für Angewandte Entomologie 69: 98-101.

80 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

Ahmad, T.R. 1988. Field studies on sex pheromone trapping of cotton leafworm Spodoptera littoralis (Boisd.) (Lep.: Noctuidae). Journal of Applied Entomology/Zeitschrift für Angewandte Entomologie 105: 212-215.

Anderson, P. and Alborn, H. 1999. Effects of oviposition behaviour and larval development of Spodoptera littoralis by herbivore-induced changes in cotton plants. Entomologia Experimentalis et. Applicata 92: 45-51.

Bishara, I. 1934. The cotton worm Prodenia litura F. in Egypt. Bulletin de la Société Entomologique d'Egypte 18: 223-404.

Blair, B.W. 1974. Identification of economically important Spodoptera larvae (Lepidoptera: Noctuidae). In: Scientific Note. Journal of the Entomological Society of Southern Africa 37: 195-196.

Brown, E.S. and Dewhurst, C.F., 1975. The genus Spodoptera (Lepidoptera, Noctuidae) in Africa and the Near East. Bulletin of Entomological Research 65(2): 221-262.

CABI. 2007. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Cayrol, R.A. 1972. Famille des Noctuidae. In: Balachowsky AS, ed. Entomologie appliquée à l'agriculture. Vol. 2. Paris, France: Masson 1411-1423.

El-Mezayyen, G.A., El-Dahan, A.A., Moawad, G.M., and Tadros, M.S. 1997. A modified light trap as a tool for insects survey in relation to the main weather factors. Egyptian Journal of Agricultural Research 75: 995-1005.

EPPO. 1997. Spodoptera littoralis and Spodoptera litura. In: Smith IM, McNamara DG, Scott PR, Holderness M, eds. Quarantine pests for Europe. 2nd edition. Wallingford, UK: CAB International, 518- 525.

Inserra, S. and Calabretta, C. 1985. Attack by noctuids: a recurring problem in greenhouse crops of the Ragusa coast. Tecnica Agricola 37(3-4): 283-297.

Kehat, M. and Dunkelblum, E. 1993. Sex pheromones: achievements in monitoring and mating disruption of cotton pests in Israel. Archives of Insect Biochemistry and Physiology 22(3-4): 425-431.

Miller, G.W. 1976. Cold storage as a quarantine treatment to prevent the introduction of Spodoptera littoralis (Boisd.) into glasshouses in the UK. Plant Pathology 25(4): 193-196.

Mochida, O. 1973. Two important insect pests, Spodoptera litura (F.) and S. littoralis (Boisd.) (Lepidoptera:Noctuidae), on various crops - morphological discrimination of the adult, pupal and larval stages. Applied Entomology and Zoology 8(4): 205-214.

Nucifora, A. 1985. Successive cultivation and systems of integrated control in protected crops of the Mediterranean area. Tecnica Agricola 37(3-4): 223-241.

Pinhey, E.C.G. 1975. Moths of Southern Africa. Descriptions and colour illustrations of 1183 species. Moths of Southern Africa.

PPQ. 1993. Fact sheet for exotic pest detection survey recommendations. Cooperative Agricultural Pest Survey (CAPS) and Plant Protection and Quarantine, US Department of Agriculture.

81 Spodoptera littoralis Primary Pest of Corn Arthropods Egyptian cotton leafworm Moth

Rizk, G.A., Soliman, M.A., and Ismael, H.M. 1990. Efficiency of sex pheromone and U. V. light traps attracting male moths of the cotton leafworm Spodoptera littoralis (Boisd.). Assiut Journal of Agricultural Sciences 21(3): 86-102.

Salama, H.S., Dimetry, N.Z., and Salem, S.A. 1970. On the host preference and biology of the cotton leaf worm Spodoptera littoralis. Zeitung für Angewandte Entomologie 67: 261-266.

Salem, S. and Salama, H.S. 1985. Sex pheromones for mass trapping of Spodoptera littoralis (Boisd.) in Egypt. Journal of Applied Entomology/Zeitschrift fur Angewandte Entomologie 100: 316-319.

Schmutterer, H. 1969. Pests of crops in Northeast and Central Africa with particular reference to the Sudan. Stuttgart, Germany: Gustav Fischer Verlag.

Sidibe, B. and Lauge, G. 1977. Effect of warm periods and of constant temperatures on some biological criteria in Spodoptera littoralis Boisduval (Lepidoptera Noctuidae). Annales de la Societe Entomologique de France 13(2): 369-379.

USDA. 1982. Pests not known to occur in the United States or of limited distribution, No. 25: Egyptian cottonworm., pp. 1-14. APHIS-PPQ, Hyattsville, MD.

Venette, R.C., Davis, E.E., Zaspel. J., Heisler, H., and Larson, M. 2003. Mini Risk Assessment Egyptian cotton leafworm, Spodoptera littoralis Boisduval [Lepidoptera: Noctuidae]. Cooperative Agricultural Pest Survey, Animal and Plant Health Inspection Service, US Department of Agriculture. Available on line at: http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/slittoralispra.pdf.

82 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

Spodoptera litura

Scientific Name Spodoptera litura Fabricius

Synonyms: Mamestra albisparsa, Noctua elata, Noctua histrionica, Noctua litura, Prodenia ciligera, Prodenia declinata, Prodenia evanescens, Prodenia glaucistriga, Prodenia litura, Prodenia subterminalis, Prodenia tasmanica, Prodenia testaceoides, Prodenia littoralis, and Spodoptera littoralis

Common Name(s) Rice cutworm, armyworm, taro caterpillar, tobacco budworm, cotton leafworm, cluster caterpillar, cotton worm, Egyptian cotton leafworm, tobacco caterpillar, tobacco cutworm, tobacco leaf caterpillar, and common cutworm

Type of Pest Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Family: Noctuidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2003 through 2012

Pest Description The two Old World cotton leafworm species, Spodoptera litura and S. littoralis, are allopatric, their ranges covering Asia and Africa, Europe and the Middle East, respectively. Many authors have regarded them as the same species, but they have been differentiated based on adult genitalia differences (Mochida, 1973; CABI, 2007). Figure 1. Egg mass with recently hatched larvae. Photo courtesy of http://australian- Eggs: Spherical, insects.com/lepidoptera/acro/litura.html somewhat flattened, sculpted with approximately 40 longitudinal ribs, 0.4 - 0.7 mm in diameter; pearly green,

83 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

turning black with time, laid in batches covered with pale orange-brown or pink hair-like scales from the females body (Pearson, 1958; CABI, 2007). The female scales look like an irregular furry mass on the underside of a leaf of a plant (Fig. 1).

Larva: Newly hatched larvae are tiny, blackish green (Fig. 1) with a distinct black band on the first abdominal segment. Fully grown larvae are stout and smooth with scattered short setae. Head is shiny black, and has conspicuous black tubercules each with a long hair on each segment. Color of fully grown larvae not constant, but varies Figure 2. S. litura larva. Photo courtesy from dark gray to dark brown, or black, of Lynn Finn. http://australian- sometimes marked with yellow dorsal and insects.com/lepidoptera/acro/litura.html. lateral stripes of unequal width. The lateral yellow stripe is bordered dorsally with series of semilunar black marks (Fig 2). Mature larvae are 40-50 mm with two large black spots on first and eighth abdominal segments (Hill, 1975; USDA, 1982; CABI, 2007). When disturbed, the larvae curl into a tight spiral with the head protected in the center.

Pupa: Reddish brown in color, enclosed inside rough earthen cases in the soil, 18-22 mm long, last abdominal segment terminates in two hooks (USDA, 1982; CABI, 2007).

Adult: Body whitish to yellowish, suffused with pale red. Forewings dark brown with lighter shaded lines and stripes (Fig. 3). Hind wings whitish with violet sheen, margin dark brown and venation brown. Thorax and abdomen orange to light brown with hair-like tufts on dorsal surface. Head clothed with tufts of light and dark brown scales. Body length 14- 18 mm, wing span 28-38 mm (Hill, 1975; USDA, 1982). Figure 3. Female (left) and male (right) S. litura. See Schmutterer (1969), Photos courtesy of K. Kiritani (JP). Cayrol (1972), and Brown and http://www.eppo.org/QUARANTINE/insects/Spodopte Dewhurst (1975) for additional ra_litura/PRODLI_images.htm. information.

84 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

Biology and Ecology The eggs of S. litura are laid in bunches of 50 to 300 on the under surface of leaves (preferred) by female moths (Chari and Patel, 1983). They hatch in 3 to 4 days. A single female lays 1500 to 2500 eggs in about 6 to 8 days. Castor bean is the most preferred host for ovipositing females (Chari and Patel, 1983). Newly irrigated fields are also very attractive to ovipositing females. Three peak periods of egg laying have been observed in the third weeks of June and July and in mid-August. Newly hatched larvae feed gregariously on the epidermis of the leaf. If the population density is high or the host is not suitable, the young larvae will hang on silken threads and migrate to other leaves or preferred hosts. There are generally six instars. The general habit of the larva is that the 1st, 2nd, and 3rd instars remain on the lower surface of leaves. The 4th, 5th, and 6th instars escape from sunshine, push to loosen the surface of the soil, and bite out soil particles to form a clay cell or cocoon in which to pupate (Chari and Patel, 1983).

Ahmed et al. (1979) showed that S. litura adults developed from first instar larvae in 23.4 days at 28°C. Mean female longevity was 8.3 days and mean fecundity was 2673 eggs. Mean male longevity was 10.4 days. No mating took place on the night of emergence and maximum mating response occurred on the second night after emergence (Yamanaka et al., 1975; Ahmed et al., 1979). According to Yamanaka et al. (1975) the female continues to lay eggs in egg masses over a period of 5 days at 25°C.

Maximum fecundity for S. litura was observed at 27°C (81°F) under 12 hours per 24 hours of light (100 foot candle light) (Hasmat and Khan, 1977, 1978). Temperatures between 24 and 30°C (75 to 86°F) were also favorable for fecundity and fertility. At 33 and 39°C (91 and 102°F), both fecundity and fertility were decreased, and in the latter, fertility was completely inhibited (Hasmat and Khan, 1977). Twenty four hours exposure to light markedly reduced both fecundity and fertility. Hatching was highest in dark conditions (Hashmat and Khan, 1978). Parasuraman and Jayaraj (1983a) noted that 25°C (77°F) and 75% relative humidity were favorable for development of S. litura with a shorter larval period, 100% pupation, a shortened pupal period, and 100% adult emergence.

Ranga Rao et al. (1989) reported that an average of 64 degree-days (DD) above a threshold of 8°C (46°F) was required for oviposition to egg hatch. The larval period required 303 DD, and the pupal stage 155 DD above a 10°C (50°F) threshold. Females needed 29 DD above a 10.8°C (51°F) threshold from emergence to oviposition. The upper developmental threshold temperature of all stages was 37°C (99°F); 40°C (104°F) was lethal.

Maheswara Reddy (1983) showed that the majority of mating occurred between 11:30 PM and 12:30 AM under controlled conditions. The duration of matings ranged between 82.5 and 90 minutes. Although males are capable of insemination throughout their lifecycle, no males inseminated more than one female in one night. Some males failed to inseminate even one female on some nights. The mean number of matings per male was 10.3 and per female was 3.1 (Ahmed et al., 1979). Ohbayashi et al. (1973) showed

85 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth two peaks in mating behavior at 11:00 PM (3 hours after initiation of a dark period) and a minor peak at 3:00 AM (1 hour before the end of the dark period).

S. litura spends its pre-pupal and pupal period inside soil. In India, Parasuraman and Jayaraj (1983b) found pupation was maximal under fallen leaves, especially in wet, sandy loam soil. Although the depth of pupation varied, no pupation was observed beyond 12 cm deep. Across soil types, most larvae pupated at a 4 cm depth.

Symptoms/Signs On most crops, damage arises from extensive feeding by larvae, leading to complete stripping of the plants. Larvae are leaf eaters but sometimes act as a cutworm with crop seedlings. S. litura feeds on the underside of leaves causing feeding scars and skeletonization of leaves. Early larval stages remain together radiating out from the egg mass. However, later stages are solitary. Initially there are numerous small feeding points, which eventually spread over the entire leaf. Because of this pest’s feeding activities, holes and bare sections are later found on leaves, young stalks, bolls, and buds. Larvae mine into young shoots. In certain cases, whole shoot tips wilt above a hole and eventually die (Hill, 1975; USDA, 1982).

Corn: The stems are often mined and young grains in the ear may be injured (CABI, 2007).

Other crops: On cotton, leaves are heavily attacked and bolls have large holes in them from which yellowish-green to dark-green larval excrement protrudes. In tobacco, leaves develop irregular, brownish-red patches and the stem base may be gnawed off.

Pest Importance S. litura larvae are polyphagous defoliators, seasonally common in annual and perennial agricultural systems in tropical and temperate Asia. This noctuid is often found as part of a complex of lepidopteran and non-lepidopteran foliar feeders but may also damage tubers and roots. Hosts include field crops grown for food and fiber, plantation and forestry crops, as well as certain weed species (CABI, 2007).

Most work on the economic impact of S. litura has been conducted in India, where it is a serious pest of a range of field crops. It has caused 12 to 23% loss to tomatoes in the monsoon season, and 9 to 24% loss in the winter (Patnaik, 1998). In a 40- to 45-day-old potato crop, damage ranged from 20 to 100% in different parts of the field depending on moisture availability. Larvae also attacked exposed tubers when young succulent leaves were unavailable (CABI, 2007). S. litura is also a pest of sugarbeet, with infestations commencing in March and peaking in late March and April (Chatterjee and Nayak, 1987). Severe infestations led to the skeletonization of leaves, as well as feeding holes in roots that rendered the crop 'virtually unfit for marketing'. Late harvested crops were most severely affected and, in extreme cases, 100% of the roots were damaged, leading to considerable yield reduction. Aroid tuber crops (including taro (Colocasia esculenta)) suffered yield losses of up to 29% as a result of infestation by S. litura, Aphis gossypii (cotton or melon ) and spider mites (Pillai et al., 1993).

86 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

S. litura causes damage to many species of forest and plantation trees and shrubs (Roychoudhury et al., 1995). It is responsible for brown flag syndrome in banana (Ranjith et al., 1997) and 5 to 10% fruit damage in grapes (Balikai et al., 1999).

S. litura is also a member of a complex that causes extensive defoliation of soybean (Bhattacharjee and Ghude, 1985). Defoliation as severe as 48.7% during the pre-bloom stage of growth caused no 'marked' difference from a control treatment in which defoliation was prevented by repeated insecticide application. Number and weight of pods and grains per plant were, however, reduced when defoliation occurred at, or after, blooming.

Insecticide resistance has been reported in India (Armes et al., 1997; Kranthi et al., 2001) and Pakistan (Ahmad et al., 2007).

Known Hosts Both S. litura and S. littoralis are widely polyphagous (Brown and Dewhurst, 1975; Holloway, 1989). The host range of S. litura covers at least 120 species (Venette et al., 2003). Among the main crop species attacked by S. litura in the tropics are taro, cotton, flax, peanuts, jute, alfalfa, corn, rice, soybeans, tea, tobacco, vegetables, aubergines (eggplant), Brassica spp., Capsicum spp., cucurbits, beans, potatoes, sweet potatoes, grape, and cowpea. Other hosts include ornamentals, wild plants, weeds and shade trees (for example, Leucaena leucocephala, a shade tree of cocoa plantations in Indonesia). Balasubramanian et al. (1984) showed better larval growth and higher adult fecundity when reared on castor bean compared to tomato, sweet potato, okra, cotton, sunflower, eggplant and alfalfa.

Major Hosts Abelmoschus esculentus (okra), Acacia mangium (brown salwood), Allium cepa (onion), Amaranthus (grain amaranth), Arachis hypogaea (peanut), Beta vulgaris var. saccharifera (sugarbeet), Boehmeria nivea (ramie), Brassica, Brassica oleracea var. botrytis (cauliflower), Brassica oleracea var. capitata (cabbage), Camellia sinensis (tea), Capsicum frutescens (chili), Castilla elastica (castilloa rubber), Cicer arietinum (chickpea), Citrus, Coffea (coffee), Colocasia esculenta (taro), Corchorus (jutes), Corchorus olitorius (jute), Coriandrum sativum (coriander), Crotalaria juncea (sunn hemp), Cynara scolymus (artichoke), Erythroxylum coca (coca), (leguminous plants), Foeniculum vulgare (fennel), Fragaria ananassa (strawberry), Gladiolus hybrids (gladiola), Glycine max (soybean), Gossypium spp.(cotton), Helianthus annuus (sunflower), Hevea brasiliensis (rubber), Ipomoea batatas (sweet potato), Jatropha curcas (Barbados nut), Lathyrus odoratus (sweet pea), Lilium spp. (lily), Linum usitatissimum (flax), , Malus domestica (apple), Manihot esculenta (cassava), Medicago sativa (alfalfa), Morus alba (mora), Musa spp. (banana), Nicotiana tabacum (tobacco), Oryza sativa (rice), Papaver (poppies), Paulownia tomentosa (paulownia), Phaseolus (beans), Piper nigrum (black pepper), Poaceae (grasses), Psophocarpus tetragonolobus (winged bean), Raphanus sativus (radish), Ricinus communis (castor bean), Rosa spp. (), Sesbania grandiflora (agati), Solanum esculentum (tomato), Solanum melongena (aubergine, eggplant), Solanum tuberosum (potato), Sorghum

87 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

bicolor (sorghum), Syzygium aromaticum (clove), Tectona grandis (teak), Theobroma cacao (cocoa), Trifolium spp. (clover), Trigonella foenum-graecum (fenugreek), Vigna mungo (black gram), Vigna radiata (mung bean), Vigna unguiculata (cowpea), Vitis vinifera (grape), Zea mays (corn), and Zinnia elegans (zinnia).

For a complete listing of hosts see Venette et al. (2003).

Known Vectors (or associated insects) S. litura is not a known vector and does not have any associated organisms.

Known Distribution The tobacco caterpillar, S. litura, is one of the most important insect pests of agricultural crops in the Asian tropics. This species is widely distributed throughout tropical and temperate Asia, Australasia and the Pacific Islands (Kranz et al., 1977).

Asia: Afghanistan, Bangladesh, Brunei Darussalam, Cambodia, China, Christmas Island, Cocos Islands, India, Indonesia, Iran, Japan, Korea, Laos, Lebanon, Malaysia, Maldives, Myanmar, Nepal, Oman, Pakistan, Philippines, Singapore, Sri Lanka, Syria, Thailand, and Vietnam. Europe: Russia. Africa: Reunion. North America: United States (Hawaii). Oceania: American Samoa, Australia, Belau, Cook Islands, Federated states of Micronesia, Fiji, French Polynesia, Guam, Kiribati, Marshall Islands, New Caledonia, New Zealand, Niue, Norfolk Island, Northern Mariana Islands, Papua New Guinea, Pitcairn Islands, Samoa, Solomon Islands, Tonga, Tuvalu, Midway Islands, Wake Island, Vanuatu, and the Wallis and Futuna Islands.

Potential Distribution within the United States The pest has been present in Hawaii since 1964 (CABI, 2007). S. litura was identified in a sample from a Miami-Dade County, Florida nursery in April 2007. Pheromone traps have been placed over a nine square mile area and have yielded no additional finds. A recent risk analysis by USDA-APHIS- PPQ-CPHST shows that portions of Alabama, Arkansas, Florida, Georgia, Louisiana, Mississippi, North Carolina, Oklahoma, South Carolina, and Texas are at the greatest risk from S. litura. Establishment of S. litura is unlikely in portions of Arizona, California, Colorado, Connecticut, Delaware, Idaho, Maine, Michigan, Minnesota, Montana (whole state), Nebraska, Nevada, New Mexico, New York, North Dakota, Oregon, Pennsylvania, Rhode Island, South Dakota, Utah, Vermont, Washington, West Virginia, Wisconsin, and Wyoming.

Survey CAPS-Approved Method: Trap with lure. A plastic bucket trap [unitrap] with dry kill strip is the approved trap for Spodoptera litura. The lure information is provided below:

Lure Dispenser Dispenser Lure Length of Compound Load Type Abbreviation Effectiveness a) 'Z,E,9,11- a) 1.76 mg laminate CL 12 weeks 14:AC b) 0.24 mg

88 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth b) 'Z,E,9,12- 14:AC

The plastic bucket trap (also known as the Universal moth trap or unitrap) should have a green canopy, yellow funnel, and white bucket and should be used with dry kill strip. For instructions on using the trap (see Appendix G “Plastic Bucket Trap Protocol”).

IMPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet). As of June 2010, S. litura and S. littoralis lures should be placed in different traps and separated by at least 20 meters (65 feet). Literature-Based Methods: Trapping: The identification of a male sex pheromone of S. litura, (Z,E)-(9,11)- tetradecadienyl acetate and (Z,E)-(9,12)-tetradecadienyl acetate by Tamaki (1973) has enabled effective monitoring of this species for several years. One milligram of a 10:1 mixture of these two compounds in a rubber septum attracted a comparable number of males as 10 caged virgin females in the field (Yushima et al., 1974). The compounds are most effective in a ratio (A:B) between 4:1 to 39:1 (Yushima et al., 1974). The two components in a ratio of 9:1 are available commercially as Litlure in Japan (Yushima et al., 1974). For early detection sampling, traps should be placed in open areas with short vegetation (Hirano, 1976). Krishnananda and Satyanarayana (1985) found that trap catches at 2.0 m above the ground level caught significantly more male S. litura than those placed at higher of lower heights (ranging from 0.5 m to 4.0 m). Ranga Rao et al. (1991) suggest trap placement at 1 m.

A standard sex pheromone trap (plastic dry funnel trap and pheromone septa) has been developed at the International Crop Research Institute for the Semi-Arid Tropics (ICRISAT) (Pawar et al., 1988; Ranga Rao et al., 1990, 1991; Singh and Sachan, 1993). Water traps baited with synthetic pheromone, box traps with rectangular windows, and cylindrical traps equipped with a blowing fan (to suck the males into a bag attached to bottom of the cylinder) have been used in Japan (Yushima et al., 1974; Hirano, 1976; Hirano, 1977; Oyama, 1977; Nakamura, 1977). Kamano et al. (1976) also mention a trap composed of two cylindrical parts and four cones made of wire screen that opened to the outside. Krishnanda and Satyanarayana (1985) used a dry trap that incorporated a tin sheet for the trap head, to which a polythene sleeve (45 x 10 cm) was attached. A small cylindrical polythene vial with 2.5 mg of pheromone was fastened to a small hook inside the dome. Rango Rao et al. (1991), however, found that at night many moths escaped from ‘sleeve’ traps and recommended either single or double funnel traps.

Visual survey: Visual survey can be used to determine the presence of S. litura. The presence of newly hatched larvae can be detected by the 'scratch' marks they make on the leaf surface. Particular attention should be given to leaves in the upper and middle

89 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

portion of the plants (Parasuraman, 1983). The older larvae are night-feeders, feeding primarily between midnight and 3:00 am and are usually found in the soil around the base of plants during the day. They chew large areas of the leaf, and can, at high population densities, strip a crop of its leaves. In such cases, larvae migrate in large groups from one field to another in search of food. S. litura may be detected any time the hosts are in an actively growing stage with foliage available, usually spring and fall. Check for 1st and 2nd instar larvae during the day on the undersurface of leaves and host plants. Watch for skeletonized foliage and perforated leaves. If no larvae are obvious, look in nearby hiding places. Third instar larvae rest in upper soil layers during the day. Sweep net for adults and larvae at dawn or dusk. Watch for external feeding damage to fruits. Watch near lights and light trap collections for adult specimens. Submit similar noctuid moths in any stage for identification (USDA, 1982).

Not recommended: Light traps have been used to monitor S. litura populations (Vaishampayan and Verma, 1983). Capture of S. litura moths was affected by the stage of the moon, with the traps being least effective during the full moon and most effective during the new moon (Parasuraman and Jayaraj, 1982).

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of S. litura is by morphological identification. It is difficult to distinguish S. litura from S. littoralis without close examination of the genitalia; consult appropriate keys by Todd and Poole (1980) and Pogue (2002). To separate from other noctuids, use the key developed by Todd and Poole (1980). S. litura can also be confused with S. dolichos, S. ornithogalli, S. pulcella and other Spodoptera species (present in the United States).

Literature-Based Methods: Wing coloration has been used to separate the sexes of S. litura (Singh et al., 1975). S. litura can be easily confused with S. littoralis. Adults are similar, and they can be distinguished only through examination of genitalia. On dissection of the genitalia, ductus and ostium bursae are the same length in female S. littoralis, different lengths in S. litura. The shape of the juxta in males is very characteristic, and the ornamentation of the aedeagus vesica is also diagnostic. The larvae of the two species are not easily separable, but some distinguishing criteria are used for the 6th instar. Mochida (1973) provides information on morphological discrimination between the adult, pupal and larval stages of the two species.

Screening aids to help identify S. litura in the field (http://caps.ceris.purdue.edu/webfm_send/555) and using wing diagnostics are available (http://caps.ceris.purdue.edu/webfm_send/556).

For additional images, including photos of host damage see http://www.padil.gov.au/viewPestDiagnosticImages.aspx?id=418.

Easily Confused Pests

90 Spodoptera litura Primary Pest of Corn Arthropods Rice cutworm Moth

Adult S. litura closely resemble Spodoptera ornithogali (yellowstriped armyworm), a pest in the United States. However, the hindwings of female S. litura are darker than those of S. ornithogalli.

References Ahmad, M., Iqbal Arif, M., and Ahmad, M. 2007. Occurrence of insecticide resistance in field populations of Spodoptera litura (Lepidoptera: Noctuidae) in Pakistan. Crop Protection 26: 809-817.

Ahmed, A.M., Etman, M., and Hooper, G.H.S. 1979. Developmental and reproductive biology of Spodoptera litura (F.) (Lepidoptera: Noctuidae). J. Aust. Ent. Soc. 18: 363-372.

Armes, N.J., Wightman, J.A., Jadhav, D.R., and Ranga Rao, G.V. 1997. Status of insecticide resistance in Spodoptera litura in Andhra Prades, India. Pesticide Science 50: 240-248.

Balasubramanian, G., Chellia, S., and Balasubramanian, M. 1984. Effect of host plants on the biology of Spodoptera litura Fabricius. Indian J. Agric. Soc. 54(12): 1075-1080.

Balikai, R.A., Bagali, A.N., and Ryagi, Y.H. 1999. Incidence of Spodoptera litura Fab. on grapevine, Vitis vinifera L. Insect Environment 5: 32.

Bhattacharjee, N.S. and Ghude, D.B. 1985. Effect of artificial and natural defoliation on the yield of soybean. Indian Journal of Agricultural Sciences 55: 427-429.

Brown, E.S. and Dewhurst, C.F. 1975. The genus Spodoptera (Lepidoptera, Noctuidae) in Africa and the Near East. Bulletin of Entomological Research 65: 221-262.

CABI. 2007. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Cayrol, R.A. 1972. Famille des Noctuidae. In: Balachowsky AS, ed. Entomologie appliquée à l'agriculture. Vol. 2. Paris, France: Masson 1411-1423.

Chari, M.S. and Patel, S.N. 1983. Cotton leaf worm Spodoptera litura Fabricius its biology and integrated control measures. Cotton Dev. 13(1): 7-8.

Chatterje, P.B. and Nayak, D.K. 1987. Occurrence of Spodoptera litura (Fabr.) as a new pest of sugar- beet in West Bengal. Pesticides 21: 21-22.

Hashmat, M. and Khan, M.A. 1977. The effect of temperature on the fecundity and fertility of Spodoptera litura (Fabr.) (Lepidoptera: Noctuidae). J. Anim. Morphol. Physiol. 24(2): 203-210.

Hashmat, M. and Khan, M.A. 1978. The fecundity and the fertility of Spodoptera litura (Fabr.) in relation to photoperiod. Z. Angew. Entomol. 85(2): 215-219.

Hill, D.S. 1975. Agricultural insect pests of the tropics and their control. Cambridge Univ. Press, London.

Hirano, C. 1976. Effect of trap location on catches of males of Spodoptera litura (Lepidoptera: Noctuidae) in pheromone baited traps. Appl. Entomol. Zool. 11(4): 335-339.

Hirano, C. 1977. Efficiency of water pan trap baited with synthetic sex pheromone in attracting male Spodoptera litura (Lepidoptera: Noctuidae). Jap. J. Appl. Entomol. Zool. 21(4): 216-219.

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Holloway, J.D. 1989. The moths of Borneo: family Noctuidae, trifine subfamilies: , Heliothinae, , Acronictinae, Amphipyrinae, Agaristinae. Malayan Nature Journal 42: 57-228.

Kamano, S., Wakamura, S., and Oyama, M. 1976. Applications of synthetic sex pheromone and its components for manipulating Spodoptera litura. Proceedings of the International Symposium on Insect Pheromones and their Applications. Pg. 135-144.

Kranthi, K.R., Jadhav, D.R., Wajari, R.R., Ali, S.S., and Russell, D. 2001. Carbamate and organophosphate resistance in cotton pests in India, 1995 to 1999. Bull. Ent. Res. 91: 37-46.

Kranz, J., Schumutterer, H., and Koch, W. eds. 1977. Diseases Pests and Weeds in Tropical Crops. Berlin and Hamburg, Germany: Verlag Paul Parley.

Krishnananda, N. and Satyanarayana, S.V.V. 1985. Effect of height of pheromone trap on the catch of Spodoptera litura moths in tobacco nurseries. Phytoparasitica 13(1): 59-62.

Maheswara Reddy, A. 1983. Observations on the mating behaviour of tobacco cut worm, Spodoptera litura (F.). Madras Agric. J. 70(2): 137-138.

Mochida, O. 1973. Two important insect pests, Spodoptera litura (F.) and S. littoralis (Boisd.) (Lepidoptera: Noctuidae), on various crops - morphological discrimination of the adult, pupal and larval stages. Applied Entomology and Zoology 8: 205-214.

Nakamura, K. 1977. The active space of the Spodoptera litura (F.) sex pheromone and the pheromone component determining this space. Appl. Entomol. Zool. 12(2): 162-177.

Ohbayashi, N., Yushima, T., Noguchi, H., and Tamaki, Y. 1973. Time of mating and sex pheromone production and release of Spodoptera litura (F.) (Lepidoptera: Noctuidae). Kontyu 41(4): 389-395.

Oyama, M. 1977. Pheromone trap equipped with a blowing electric fan for Spodoptera litura. Jap. J. Appl. Entomol. Zool. 21(2): 103-105.

Parasuraman, S. 1983. Larval behaviour of Spodoptera litura (F.) in cotton agro-ecosystem. Madras Agric. J. 70(2): 131-134.

Parasuraman, S. and Jayaraj, S. 1982. Studies on light trap catches of tobacco caterpillar Spodoptera litura (F.). Cotton Dev. 13(1): 15-16.

Parasuraman, S. and Jayaraj, S. 1983a. Effect of temperature and relative humidity on the development and adult longevity of the polyphagous Spodoptera litura (Fabr.) (Lepidoptera: Noctuidae). Indian J. Agric. Soc. 53(7): 582-584.

Parasuraman, S. and Jayaraj, S. 1983b.Pupation behaviour of Spodoptera litura (F). Madras Agric. J. 69(12): 830-831.

Patnaik, H.P. 1998. Pheromone trap catches of Spodoptera litura F. and extent of damage on hybrid tomato in Orissa. Advances in IPM for horticultural crops. Proceedings of the First National Symposium on Pest Management in Horticultural Crops: environmental implications and thrusts, Bangalore, India, 15- 17 October 1997, 68-72.

Pawar, C.S., Sithanantham, S., Bhatnagar, V.S., Srivastava, C.P., and Reed, W. 1988. The development of sex pheromone trapping of Heliothis armigera at ICRISAT, India. Trop. Pest Manage. 34: 39-43.

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Pearson, E.O. 1958. The insect pests of cotton in tropical Africa. Commonwealth Inst. Entomol., London.

Pillai, K.S., Palaniswami, M.S., Rajamma, P., Mohandas, C., and Jayaprakas, C.A. 1993. Pest management in tuber crops. Indian Horticulture 38: 20-23.

Pogue, M. G. 2002. A world revision of the genus Spodoptera guenee (Lepidoptera: Noctuidae). Mem. American Entomol. Soc. 43: 1-202.

Ranga Rao, G.V., Wightman, J.A., and Rango Rao, D.V. 1990. The development of a standard pheromone trapping procedure for Spodoptera litura (F.) (Lepidoptera: Noctuidae) population in groundnut (Arachis hyopogaea L) crops. Tropical Pest Management 37(1): 37-40.

Ranga Rao, G.V., Wightman, J.A., and Ranga Rao, D.V. 1989. Threshold temperatures and thermal requirements for the development of Spodoptera litura (Lepidoptera: Noctuidae). Environ. Entomol. 18(4): 548-551.

Ranga Rao, G.V., Wightman, J.A., and Ranga Rao, D.V. 1991. Monitoring Spodoptera litura (F.) (Lepidoptera: Noctuidae) using sex attractant traps: Effect of trap height and time of the night on moth catch. Insect. Sci. Applic. 12(4): 443-447.

Ranjith, A.M., Haseena, Bhaskar., and Nambiar, S.S. 1997. Spodoptera litura causes brown flag syndrome in banana. Insect Environment 3: 44.

Roychoudhury, N., Shamila, Kalia., and Joshi, K.C. 1995. Pest status and larval feeding preference of Spodoptera litura (Fabricius) Boursin (Lepidoptera: Noctuidae) on teak. Indian Forester 121: 581-583.

Schmutterer, H. 1969. Pests of crops in Northeast and Central Africa. Stuttgart, Germany: Gustav Fischer Verlag 186-188.

Singh, K.N. and Sachan, G.C. 1993. Spodoptera litura male moth catches in pheromone traps and their relationship with oviposition in groundnut field at Pantnagar, India. Insect. Sci. Applic. 14(1): 11-14.

Singh, O.P., Matkar, S.M., and Gangrade, G.M. 1975. Sex identification of Spodoptera (Prodenia) litura (Fabricius) by wing coloration. Curr. Sci. 44(18): 673.

Tamaki, Y. 1973. Sex pheromone of Spodoptera litura (F.) (Lepidoptera:Nocutidae): isolation, identification, and synthesis, Appl. Entomol. Zool. 8(3): 200-203.

Todd, E. L. and Poole, R.W. 1980. Keys and illustrations for the armyworm moths of the Noctuidae genus Spodoptera guenee from the Western Hemisphere. Ann. Entomol. Soc. Am. 73: 722-738.

USDA. 1982. Pests not known to occur in the United States or of limited distribution: Rice cutworm. USDA-APHIS-PPQ.

Vaishampayan, S.Y. and Verma, R. 1983. Comparative efficiency of various light sources in trapping adults of Heliothis armigera (Hubn.), Spodoptera litura (Bois.) and Agrotis ipsilon (Hufn.) (Lepidoptera: Noctuidae). Indian J. Agric. Sci. 53(3): 163-167.

Venette, R.C., Davis, E.E., Zaspel, J., Heisler, H., and Larson, M. 2003. Mini-risk assessment: Rice cutworm, Spodoptera litura Fabricius [Lepidoptera: Noctuidae].

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Yamanaka, H., Nakasuji, F., and Kiritani, K. 1975. Development of the tobacco cutworm Spodoptera litura in special reference to the density of larvae. Bull. Koch. Inst. Agric. For Sci. 7: 1-7. (English summary).

Yushima, T., Tamaki, Y., Kamano, S., Oyama. M. 1974. Field evaluation of synthetic sex pheromone 'Litlure' as an attractant for males of Spodoptera litura (F.) (Lepidoptera: Noctuidae). Applied Entomology and Zoology 9: 147-152.

94 Striacosta albicosta Primary Pest of Corn Arthropods Moth

Striacosta albicosta

Scientific name Striacosta albicosta (Smith)

Synonyms: Agrotis albicosta, Loxagrotis albicosta, Richia albicosta

Common names Western bean cutworm

Type of pest Moth

Taxonomic position Class: Insecta, Order: Lepidoptera, Family: Noctuidae

Reason for Inclusion in Manual Requested by CAPS Community - Native to Western United States, now spreading east (records now to Pennsylvania and Ontario, Canada), and of economic concern in corn and beans, but not soybeans.

Pest Description Eggs: Creamy white when first placed on the upper leaf surface, laid in clusters of 5-200, circular with ridges running from top to bottom (Fig.1, top). Maturing eggs turn more tan, then a Top: Newly laid purplish-grey color 1-2 days before egg hatch (Fig. Figure 1. eggs of S. albicosta. Bottom: 1, bottom). Eggs hatch in about 5-7 days. Mature eggs of S. albicosta.

Photos courtesy of Frank Larvae: There a 6-7 larval stages (instars); newly Peairs, Colorado State emerged larvae have a dull orange body, black University www.bugwood.org head, pronotum with a dark brown band on both and Mark Moore, sides (Fig. 2). On corn, these larvae will move up www.mooreagphotos.com, to the tassels (or directly to silks, if available) to respectively. begin feeding. Later instars move down to the ear, and have a tan or brown body with a broad and pale stripe on the dorsal side of their abdomen, two dark brown bands on the pronotum and no distinctive spots on the abdomen (Fig. 3). When fully mature, larvae are about 1.5 inches long. Unlike other cutworm pests, S. albicosta larvae are not

95 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

cannibalistic, so multiple larvae in one ear are likely. Mature larvae drop to the ground, burrow and overwinter in soil chambers.

Pupae: Orange-brown in color, and occur in the soil. Rarely seen.

Adults: Body length 0.75 inches long with a wingspan of 1.5 inches. Forewing with a white to cream-colored stripe along the outer, leading edge, with 2 markings (a comma- shaped mark and a circle marking) interior to this cream-colored strip (Fig. 4). Hindwings are cream colored without any pattern or markings (Fig. 4). Adults eclose (hatch) and begin to fly in mid to late June; this varies with latitude and climate. Adults are active at night.

Biology and Ecology Figure 2. Newly emerged first Striacosta albicosta is native to the Western instar larvae of S. albicosta. United States and has been extending its These are consuming part of their historical range to the east; it now has been egg shells before traveling up to found as far east as New York and Ontario, the tassels to begin feeding on Canada. S. albicosta appears to build to corn. Photo courtesy of Purdue higher population densities in sandier soils Extension Entomology. (Tooker, 2010). Populations of the moth in Ohio, Indiana, Michigan, and Ontario are building in areas bordering the Great Lakes. This Noctuid moth feeds on corn and beans, but not soybeans, and its preference is for corn.

Due to their long development time (about 60 days in a 27ºC (81ºF) laboratory environment), there is only one generation per year in the United States. Striacosta albicosta is a mid-to late-season pest, and has been recorded to cause as much as a 40% yield Figure 3. Later instars of S. loss in corn. Later maturing corn lines tend to albicosta. Note: The dark brown suffer more damage from S. albicosta than bands on the pronotum (segment shorter season hybrids (Holtzer, 1983; directly behind the head) and the Eichenseer et al., 2008). This pest pale stripes running lengthwise overwinters as larvae in the soil (within self- down the entire dorsal side of the constructed soil chambers), and become abdominal segments. Photo active when soil temperatures reach 10ºC courtesy of Purdue Extension (50ºF). Larvae then pupate and begin to Entomology. emerge from the soil as adults, starting in

96 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

June, with peak emergence depending on latitude and climate. Females lay eggs on the upper surface of leaves (normally on the top three or four leaves of the corn plant) just before the tassel stage (also called anthesis). After the tassel stage in corn, females shift to ovipositing on beans. Females lay an average of about 400 eggs in their lifetime.

Larvae hatch after 5-7 days, and move up the plant to feed on the tassel, including the pollen. Larvae are also mobile, and can move between plants with a 10-foot radius of their egg mass. After tassels have matured, larvae feed on shed pollen, leaf tissue, and silks, finally on the developing ear. This ear/kernel feeding can occur along the entire length of the ear, not just at the tip. Multiple larvae can be found within a single ear. Larvae DO NOT tunnel into stalks, but may tunnel throughout the ear.

After about two months of development (most spent within the ears), larvae drop to the ground in September, burrow into the ground, Figure 4. Adult S. albicosta. Top: The cream create a chamber and overwinter stripe along the outer edge of each forewing, below the frost line. and the circle and comma shape markings interior to the stripe. Bottom: Adult with wings Niche overlap with the European spread to show their non-descript hind wings. corn borer and corn earworm may Photos courtesy of Purdue Extension exist (Catangui and Berg, 2006, Entomology. Dorhout and Rice, 2010), and it has been speculated that the current range expansion is, at least in part, due to the widespread use of transgenic Cry 1Ab corn lines. Cry 1Ab controls other corn ear pests, but is susceptible to S. albicosta. The use of Cry 1Ab may eliminate the competition between S. albicosta and other corn ear pests.

Estimates indicate that a field infestation rate of one S. albicosta larva per ear can cause 3.7 bushels/acre in losses. In addition, secondary infection can decrease the quality of the yield.

97 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

Spraying to control larvae is recommended if there is an 8% infestation rate (presence of eggs or small larvae) in field corn, 4% in sweet corn for processing, and >1% infestation for fresh market sweet corn. Recommend insecticides include cyhalothrin- lambda (Matador 120 E) or deltamethrin (Decis 5.0 EC); local extension agents will have good pesticide recommendations for local laws and climates.

Note: Scout for spider mite colonies; some insecticides recommended for S. albicosta stimulate a secondary outbreak of spider mites; check label instructions and with local extension personnel for regional recommendations.

Biological control organisms specific to S. albicosta are not available. General predators (lacewings, lady beetles, spiders, etc.) can be significant feeders on larvae of S. albicosta, and a microsporidian (Nosema spp.) has been seen to affect larvae as well, but data quantifying the effect of these potential control agents has not been published.

Symptoms/Signs Because symptoms of this pest (damaged tassels, holes in the ear of corn) are common signs of other species moth larvae (corn earworm, European corn borer), traps that attract adults are the best way to identify this Figure 5. Late instar larvae pest. Harvested corn can have significant of S. albicosta feeding within damage from this pest, even with only one a maturing ear of corn. larvae per ear (Fig. 5). Photo courtesy of Purdue Extension Entomology.

Known Hosts Major hosts: Phaseolus vulgaris (bean) and Zea mays (corn).

Minor hosts: Phaseolus acutifolis, Phaseolus coccineus (tepary bean), Phaseolus lunatus (adzuki bean), Pisum sativium (garden pea), Vicia faba (lima bena), Vigna angularis (scarlet runner bean), and Vigna sinensis (horse bean) (Blickenstaff and Jolly, 1982).

Ground cherry, black nightshade, and tomato although previously reported as hosts were found to be unlikely hosts (Blickenstaff and Jolly, 1982). Older larvae can feed and complete development on these hosts but younger larvae cannot. Soybean was considered a poor host (Blickenstaff and Jolly, 1982).

98 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

Known Vectors (or associated organisms) Secondary damage from fungal pathogens has been seen with the western corn rootworm. Elevated levels of mycotoxins in corn harvests have been seen with S. albicosta, but this is not always the case (Catangui & Berg, 2006).

Known Distribution S. albicosta has been expanding its range eastward. As of July 2010, the pest has been found as far east as Long Island New York. The presence Figure 6. Pheromone trap for S. of the western bean cutworm has albicosta. Photo courtesy of Purdue been confirmed in Arizona, Extension Entomology. Colorado, Idaho, Illinois, Indiana, Iowa, Kansas, Michigan, Minnesota, Missouri, Nebraska, New Mexico, New York, North Dakota, Ohio, Oklahoma, Pennsylvania, South Dakota, Texas, Utah, Wisconsin, and Wyoming.

Potential Distribution within the United States This pest has the potential to occur wherever corn is grown in the contiguous United States.

Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: In Wisconsin, it is recommended that scouting for adults Figure 7. Adult S. albicosta, with arrows identifying distinctive begin when 1320 total marks: the small white circle, the comma shaped band and the degree days (DD) have cream-colored band together make identifying the moth in North accumulated, based on America very simple. No current moth has all three of these a base temperature of readily apparent marks. Photo courtesy of Purdue Extension Entomology.

99 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

10ºC (50ºF). Pheromone traps can indicate of the presence of this moth, and visual inspection of the top three or four leaves of corn after adult flight has been confirmed as important for the timing of insecticide spraying. In general, moth catches do not correlate well with larval populations. Several eastern states would like to assess larval populations (visual survey) to better ascertain the risk posed by this pest.

Trapping: Pheromone lures to monitor the activity and peak male adult populations are commercially available (ex., Scentry western bean cutworm pheromone lure, Great Lakes IPM, Vestaburg, MI and Suterra, Inc. Bend, OR). The chemical composition of the pheromone lure is not known at this time. Placement of traps between two cornfields collects more moths than traps placed between corn and soybeans (Dorhout and Rice, 2008). Figure 8. Fall armyworm adult. Photo courtesy Pheromone lures are of William Lambert, University of Georgia, commonly placed at the top of www.bugwood.org. modified gallon milk jugs (Fig. 6); traps are baited with the lure near the top with a 1:1 or 4:1 mix of water: antifreeze plus a drop of dish soap in the base. These are placed 1.5 meters (4 feet) above the ground near the edge of the corn field. Traps placed under 1.2 meters tend to catch significantly less moths, Traps are checked once a week, and the lures are changed once in the growing season. The Wisconsin protocol can be found at: http://www.entomology.wisc.ed u/cullenlab/extension/xtras/PD Fs/WBCWtrapping2008.pdf. Figure 9. Yellow striped armyworm adult. Photo Tooker and Fleischer (2010) courtesy of Natasha Wright, Florida Dept. of Ag., used tricolor (green, white, and www.bugwood.org. yellow) universal traps (unitraps or bucket traps). The universal traps contained insectidal strips as a killing agent. Traps were placed at the edge of the corn fields and were baited with synthetic female western bean cutworm pheromone. The closest traps were separated by approximately 4 km. Pheromone lures were changed every two weeks. The universal

100 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth traps, which produce a better specimen than milk jug traps, are simpler to use because they do not require antifreeze solution.

Visual Survey: Begin larval scouting when adult moths can be found flying, normally after May 1st and 1320 degree days have accumulated. Focus on the top 3 or 4 leaves for young larvae or egg masses. Inspect these leaves on 20 consecutive plants at five locations.

Visual surveys should target corn varieties susceptible to S. albicosta. Transgenic Cry1Ab corn is susceptible to attack by this pest; YieldGard and StarLink genes, as examples, are readily infested (StarLink was removed from the market in 2000), purportedly by eliminating the susceptible European corn borer (Catangui & Berg, 2006). Conventional corn (when in the presence of the European corn borer) does not seem to be as susceptible as the Cry1Ab corn lines. The Cry1 F expressed by some corn hybrids (examples Herculex I, Herculex Xtra) seems to be effective at avoiding damage by S. albicosta. Newer transgenic corn lines are also targeting this pest.

Sticky-wing traps and black light traps have been Figure 10. Dingy used, and can be as effective as the milk-jug traps if cutworm adult. Photo placed at the same 4-foot height. Regardless of trap courtesy of Heidi type used, take into consideration the prevailing night Genter, wind patterns, since S. albicosta moths are nocturnal. www.bugguide.net.

Key Diagnostics/Identification

CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Adults have a white band running the outside margin of each wing, along with a small white circle and comma-like mark on each wing (Fig. 7). Larvae also have two broad brown bands on the pronotum (on the first segment behind the head, See Fig. 3)

Easily Confused Pests Striacosta albicosta can be confused with other moths including: Spodoptera frugiperda, Spodoptera ornithogalli, and Feltia jaculifera. The fall armyworm, Spodoptera frugiperda (Fig. 8), the yellow striped armyworm, Spodoptera ornithogalli (Fig. 9) and the dingy cutworm, Feltia jaculifera (Fig. 10) adults have distinct markings, but they do not have the white margins, the circle and the comma-like markings that are distinctive to the Western bean cutworm (Fig. 7).

101 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

The dingy cutworm adult has the crescent-shape and the adults is similar in size, shape and distribution as the western corn rootworm, but normally is not the same color, nor does it have the circle shape and cream stripe on the outer margin of the forewing.

European corn borer (Ostrinia nubilalis) larvae can be differentiated by three main features; (1) western bean cutworm larvae do not have tubercles (spots) on their bodies, and European corn borer larvae do; (2) The western corn borer has two dark brown bands on their pronotum, whereas the European corn borer does not have anything like this on the pronotum, and (3) European corn borer larvae are without any distinctive stripes running lengthwise, which are present on western bean cutworm larvae (Figs. 3,11).

The fall armyworm (Spodoptera frugiperda) larvae may occur within corn at the same time as S. albicosta larvae, and have similar characteristics. Two important differences can be identified between the two larvae; (1) fall armyworm larvae have a distinct inverted ‘Y’ shaped suture on their head which is not pronounced on the western bean cutworm larvae, and (2) there are also four distinct ‘bumps’ on the most posterior segment of the fall armyworm larvae, which are also lacking on the western bean cutworm (Figs. 3 and 12 for reference).

The corn earworm larva, Helicoverpa zea (Fig. 12) has similar characteristics as the S. albicosta larva, but, again, with distinct differences. The pronotum bands on the S. albicosta larvae are not present on the corn earworm, and the skin of the corn earworm is rough; not so for the western bean cutworm. Figure 11. Top: Western bean cutworm larva.

Please also see Figure 3 for more detail. Bottom: Western bean cutworm eggs are European corn borer larva. Photos courtesy of laid on the upper surface of the Purdue Extension Entomology and Keith Weller, leaves close to the developing USDA-ARS, www.bugwood.org. ears; European corn borer

102 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

(second gen. adults) lay their eggs close to the developing ears, on the leaf collar, the underside of leaves and directly on the husk, not on the upper side of the top three or so leaves as S. albicosta. Western bean cutworm also has a peak moth flight about a month before second generation European corn borer adults.

Larval feeding damage can be confused with damage caused by the corn earworm and European corn borer. Larval identification is required to determine which species is responsible for the damage.

Figure 12. Top: Fall armyworm larva. Bottom: Corn earworm larva. Photos courtesy of Purdue Extension Entomology.

References Appel, L.L. 1993. Economic injury levels for western bean cutworm, Loxagrotis albicosta (Smith), with notes on biomics and culture of the cutworm. PhD. Dissertation, University of Idaho, Moscow.

103 Striacosta albicosta Primary Pest of Corn Arthropods Western bean cutworm Moth

Blickenstaff, C.C., and Jolly, P.M. 1982. Host plants of western bean cutworm. Environ. Entomol. 11(2): 421-425. Catangui, M.A. and Berg, R.K. 2006. Western bean cutworm, Striacosta albicosta (Smith) (Lepidoptera: Noctuidae), as a potential pest of transgenic Cry1Ab Bacillus thuringiensis corn hybrids in South Dakota. Environmental Entomology, 35(5):1439-1452.

Dorhout, D.L. and Rice, R.K. 2008. An evaluation of western bean cutworm pheromone trapping techniques (Lepidoptera: Noctuidae) in a corn and soybean agrosystem. Journal of Economic Entomology, 101(2): 404-408.

Dourhout, D.L., and Rice, M.E. 2010. Intraguild competition and enhanced survival of western bean cutworm (Lepidoptera: Noctuidae) on transgenic Cry1Ab (MON810) Bacillus thuringiensis corn. Journal of Economic Entomology, 103(1): 54-62.

Eichensee, H., Strohbehn, R., and Burks, J. 2008. Frequency and severity of western bean cutworm (Lepidoptera: Noctuidae) ear damage in transgenic corn hybrids expressing different Bacillus thuringiensis Cry toxins. Journal of Economic Entomology, 101(2): 555-563.

Holtzer, T.O. 1983. Distribution of western bean cutworm eggs among short-, mid-, and long-season hybrids planted on different dates. Environmental Entomology, 12:1375-1379.

Tooker, J.F. 2010. Personal communication.

Tooker, J.F., and Fleisher, S.J. 2010. First report of western bean cutwork (Striacosta albicosta) in Pennsylvania. Online: Crop Management DOI: 10.1094CM-2010-0616-01-RS.

104 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Thaumatotibia leucotreta

Scientific Name Thaumatotibia leucotreta Meyrick

Synonyms: leucotreta

Common Name(s) False codling moth, citrus codling moth, orange moth, and orange codling moth.

Type of Pest: Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description False codling moth (FCM), T. leucotreta, is a pest of economic importance to many crops throughout sub-Saharan Africa, South Africa and the islands of the Atlantic and Indian Oceans (Stibick, 2006). The FCM is an internal fruit feeding tortricid that does not undergo diapause and may be found throughout the year in warm climates on suitable host crops. Larval feeding and development can affect fruit development at any stage, causing premature ripening and fruit drop. T. leucotreta is a generalist with respect to host plant selection and has been recorded as feeding on over 50 different plant species. The generalist feeding strategy enables survival in marginal conditions as is necessary due to lack of diapause. Important host crops include avocado (Persea americana), citrus (Citrus spp.), corn (Zea mays), cotton (Gossypium spp.), macadamia (Macadamia spp.), and peach and plum (Prunus spp.) (USDA, 1984; Stibick, 2006).

Eggs: Eggs are flat, oval (0.77 mm long by 0.60 mm wide) shaped discs with a granulated surface. The eggs are white to Figure 1. Larvae of T. leucotreta. cream colored when initially laid. They Photo courtesy of T. Grove and W. change to a reddish color before the black Styn. www.bugwood.org. head capsule of the larvae becomes visible

105 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

under the chorion prior to eclosion (Daiber, 1979a).

Larvae: First instar (neonate) larvae approximately 1 to 1.2 mm in length with dark pinacula giving a spotted appearance, fifth instar larvae are orangey-pink, becoming more pale on sides and yellow in ventral region, 12 to 18 mm long, with a brown head capsule and first thoracic segment (Fig. 1). The last abdominal segment bears an anal comb with 2 to 7 spines. The mean head capsule width (mm) for the first through fifth instar larvae has been recorded as: 0.22, 0.37, 0.61, 0.94 and 1.37, respectively (Daiber, 1979b).

Pupae: Prepupa and pupa form inside a lightly woven silk and soil cocoon created by the fifth instar larvae on ground. Length is 8 to 10 mm and sexual determination through morphological differences on pupal case is possible (Daiber, 1979c)

Adult: Adult body length 6 to 8 mm, wingspan of female and male moth is 15 to 20 mm and 15 to 18 mm, respectively. The body is brown with a thorax with a posterior double crest. Forewing is a mixture of plumbeous, brown, black, and ferruginous markings, most conspicuous being blackish triangular pre-tornal marking and crescent-shaped marking above it, and minute white spot in discal area. Male is distinguished from female by its large, pale grayish genital tuft, large dense grayish white brush hindlegs, and its heavily tufted hind tibia (Gunn, 1921; Couilloud, 1988; CABI, 2007).

Biology and Ecology In South Africa, FCM has four to six non-discrete generations per year (Georgala, 1969; Stofberg, 1954). Females lay individual eggs (100-250 per female) on fruit or foliage (Catling and Aschenborn, 1974; Daiber, 1978). Females tend to oviposit on prematurely ripened fruit or wounded fruit when compared to healthy fruit at a normal state of development Figure 2. Adult false codling moth. Photo (Newton and Mastro, 1989). courtesy of CABI, 2007. Neonate larvae penetrate the fruit where larval development is completed. Mature larvae leave the fruit and spin cocoons near the soil or in bark crevices. Diapause or a resting stage has not been recorded.

Daiber (1980) showed that T. leucotreta adults live longest at 15°C (59°F) while most eggs were laid at 25°C (77°F). Egg laying at 20 and 25°C (68 and 77°F) increased rapidly soon after the first egg was laid but only gradually at 15°C to reach peak numbers some time after the initial egg lay. Very few eggs were laid at 10°C (50°F).

106 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Symptoms/Signs In general, the habit of internal feeding by FCM larvae displays few symptoms. Emerging larvae bore into the albedo and usually feed just below the fruit surface. Cannibalism among young larvae ensures that usually only one caterpillar matures in each fruit. When full-grown the larvae bore their way out of the fruit to seek a site for pupation, the rind around the point of infestation takes on a yellowish-brown color as the tissue decays and collapses. Larval feeding and development can affect fruit development at any stage, causing premature ripening and fruit drop.

Corn: Larvae damage corn by entering the ear from the husk through the silk channel (Stibick, 2006). Larvae can be found in the corn stem as well (Reed, 1974). On corn, T. leucotreta has been reported laying eggs on the husk of the ear.

Grape: Fresh larval penetration holes in grapes can be seen, but require careful inspection of the fruit. Sometimes a few granules of frass can be found around a fresh penetration hole or a mass of frass may be found around older penetration holes. Other times, however, frass is not visible. The area around the penetration hole can become sunken and brown as damaged tissue decays (Johnson, date not known).

Citrus: All stages of citrus fruit are vulnerable to attack. FCM larvae are capable of developing in hard green fruit before control measures can be started. Once a fruit is damaged, it becomes vulnerable to fungal organisms and scavengers. There is sometimes a scar visible on infested fruit (Stibick, 2006).

Cotton: FCM feeds mainly on large green bolls. The younger larvae feed almost entirely inside the boll wall itself, but the older larvae penetrate the inner septum and feed on the developing seeds and lint (Reed, 1974). Larval penetration of cotton bolls facilitates entry of other microorganisms that can rot and destroy the boll. The cultivars Edranol, Hass and Pinkerton were the most susceptible to attack by FCM (Stibick, 2006).

Macadamia: Larvae damage the nuts by feeding on the developing kernel after they pierce the husk and shell. Nuts reaching 14 to 19 mm diameter size are at the most risk because nutrient content is the greatest; concurrently, false codling moth reaches the adult stage by this point and is able to oviposit on the nuts (Stibick, 2006).

Avocado: Moths lay eggs superficially on the fruit of avocado. Larvae hatch, develop, and can enter through the skin. Larvae are unable to develop in avocado fruit. However, their entrance creates lesions that lessen the marketability of fruit. Lesions develop into a raised crater on the fruit surface, with an inconspicuous hole in the center where the larva has entered. Granular excreta can also be seen (Stibick, 2006).

Stone Fruit: All stages of stone fruits are vulnerable to attack. False codling moth larvae are capable of developing in hard green fruit before control measures can be started. Once a fruit is damaged, it becomes vulnerable to fungal organisms and scavengers. Larvae damage stone fruits as they burrow into the fruit at the stem end and begin to feed around the stone. Infestations can be identified by the brown spots and dark brown

107 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

frass (Daiber, 1976). Peaches become susceptible to damage about six weeks before harvest. Detecting infested peaches can be difficult if the fruit is still firm and abscission has not occurred; consequently, the danger of selling potentially infested fruit poses a serious threat to the peach industry (Daiber, 1976; USDA, 1984; Stibick, 2006).

Pest Importance In Africa, FCM is a major pest of citrus and cotton. In the Citrusdal Valley region of South Africa, FCM caused from 2.9 to 15.2% crop loss on citrus depending on the farm and the pest control program (Schwartz and Anderson, 1983). In Ugandan cotton, boll rotting is a major cause of crop loss. Over 90% of rotten bolls had insect attack symptoms and at least 60% of those were caused by T. leucotreta (Reed, 1974). Approximately 44% of corn cobs examined contained larvae of T. leucotreta in Uganda (Reed, 1974).

According to CDFA (2008), commonly grown agricultural hosts in California for FCM include citrus, grapes, peach, plum, cherry, beans, tomato, pepper, persimmon, apricot, olive, pomegranate, English walnut, and corn. Based on its status as a pest in Africa, establishment of FCM in California and/or in other parts of the United States could result in significant economic losses. FCM would likely be a significant production and quarantine issue for numerous agricultural commodities. In California alone, the annual combined gross value of the top ten agricultural commodities which would be directly impacted by this pest is over $7.1 billion, which amounts to 22% of the total agricultural value for the State (USDA NASS, 2007).

Hoffmeyr and Pringle (1998) report resistance in FCM to the chitin synthesis inhibitor trifluron, commonly used for FCM control.

Known Hosts Major Hosts: Abelmoschus esculentus (okra), Abutilon hybridum (flowering maple), Abutilon × hybridum (Chinese lantern), Ananas comosus (pineapple), Averrhoa carambola (carambola), Camellia sinensis (tea), Capsicum spp. (peppers), Citrus spp., Coffea arabica (coffee), Gossypium spp. (cotton), Litchi chinensis (litchi), Macadamia spp. (macadamia), Mangifera indica (mango), Olea spp. (olive), Persea americana (avocado), Prunus armeniaca (apricot), Prunus domestica (plum), Prunus persica (peach), Prunus spp. (cherry), Psidium guajava (guava), Punica granatum (pomegranate), Quercus spp. (oak, acorns), Ricinus communis (castor bean), Sorghum bicolor (sorghum), and Zea mays (corn).

Minor/Wild hosts: Abutilon spp. (Indian mallow), Acacia nilotica (acacia), Acacia tortilis (umbrella thorn), Annona cherimola (cherimoya), Annona glabra (pond apple), Annona muricata (soursop), Annona reticulata (Bullock's heart, custard apple), Annona squamosa (sugar apple), Azanza garckeana (snot apple), galpinii (red bauhinia), Bequaertiodendron magalismontanum (stamvrug), Butyrospermum parkii (shea tree), Caesalpinia pulcherrima (pride-of-Barbados), Caesalpinia spp. (nicker), Calotropis

108 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

procera (sodom apple), Capparis tomentosa (African caper), Carya illinoensis (pecan), petersiana (monkey pod), Catha edulis (khat), Ceiba pentandra (kapok), Chrysophyllum cainito (star apple), Chrysophyllum palismontatum (stamvrugte), Cola nitida (bitter cola), Combretum apiculatum (apiculatum), Combretum apiculatum (rooibos), Combretum zeyheri (raasblaar), Cyphomandra betacea (tree tomato), Diospyros mespiliformis (Jjakkalsbessie), Diospyros spp. (persimmon), Englerophytum magalismontanum, Eriobotrya japonica (loquat), Eugenia uniflora (Surinam-cherry), Ficus capensis (wild fig), Flacourtia indica (governor's-plum), Garcinia mangostana (mangosteen), Harpephyllum caffrum (kaffir-plum), Hibiscus cannabinus (kenaf), Hibiscus spp. (hibiscus), Juglans regia (English walnut), Juglans spp. (walnut), Solanum (Lycopersicon) esculentum (tomato), Mimusops zeyheri (Transvaal red milkwood), Musa paradisiaca (banana), Pennisetum purpureum (elephant grass), Phaseolus lunatus (lima bean), Phaseolus spp. (bean), Physalis ixocarpa (tomatillo) Physalis spp. (groundcherry), Piper spp. (pepper), Podocarpus falcatus (yellowwood), Podocarpus spp. (plum pine), Pseudolachnostylis maprouneifolia (kudu-berry), Royena pallens (pale-branched Royena), Saccharum officinarum (sugarcane), Schotia spp. (boerboon), Sclerocarya birrea (marula) Sechium edule (chayote), Sida spp. (fanpetals), Solanum melongena (eggplant), Synsepalum dulcificum (miraculous berry), Syzygium cordatum (waterbessie), Syzygium jambos (rose-apple), Theobroma cacao (cacao),Triumfetta spp. (bur weed), Vangueria infausta (wild medlar) Vigna spp. (cowpea), Vitis spp. (grape), Xeroderris stuhlmannii (wing bean), Ximenia caffra (suurpruim), Yucca spp. (yucca), and Ziziphus spp. (jujube).

Known Vectors (or associated organisms) T. leucotreta is not a known vector and does not have any associated organisms. The wounds produced by T. leucotreta, however, can provide an entrance for pathogens and can damage host plants under humid conditions.

Known Distribution False codling moth is indigenous to Southern Africa and the Ethiopian region. It also occurs on the islands of Madagascar, Mauritius, Reunion, and St. Helena.

Africa: Angola, Benin, Burkina Faso, Burundi, Cameroon, Cape Verde, Central African Republic, Chad, Congo, Dahomey, Eritrea, Ethiopia, Gambia, Ghana, Ivory Coast, Kenya, Madagascar, Malawi, Mali, Mauritius, Mozambique, Niger, Nigeria, Nyasaland, Réunion, Rhodesia, Rwanda, Saint Helena, Senegal, Sierra Leone, Somalia, South Africa, Sudan, Swaziland, Tanganyika,Tanzania, Togo, Uganda, Upper Volta, Zaire, Zambia, Zanzibar, and Zimbabwe.

False codling moth has occasionally been found in Europe, where it was imported with produce from Africa (Bradley et al., 1979; Karvonen, 1983). Border inspections have intercepted false codling moth in Denmark, Finland, Netherlands and United Kingdom; the countries have remained free of the pest.

109 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Potential Distribution within the United States Infestation by FCM generally causes the fruit to drop before harvest. Larval entries, however, can take a few days to become visible. Those that occur near fruit harvest, therefore, are often not detected by the packing house fruit graders and infested fruit can be inadvertently packaged for export.

Increased international trade and tourism between the United States and many African countries in recent years has increased the risk of introduction of this pest. Since 1984, FCM has been intercepted over 1500 times on 99 plant taxa at 34 U.S. ports of entry. In June 2005, live FCM caterpillars were found at California’s border stations inside previously cold treated Clementine citrus from South Africa. Its discovery in California is a new record for the Americas. FCM is not known to be established in California.

On June 16, 2005, California Department of Food and Agriculture (CDFA) inspectors found 1 live and 1 dead larva on a shipment of South African clementines at the California border station in Needles. The larvae were identified by both a CDFA lab and the USDA Systematic Entomology Laboratory (SEL) Specialist as False Codling Moth (FCM), Thaumatotibia leucotreta, Meyrick. The fruit had entered the United States in the port of Philadelphia (PA) off the vessel Nova Zembla. Initial review of the cold treatment records did not reveal failures in the treatment. On June 20, a second live larva was intercepted on a separate shipment of South African clementines in California. This shipment came on the vessel Fuji Star on June 14, 2005. This larva was identified by CDFA as FCM. An eradication program would be triggered if two moths were detected within one life cycle and within three miles of each other, or a mated female was found, or any immature stage (egg, larva, or pupa) was found. FCM has not triggered an eradication project in California at this time. Survey using traps and some fruit sampling is continuing around the Ventura County find.

Survey Surveys should be focused in areas of high risk. A recent risk analysis by USDA- APHIS-PPQ-CPHST shows that portions of Alabama, Arkansas, Florida, Georgia, Illinois, Indiana, Kentucky, Louisiana, Mississippi, Missouri, North Carolina, Ohio, Oklahoma, South Carolina, Tennessee, Texas, Virginia, and West Virginia are at the greatest risk from T. leucotreta. Most areas of the continental United States are at moderate to high risk from T. leucotreta based on climate and host range.

CAPS-Approved Method: Trap with lure. A wing trap is the approved trap for T. leucotreta. The lure information is provided below:

Lure Compound Dispenser Load Dispenser Type Lure Abbreviation a) 'E,8-12:AC a) 0.9 mg gray rubber septum FCM b) 'Z,8-12:AC b) 0.1 mg

IMPORTANT: Placing lures for two or more target species in a trap, should never be done unless otherwise noted here.

110 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters.

Literature-Based Methods: For early detection surveys in corn, fields in close proximity to high risk areas such as citrus and stone fruit should be monitored utilizing pheromone traps. The pheromone traps should be placed at a frequency of 1 trap per 4 hectares and traps should be no closer than 150 to 200 m to each other. Traps should be inspected weekly. Corn should also be inspected visually for the presence of FCM during the growing season. The first four rows bordering citrus or stone fruit orchards should be examined carefully.

Trapping: Male T. leucotreta are attracted to a two component blend of (E)-8-dodecenyl acetate and (Z)-8-dodecenyl acetate. These components are most effective when used in a ratio between 70:30 and 30:70 (E:Z) (Persoons et al., 1977; Venette et al., 2003). More recently, Newton et al. (1993) refined the sex pheromone and reported that a 90:10 ratio was optimal. Stibick (2006) recommends utilizing a 50:50 ratio. The lure is available from the CPHST- Otis lab. Burger et al. (1990) showed that 7-vinyldecyl acetate, a by-product of the synthesis of one of the constituents of the pheromone blend, effectively disrupts the attraction of the male moths to virgin females or to synthetic lures.

A loading rate between 0.5 and 1.0 mg per septum was found to attract the greatest number of males. The pheromone blend (1 mg applied to a rubber septum) has been used effectively with Pherocon 1C traps to capture male T. leucotreta (Newton et al., 1993). Delta traps have also been used, but these have performed less well than either the Hoechst Biotrap or Pherocon 1C traps. Traps using closed polyethylene vials to dispense pheromones captured more moths than traps using rubber septa (using a 50:50 blend of (E)- and (Z)-8-dodecenyl acetate). Lures should be replaced every 8 weeks. Traps should be placed approximately 5 ft (1.5m) high. Hofmeyr and Burger (1995) developed a prototype controlled release dispenser that was capable of releasing sex pheromone without replacement for more than seven months. Pheromone traps (homemade sticky trap with unspecified pheromone blend) have been used to monitor the number of T. leucotreta adult males in citrus orchards (Daiber, 1978) and detect the presence of the pest in peach orchards (Daiber, 1981).

Pheromone lures with (E)- and (Z)-8-dodecenyl acetate may also attract Cydia cupressana (native), Hyperstrotia spp., Cydia atlantica (exotic), Cydia phaulomorpha (exotic) and Cryptophlebia peltastica (exotic).

Visual survey: Visual inspections of plant materials may be used to detect eggs, larvae, and adults of T. leucotreta (USDA, 1984). Look for plants showing signs of poor growth or rot; holes in fruit, nuts or bolls; adults hidden in foliage; and crawling larvae. Surveys are best conducted during warm, wet weather when the population of the pest increases (USDA, 1984). Eggs will commonly be found on fruits, foliage, and occasionally on branches (USDA, 1984). However, eggs are small and laid singly, which makes them

111 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth difficult to detect. On corn, T. leucotreta has been reported laying eggs on the husk of the ear.

Fruit should be inspected for spots, mold, or shrunken areas with 1 mm exit holes in the center. On citrus fruits and other fleshy hosts, dissections are needed to detect larvae; larvae are likely to be found in the pulp (USDA, 1984). Infested fruits may be on or off the tree. In cotton, older larvae may be found in open bolls and cotton seed (USDA, 1984). Occasionally adults may be observed on the trunk and leaves of trees in infested orchards (USDA, 1984). For field crops, such as corn, the whole plant is the recommended sample unit. Because larvae of T. leucotreta have a strongly aggregated spatial distribution among corn plants, a large sample size (>60 plants) is recommended; however at low densities of the pest (<1 larva/plant) sample sizes needed to detect the pest may be prohibitively large.

Soil Sampling: Collect soil samples within 200 yards of any larval or egg detection and at any spot where dropped, especially prematurely dropped, fruit occur. Soil samples should consist of loose surface soil and any debris. Examine soil for larvae, cocoons and pupae.

Not recommended: Robinson black light traps are ineffective at attracting adult T. leucotreta (Begemann and Schoeman, 1999). Therefore, black light traps should not be used. The effectiveness of black light traps may be improved if used in conjunction with pheromone lures (Möhr, 1973). Möhr (1973) speculates that pheromone may provide a long-distant attractant, but that attraction to black light becomes much stronger when moths are in close proximity to light traps.

Key Diagnostics CAPS-Approved Method: Confirmation of T. leucotreta is by morphological identification. Larval specimens must be examined under a dissecting microscope preferably by a screener experienced with the arrangement of setae on Lepidoptera larvae.

Literature-Based Methods: Thaumatotibia leucotreta can be distinguished from other species by host range and morphological characteristics. A tool for identifying larvae of leafrollers and a job aid is provided in Appendix D and Appendix E, respectively of the New Pest Response Guideline to False Codling Moth (available at http://www.aphis.usda.gov/import_export/plants/manuals/emergency/downloads/nprg- fcm.pdf) that can help you determine if you have a possible larva of false codling moth. The job aid from Appendix E is also available at http://caps.ceris.purdue.edu/webfm_send/544. Stofberg (1948) provides a detailed description of larval structures that distinguish FCM from other larvae.

See Padil website for additional FCM images, including diagnostic characters (http://www.padil.gov.au/viewPestDiagnosticImages.aspx?id=314).

Easily Confused Pests

112 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Thaumatotibia leucotreta can be confused with many Cydia spp. including C. pomonella (codling moth) because of similar appearance and damage, however, unlike codling moth, its host range does not include apples, pears or quince (USDA, 1984). In West Africa, T. leucotreta is often found in conjunction with Mussidia nigrevenella (pyralid moth), but they can be distinguished by close examination of morphological characters (CABI, 2007). In South Africa, there is also an overlapping host range for T. leucotreta, T. batrachopa (macadamia nut borer) and Cryptophelbia peltastica (litchi moth), particularly on litchi and macadamia (Venette et al., 2003; Stibick, 2006). The male litchi moth can be distinguished from similar species by a subtriangular or Y-shaped T8 with a pair of tufts of filiform scales from membranous pockets on each side (Stibick, 2006).

Male T. leucotreta can be distinguished from other tortricid species by its specialized hindwing, which is slightly reduced and has a circular pocket of fine hairlike black scales overlaid with broad weakly shining whitish scales in anal angle, and its heavily tufted hind tibia (Bradley et al., 1979).

The larvae are creamy white in color and can be confused with fruit fly larvae in some cases. The distinct brown black head of T. leucotreta larvae, however, make it readily distinguishable from fruit fly larvae (Economides, 1979).

References Begemann, G. and Schoeman, A.1999. The phenology of Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae), Tortrix capsensana (Walker) and Cryptophlebia leucotreta (Meyrick) (Lepidoptera: Tortricidae) on citrus at ebediela, South Africa. African Entomology 7: 131-148.

Bradley, J.D., Tremewan, W.G., and Smith, A. 1979. British tortricid moths. Tortricidae: . Vol. 2. London, England: British Museum (Natural History).

Burger, B.V., le Roux, M., Mackenroth, W.M., Spies, H.S.C., and Hofmeyr, J.H. 1990. 7-vinyl acetate, novel inhibitor of pheromonal attraction in the false codling moth, Cryptophlebia leucotreta. Tetrahedron Letters 31(40): 5771-5772.

CABI. 2007. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Catling, H.D. and Aschenborn, H. 1974. Population studies of the false coding moth, Cryptophlebia leucotreta Meyr., on citrus in the Transvaal. Phytophylactica 6: 31-38.

CDFA (California Department of Food and Agriculture). 2008. False Codling Moth Pest Profile.

Couilloud, R. 1988. Cryptophlebia (Argyoploce) leucotreta (Meyrick). Lepidoptera, Tortricidae, Olethreutinae. Coton et Fibres Tropicales 43: 319-351.

Daiber, C.C. 1976. A survey of false codling moth (Cryptophlebia leucotreta Meyr.) in peach orchards. Phytophylactica 8(4): 97-102.

Daiber, C. 1978. A survey of male flight of the false codling moth, Cryptophliebia leucotreta Meyr., by the use of the synthetic sex pheromone. Phytophylactica 10: 65-72.

Daiber, C.C. 1979a. A study of the biology of the false codling moth [Cryptophlebia leucotreta (Meyr.)]: the egg. Phytophylactica 11: 129-132.

113 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Daiber, C.C. 1979b. A study of the biology of the false codling moth [Cryptophlebia leucotreta (Meyr.)]: the larvae. Phytophylactica 11: 141-144.

Daiber, C.C. 1979c. A study of the biology of the false codling moth [Cryptophlebia leucotreta (Meyr.)]: the cocoon. Phytophylactica 11: 151-157.

Daiber, C.C. 1980. A study of the biology of the false codling moth [Cryptophlebia leucotreta (Meyr.)]: the adult and generations during the year. Phytophylactica 12: 187-193.

Daiber, C. 1981. False codling moth, Cryptophlebia leucotreta (Meyr.) in peach orchards and home gardens of the summer rainfall area of South Africa. Phytophylactica 13: 105-107.

Economides, C.V. 1979. False codling moth, a serious pest of citrus in Zambia. Farming in Zambia 12(2): 4.

Georgala, M.B. 1969. Control of false codling moth and fruit flies in citrus orchards S. Afr. Citrus J. 421: 3, 5, 7.

Gunn, D. 1921. The false-codling moth (Argyroploce leucotreta Meyr.). Science Bulletin No. 21. Pretoria, South Africa: The Government Printing and Stationary office.

Hofmeyr, J.H. and Pringle, K.L.1998. Resistance of false codling moth, Cryptophlebia leucotreta (Meyrick) (Lepidoptera: Tortricidae), to the chitin synthesis inhibitor, triflumuron. Afr. Entomol. 6: 373-375.

Hofmeyr, J.H. and Burger, B.V. 1995. Controlled-release pheromone dispenser for use in traps to monitor flight activity of false codling moth. Journal of Chemical Ecology 21(3): 355-363.

Johnson, S. Date unknown. False codling moth on table grapes. http://www.satgi.co.za/export/sites/sati/galleries/protected/other/FCM_on_Table_Grapes.pdf.

Karvonen, J. 1983. Cryptophlebia leucotreta imported into Finland (Lepidoptera, Tortricidae). Notulae Entomologicae 63: 94.

Möhr, J.D. 1973. Light trap studies with the false codling moth. Citrus and Sub-tropical Fruit Journal 20- 22.

Newton, P.J. and V. Mastro. 1989. Field evaluations of commercially available traps and synthetic sex pheromone lures of the false codling moth, Cryptophlebia leucotreta (Meyr.) (Lepidoptera: Tortricidae). Tropical Pest Management 35: 100-104.

Newton, P. J., Thomas, C.D., Mastro, V.C., and Schwalbe, C.P. 1993. Improved two component blend of the synthetic female sex pheromone of Cryptophlebia eucotreta, and identification of an attractant for C. peltastica. Entomologia Experimentalis et Applicata 66: 75-82.

Persoons, C.J., Ritter, F.J., and Nooijen, W.J. 1977. Sex pheromone of the false codling moth, Cryptophlebia leucotreta (Lepidoptera:Tortricidae): Evidence for a two component system. Journal Chemical Ecology 3: 717-722.

Reed, W. 1974. The false codling moth, Cryptophlebia leucotreta Meyr. (Lepidoptera: Olethreutidae) as a pest of cotton in Uganda. Cotton Grow. Rev. 51(3): 213-225.

Schwartz, A. and Anderson, T. 1983. Economic importance of false codling moth on navels in the Citrusdal Valley. Inligtingsbull Navorsingsinst Sitrus Subtop. Vrugte 130: 14-15.

114 Thaumatotibia leucotreta Primary Pest of Corn Arthropods False codling moth Moth

Stibick, J. 2006. New Pest Response Guidelines: False Codling Moth Thaumatotibia leucotreta. USDA– APHIS–PPQ–Emergency and Domestic Programs, Riverdale, Maryland http://www.aphis.usda.gov/import_export/plants/manuals/emergency/downloads/nprg_false_codling_mot h.pdf.

Sofberg, F.J. 1948. Larvae structures as a basis for identification of false codling moth (Argyroploce leucotreta, Meyr.) larvae. Journal of Entomol. Soc. Of South Africa XI: 68-75.

Stofberg, F.J. 1954. False codling moth of citrus. Farm S. Afr. 29: 273-276, 294.

USDA. 1984. Pests not known to occur in the United States or of limited distribution, No. 48: False codling moth. USDA- APHIS-PPQ.

USDA NASS. 2007. California Agricultural Statistics, 2006 Crop Year. USDA, National Agricultural Statistics Service, California Field Office, Sacramento, California. 92 pp. http://www.nass.usda.gov/Statistics_by_State/California/Publications/California_Ag_Statistics/2006cas- all.pdf.

Venette, R.C., Davis, E.E., DaCosta, M., Heisler, H., and Larson, M. 2003. Mini Risk Assessment - False codling moth, Thaumatotibia (=Cryptophlebia) leucotreta (Meyrick)[Lepidoptera: Tortricidae]. Cooperative Agricultural Pest Survey, Animal and Plant Health Inspection Service, US Department of Agriculture. http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/tleucotretapra.pdf.

115 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

Secondary Pests of Corn (Truncated Pest Datasheet)

Copitarsia spp.

Scientific Names of Species of Concern Copitarsia incommoda Walker Copitarsia decolora Herrich-Schaffer

Most of this pest data sheet is at the genus level due to taxonomic confusion (see note below); however, detailed pest descriptions are given for the two most economically important pest species, C. incommoda and C. decolora.

Note: Systematics and nomenclature within the genus Copitarsia are particularly problematic. Over time, the genus has included from six to twenty two species, depending on which taxonomic authority is consulted (Angulo and Olivares, 2003; Venette and Gould, 2006; Pogue and Simmons, 2008).

Synonyms: C. decolora: Agrotis heydenreichii, Mamestra decolora, Polia turbata, Copitarsia turbata, Mamestra inducta, Copitarsia inducta, Spaelotis subsignata, Copitarsia subsignata, Agrois hostilis, Copitarsia hostilis, Graphiphora sobria, Copitarsia sobria, Lycophotia margaritella, and Copitarsia margaritella.

C. incommoda: Agrotis consueta, Copitarsia consueta, Agrotis incommoda, Agrotis peruviana, Copitarsia peruviana, and Allorhodecia hampsoni.

Common Names Owlet moths, cutworms, army worms, leaf worms

Type of Pest Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Family: Noctuidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List - 2009

Pest Description Two species, Copitarsia incommoda and Copitarsia decolora, are the most economically important members of the genus. C. incommoda is reported from Mexico to northern Chile. Documented hosts of C. incommoda include asparagus, , and alfalfa. C. decolora is widely distributed in Central America and South America and has been reported from Mexico to Chile and east to Argentina. C. decolora feeds on a

116 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

variety of crops, including artichokes, cut flowers, lettuce, peas, beets, cabbage, carrots, corn, beans, and potatoes. C. decolora is routinely intercepted on produce at U.S. ports of entry. This species has historically been misidentified as C. incommoda in both agricultural and taxonomic literature.

General (all species in genus): Copitarsia spp. begin life as eggs (Fig. 1A), deposited singly or in egg masses. A single female may produce between 570 and 1640 eggs, depending on the quality of the environment and the host. Larvae complete five to six instars during development and reach a length of approximately 2 to 4 cm. The larvae tend to be green in color (Fig. 1B), but green, black, and gray phases occur that vary with habitat and crops attacked. Development time from egg to adult depends on many factors including temperature, humidity, and host. Reported larval development times vary from approximately 43 days at 24.5°C (76°F) on lettuce to 82.5 days at 20.4°C (69°F) on artificial diet (Arce de Hamity and Neder de Roman, 1992; Lopez-Avila, 1996; Velasquez, 1998).

A B

C D E

Figure 1. Copitarsia life stages. Eggs (A), larva (B), adult (C, D), and pupae (E). Photos courtesy of Julie Gould and Charles Olsen (USDA-APHIS-PPQ).

117 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

Copitarsia spp. pupate in the soil (Fig. 1E) and emerge as gray or brown moths (Fig. 1C, D) that are difficult to distinguish from other noctuids. Diapause has not been reported for any member of the genus. The literature suggests that Copitarsia spp. are multivoltine through much of their range. In general, Copitarsia spp. appear to have two to four generations per year.

Copitarsia decolora: (from Simmons and Pogue, 2004) Description. Medium-sized, light brown or gray moths with well-defined orbicular and reniform spots.

Discussion. C. decolora varies slightly in coloration from light to medium brown. Females tend to be larger and have darker hindwings than males. Mitochondrial DNA evidence indicates at least two morphologically cryptic species within C. decolora: one ranging from southern Mexico to Ecuador, the other occurring in Ecuador, Mexico, Colombia, and Peru (Simmons and Scheffer, 2004). The second species was recently named as Copitarsia corruda by Pogue and Simmons (2008). Larval host genera include: Asparagus (Liliaceae), Iris (Iridaceae), Ammi (Apiaceae), Lysimachia (Primulaceae), Callistephus (), and Aster (Asteraceae).

Diagnosis. C. decolora lacks the brush-like androconia found in male C. incommoda. Male C. decolora have a blunt digitus and corona of spines on the valve. Female C. decolora are recognizable due to the speculate, heavily sclerotized antevaginal plate.

Male. Head. Brown; antenna light brown, biserrate and ciliated; palpus light brown, apex white.

Thorax. Patagium brownish gray; mesothorax pale brown; metathorax gray to white; fore, mid, and hindleg mixed with white and brown scales, tibial spurs striped with brown; tarsi white.

Wings. Forewing. Length = 13 to 18 mm (average = 16.1 mm, SD = 1.3 mm, n= 14). Ground color light brown or gray; antemedial and postmedial lines, double row of brown zigzag lines with white between them; basal area with well-defined brown lines; brown reniform spot outlined in white; orbicular spot ground color with white inner and black outer margin; outer margin with triangular black spots between wing veins; fringe grayish- brown.

Hindwing. Ground color white; wide marginal band brown; veins toward wing margin brown; fringe brown basally, remainder white.

Abdomen. First three abdominal segments light gray, remainder of abdomen gray; genital tuft gray; sclerotized patches present in pleural membrane near second abdominal segment; hair brushes, scent pouches, and modified S2

118 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

absent; terminal tergite weakly sclerotized medially, more heavily sclerotized laterally, forming two circular areas.

Genitalia. Tegumen rounded; uncus apically swollen, bearing long setae; saccus extended into narrow point; valve sinuate, tapering to pointed apex; corona present; ampulla attenuate, apex extending beyond costal margin of valve; digitus spatulate; juxta a broad plate with pointed lateral margins, medio-ventral plate with rounded, sinnuate margins, dorsal margin V-shaped with a pair of ventrally produced arms with dorsally curved apices; spinose pad present above aedeagus; apex of aedeagus with a small sclerotized plate (sp) consisting of one large and two pointed projections, a large serrate sclerotized plate (lp) opposite small plate; vesica elongate; cornuti various sized elongate spines in both clusters and solitary in a spiral line in basal one-quarter of vesica.

Female. As in male, except antennae filiform and ciliated; forewing length = 14 to 18 mm (average = 16.8 mm, SD = 1.2 mm, n = 24); hindwing darker than males.

Genitalia. Papillae anales, posterior apophyses unmodified; anterior apophyses reduced in length, thickened; S8 unmodified; antevaginal plate U-shaped, spiculate texture, symmetrical; ductus bursae sclerotized, spinose; corpus bursae deeply ridged, spherical, three lines of signa; appendix bursae larger than corpus bursae, membranous, irregular in shape; ductus seminalis from posterior of appendix bursae.

Copitarsia incommoda: (From Simmons and Pogue, 2004)

Description. Medium-sized, pale brown moths, with well-defined orbicular and reniform spots and light brown hindwings.

Discussion. C. incommoda varies slightly in coloration from lighter to medium brown. Females tend to be larger and have darker hindwings than males.

Diagnosis. C. incommoda is often confused with C. decolora. Males of C. incommoda can be identified externally by their brush-like androconia on the second abdominal segment (sometimes only after dissection), which are absent in C. decolora. Male C. incommoda has a rounded digitus, and valves lack a corona of spines that is present in C. decolora. Female C. incommoda can be identified by the smooth texture of the U- shaped antevaginal plate, compared with the spiculate antevaginal plate found in C. decolora.

Male. Head. Brown; antenna pale brown, filiform and ciliated; palpus brown.

Thorax. Patagium brown; mesothorax lighter, tawny brown; metathorax cream to white; fore, mid, and hindleg mixed white and brown, tibial spurs striped with brown, tarsi white.

119 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

Wings. Forewing. Length = 14 to 18 mm (average = 16 mm, SD = 1.3 mm, n = 15). Ground color light brown; antemedial and postmedial lines, a double row of brown zigzag lines with white between them; basal area with well- defined brown lines; reniform spot ground color with white inner and black outer margin; orbicular spot ground color outlined in black; outer margin with triangular black spots between wing veins; fringe brown.

Hindwing. Ground color brown mixed with white scales basally; fringe light brown basally, rest white.

Abdomen. Brown, genital tuft white; hair brushes, scent pouches, and modified S2 present; terminal tergite as in C. decolora.

Genitalia. As in C. decolora, except corona absent; digitus slender, apex round, not spatulate; apex of aedeagus with a small sclerotized plate (sp) consisting of one large, one small, and three minute pointed projections; a series of variously sized, heavily sclerotized spines opposite small plate (ss); cornuti in a similar pattern to that of C. decolora, but more robust.

Female. As in male, except forewing length = 14 to 19 mm (average = 17.2 mm, SD = 1.3 mm, n = 18); hindwing darker than males.

Genitalia. As in C. decolora, except lateral lobes of U-shaped antevaginal plate larger than C. decolora.

Symptoms/Signs Eggs and larvae may be present on plant parts. Larvae generally feed externally on leaves, stems, and fruits of host plants but will occasionally bore into thicker non-woody tissues (Venette and Gould, 2006).

Survey CAPS-Approved Method: Blacklight trapping.

Literature-Based Methods: Surveys should occur at locations most at risk for Copitarsia spp. A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that large portions of California and the southeastern United States have the greatest risk for Copitarsia spp. establishment based on host availability and climate. Establishment is precluded in many areas of the western United States.

Trapping: A pheromone consisting of (Z)-9-tetradecenyl acetate (Z9-14:Ac) and Z-9- tetradecenol (Z9-14:0H) has been previously identified for C. decolora (Rojas et al., 2006). Captures in traps baited with a mixture of Z9-14:Ac and Z9-14:0H at 4:1, 10:1, and 100:1 ratios were not significantly different from traps baited with virgin females.

120 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

The commercial availability of this pheromone, however, is unknown at this time. The same components were identified for C. incommoda (Cibrian-Tovar et al., 2003).

Early detection surveys have traditionally utilized non-selective black light trapping.

Visual survey: Survey for Copitarsia spp. generally has been conducted visually at the ports by examining cut flowers and vegetable products destined for entry into the United States. All products are examined for the presence of egg masses and/or larvae.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of Copitarsia spp. is by morphological identification. Adults can be identified through genitalia dissections. Larvae can only be identified to genus.

Literature-Based Methods: Adult Copitarsia spp. have few external characteristics to distinguish them from other noctuid moths and can only be identified with confidence by genitalia dissections. At times, members of Copitarsia have been confused with the genera Agrotis, Euxoa, Polia, and Orthosia (Venette and Gould, 2006).

Copitarsia larvae can be distinguished from other genera based on external characteristics. For example, Copitarsia larvae have dark bars at the base of the two medial setae, white dorsal setae, misaligned head setae (dorsal ventrally), and two dark triangles on the posterior abdominal segments (Riley, 1998). Within the Copitarsia genus, adults can be identified by the presence of large spines on the foretarsi; however, larval and egg identification characters are inconsistent or nonexistent (Simmons and Pogue, 2004). Angulo and Olivares (2005), however, state that C. incommoda and C. decolora larvae can be distinguished by examining the spinneret and pinnaculae. Pogue and Simmons (2008) give characters to separate C. decolora from C. corruda, a newly described species of Copitarsia formerly classified as a cryptic species within C. decolora (Simmons and Scheffer, 2004).

References Angulo, A.O. and Olivares, T.S. 2003. Taxonomic update of the species of Copitarsia Hampson 1906, (Lepidoptera: Noctuidae: Cucilliinae). Gayana 67(1): 33-38.

Angulo, A.O. and Olivares, T.S. 2005. Two larval characters to separate Copitarsia incommoda (Walker) from C. decolora (Guenee) (Lepidoptera: Noctuidae). Gayanna 69(2): 409-410.

Arce de Hamity, M.G. and Neder de Roman, L.E. 1992. Aspectos bioecologicos de Copitarsia turbata (Herrich-Schaffer) (Lepidoptera: Noctuidae) imporantes en la determinacion del dano economico en culitvos de Lactuca sativa L. de la Quebrada de Humahuaca, Jujuy, Argentine. Review of the Society of Entomology of Argentina 50: 73-87.

Cibrian-Tover, J., Rojas, J., Putnam, S., Calyecac, G., Cruz-Lopez, L., Malo, E., and Diaz-Gomez, O. 2003. Identification of sexual pheromone of Copitarsia incommoda (Lepidoptera: Noctuidae) (Abstr.). Annual Meeting of the Entomological Society of America.

121 Copitarsia spp. Secondary Pest of Corn Arthropods Owlet moths Moth

Lopez-Avila, A. 1996. Insectos plagas del cultivo de la papa en Colombia y su manejo. In: Anon. (Ed.), Papas Colombianas con el major entorno ambiental, pp. 146-148, 150-154. Communicaciones y Asociados Ltda, Santafe de Bogota, Colombia.

Pogue, M.G. and Simmons, R.B. 2008. A new pest species of Copitarsia (Lepidoptera: Noctuidae) from the neotropical region feeding on asparagus and cut flowers. Ann. Entomol. Soc. Am. 101(4): 743-762.

Riley, D.R. 1998. Identification key for Copitarsia, Spodoptera exigua, and Peridorma saucia, US Department of Agriculture, Animal and Plant Health Inspection Service (Internal Report). Pharr, TX.

Rojas, J.C., Cruz-Lopez, L., Malo, E.A., Diaz-Gomez, O., Calyecac, G., and Cibrian Tovar, J. 2006. Identification of the sex pheromone of Copitarsia decolora (Lepidoptera: Noctuidae). Journal of Economic Entomology 99(3): 797-802.

Simmons, R.B. and Pogue, M.G. 2004. Redescription of two often-confused Noctuid pests, Copitarsia decolora and Copitarsia incommoda (Lepidoptera: Noctuidae: Cucullinae). Annual Entomology Society of America 97(6) 1159-1164.

Simmons. R.B. and Scheffer, S.J. 2004. Evidence of cryptic species within the pest Copitarsia turbata (Herrich-Shaffer) (Lepidoptera: Noctuidae). Annals of Entomology Society of America 97(4): 675-680.

Velasquez, Z.L.D. 1988. Ciclo biologico de Copitarsia turbata (Lep: Nocituidae) sobre cebolla, en Arequipa. Rev. Peru Entomology 30:108-110.

Venette, R.C. and Gould, J. 2006. A pest risk assessment for Copitarsia spp., Insects associated with importation of commodities into the United States. Euphytica 148: 165-183.

122 Darna pallivitta Secondary Pest of Corn Arthropods Nettle caterpillar Moth

Darna pallivitta

Scientific name Darna pallivitta Moore

Synonyms Miresa pallivitta and Oxyplax ochracea

Common names Nettle caterpillar and stinging Caterpillar

Type of pest Moth

Taxonomic position Class: Insecta, Order: Lepidoptera, Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description The nettle caterpillar’s stinging, spiny hairs Figure 1. Darna pallivitta eggs on the have a physical effect on human skin much underside of a leaf; in a row (top) and in a like fiberglass. In addition, the spines cluster (bottom). These eggs are small release a mixture of histamines produced by (0.8 mm) and nearly translucent, and are a poison gland, causing further irritation easily overlooked. Photos courtesy of (burning and itching) that might require Walter T. Nagamine, Hawaii Department of medical attention. If spines get into the Agriculture. eyes, the irritation can be acute; seek medical attention quickly.

Eggs: Deposited in small clusters, a line, or singly, usually on the undersides of older leaves. Eggs are flattened, transparent ovals, 1/32 inch (0.8 mm) wide and 1/16 inch (1.6 mm) long, appearing as a glassy sheen on the leaf surface that can easily be overlooked (Fig. 1) (Chun et al., 2005).

Larvae: The larvae (Fig. 2) can be up to 1 inch (25 mm) long and are covered with many rows of stinging spines. Larvae vary from white to light gray, with a dark longitudinal stripe down the back. Larvae have two rows of moderately long spiny tubercles which, like the body, are marbled with black and white on light gray. There is a dorsal white band and sometimes yellow or light orange flecks on the flanks anteriorly. The head is yellow. The larva leaves a viscous trail over the leaves as it progresses (Chun et al., 2005).

123 Darna pallivitta Secondary Pest of Corn Arthropods Nettle caterpillar Moth

Pupa: Pupation occurs within a light brown to lilac cocoon covered with coarse silk (Fig. 3).

Adult: The adult moth is approximately ½ inch (12.7 mm) long, with females usually larger than males. The forewing is divided by a white diagonal marking, with the upper portion rust-colored and the lower portion lighter brown; the hind wings are uniform light brown. These nocturnal moths have not been observed feeding. During the day they are inactive and retreat into vegetation, usually in an upside-down, perching position. Male and female moths are very similar, except in their antennae; females have filiform antennae, whereas males have bipectinate antennae (Fig. 4) (Chun et al., 2005).

Symptoms/Signs Significant leaf damage and defoliation can occur (Fig. 5). When these larvae are present, a distinctive black or green frass can also be seen on this vegetation (Figs. 5, 6). Figure 2. Early (top) and late instar (bottom) larva of D pallivitta. Photo Survey courtesy of Walter T. Nagamine, Hawaii CAPS-Approved Method: Has not been Department of Agriculture. evaluated at this time.

Literature-Based Methods: Darna pallivitta is a tropical pest that can cause significant damage to a wide range of agricultural, landscape and endemic vegetation. To date, it has been detected on three Hawaiian Islands, and been intercepted as larvae and cocoons several times in California.

Darna pallivitta would probably only be able to establish in southern California, Florida, Puerto Rico and the U. S. Virgin Islands; potentially southern Texas as well (Koop, Figure 3. Newly emerged D. pallivitta 2006). Jang et al. (2009) reports that while adult from cocoon (cocoon at right, about precipitation does not seem to be a factor in the size of a bean). Photo courtesy of their range expansion in Hawaii, there is Walter T. Nagamine, Hawaii Department evidence to suggest higher elevations and of Agriculture.

124 Darna pallivitta Secondary Pest of Corn Arthropods Nettle caterpillar Moth lower temperatures are potential limitations to the spread of D. pallivitta.

Trapping: Siderhurst et al. (2007) identified two pheromone components, n-butyl (E)-7,9- decadienoate (E7,9-10:COOn-Bu) and ethyl (E)-7,9-decadienoate, for D. pallivitta. The n-butyl ester pheromone component attracted male Figure 4. Adult male (left) and female moths at equal or greater rates than (right) D. pallivitta. Notice the bipectinate trapped virgin females. The ethyl antennae on the male, and the filiform ester, however, did not increase trap antennae on the female. Photo courtesy of captures at the levels and ratios Walter T. Nagamine, Hawaii Department of tested. Agriculture.

Jackson traps, delta traps with a sticky surface of approximately 130 cm2 (Fig. 7) baited with 250 µg of E7,9-10:COOn-Bu have been used to monitor D. pallivitta to in Hawaii (Jang et al., 2009). Traps suspended at 1 m trapped more males than traps at 3 and 5 meters.

Visual survey: Visual inspection for the larvae and their frass can also be used as a survey tool (Figs. 5, 6).

Surveys should occur at locations most at risk for D. pallivita. A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that portions of Florida, Illinois, Indiana, Iowa, Kansas, Louisiana, Minnesota, Mississippi, Nebraska, North Dakota, Ohio, South Dakota, and have the greatest risk for D. Figure 5. Young D. pallivitta larvae consume guinea pallivita establishment based on grass (Panicum maximum). The bottom image is a host availability alone. close-up of the top image. Notice the green balls of frass on the upper right portion of the bottom photo. Photo courtesy of Walter T. Nagamine, Hawaii Department of Agriculture.

125 Darna pallivitta Secondary Pest of Corn Arthropods Nettle caterpillar Moth

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Adult: The bronzy brown forewings traversed by an oblique white fascia are distinctive (Holloway, 2006). The valve of the male genitalia lacks a costal process.

Larva: The distinctive stinging spines and 4 orange spots on the dorsal side of its body in later instars are unlike moth larvae of the United States. Its frass can also be distinctive in color and shape.

Some of the caterpillars from the Nyphalidae family also are covered in spines, though non-stinging. Butterflies in the family include Kamehmeha butterfly (Vanessa tameame), painted lady (Vanessa cardui), red admiral (Vanessa atalanta), American lady (Vanessa virginiensis), and California tortoiseshell ( californica). The Kamehmeha butterfly is native to Hawaii. The painted lady occurs everywhere, including the United States, with the exception of South America, Australia, and the Arctic. Both the American lady and California tortoiseshell are present in the United States, although the California tortoiseshell primarily occurs in the western United States.

Figure 7. A Jackson trap with male D. pallivitta adults. The pheromone impregnated lure is seen in the middle. Photo courtesy of Hawaii Department of Agriculture. Figure 6. Young D. pallivitta larva with black frass. Photo courtesy of Walter T. Nagamine, Hawaii Department of Agriculture.

126 Darna pallivitta Secondary Pest of Corn Arthropods Nettle caterpillar Moth

References Chun, S., Hara A., Niino-DuPonte R., Nagamine W., Conant P., and Hirayama, C. 2005. Stinging nettle caterpillar, Darna pallivitta: pest alert. Cooperative Extension Service, University of Hawaii at Manoa, Manoa, Hawaii, U.S.A. http://www.ctahr.hawaii.edu/oc/freepubs/pdf/IP-22.pdf.

Holloway, J. D. 2006. The Moths of Borneo. http://www.mothsofborneo.com/part- 1/limacodidae/limacodidae-42-7.php.

Jang, E.B Siderhurst, M.S., Connant, P., and Siderhurst, L.A. 2009. Phenology and population radiation of the nettle caterpillar, Darna pallivitta (Moore) (Lepidoptera: Limacodidae) in Hawai’i. Chemoecology 19: 7-12.

Koop, A.L. 2006. New Pest Advisory Group Report. Darna pallivitta Moore: Nettle Caterpillar.

Siderhurst, M.S., Jang, E.B., Hara, A.H., and Conant, P. 2007. n-Butyl (E)-7,9-decadienoate: sex pheromone component of the nettle caterpillar, Darna pallivitta. Entomologia Experimentalis et Applicata 125: 63-69.

127 Eutetranychus orientalis Secondary Pest of Corn Arthropods Citrus brown mite Mite

Eutetranychus orientalis

Scientific Name Eutetranychus orientalis Klein

Synonyms: Anychus orientalis, Anychus ricini, Eutetranychus monodi, Eutetranychus sudanicaus, Eutetranychus anneckei, Anychus latus, and Eutetranychus latus

Common Name(s) Citrus brown mite, oriental mite, oriental red mite, oriental spider mite, and Lowveld citrus mite

Type of Pest Mite

Taxonomic Position Class: Arachnida, Order: Acarina, Family: Tetranychidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description The genus Eutetranychus is characterized by its empodium, which is reduced to a small protuberance (Avidov and Harper, 1969). The life cycle of E. orientalis is completed in four active (larva, protonymph, deutonymph, and adult) and three quiescent stages (nymphochrysalis, deutochrysalis, and teleochrysalis) (Lal, 1977).

Eggs: The eggs of E. orientalis are oval or circular (Fig. 1) and flattened, coming to a point dorsally, but lacking the long dorsal stalk of other spider mites. Newly laid, the eggs are bright and hyaline, but later they take on a yellow, parchment-like color (Smith-Meyer, 1981). Diameter of the eggs is 0.14 mm (Avidov and Figure 1. Eggs (left) and adult (right ) of E. orientalis. Harper, 1969). Photos courtesy of Pedro Torrent Chocarro.

Larvae: Average size of the larva of E. orientalis is 190 x 120 µm. The abdomen of female larvae and nymphs is greenish brown, while the abdomen of male larvae is reddish brown. The protonymph is pale-brown to light-green, with legs shorter than the

128 Eutetranychus orientalis Secondary Pest of Corn Arthropods Citrus brown mite Mite

body, average size 240 x 140 µm. The deutonymph is pale-brown to light-green, average size 300 x 220 µm.

Adults: Adult female E. orientalis are broad, oval and flattened. They vary in color from pale brown through brownish-green to dark green with darker spots within the body. The legs are about as long as the body and are yellow-brown (Fig. 1). Average size is 410 x 280 µm. Females have the dorsal striae of the prodosoma more or less parallel and slightly but distinctly lobed. The dorsal setae of the body are set on small tubercles, and the lateral setae of the body are moderately slender and spatulate.

Technical description: Empodia lacking on all tarsi; true claws slender, padlike, each with pair of tenent hairs; duplex setae of tarsi loosely associated, not paired as in other spider mites; 2 pairs of anal setae; 3 pairs of dorsal propodosomal setae, and 10 pairs of dorsal hysterosomal setae, all setae stout, serrate; dorsal striae of hysterosoma form V-pattern between setae D1 and E1, and setal bases E1 and F1 form a square; setal cout (solenidia or sensory rodlike setae in parentheses) of legs (Meyer, 1974). L coxa 2- 1-1-1, trochanter 1-1-1-1, femur (8-6-3/4-1/2), genu (5-5-2-2), tibia 9(1/4)-6(0/2)-6(0/1)- 7, and tarsus 15(3)-13(1/2)-10(1)-10(1).

Adult male E. orientalis are much smaller than the females. They are elongate and triangular in shape with long legs (leg about 1.5 x body length). Usually males have a higher solenidia count.

Short setae are found on legs and body of both sexes at all stages. The body setae are short, however, and cannot be seen with a 10x lens (Smith-Meyer, 1981; Dhooria and Butani, 1984).

The outstanding characteristic in the adult is that the legs are equal to, or longer than, the body length (Avidov and Harper, 1969).

Symptoms/Signs E. orientalis begins feeding on the upper side of the leaf along the midrib and then spreads to the lateral veins, causing the leaves to become chlorotic. Pale yellow streaks develop along the midrib and veins (Fig. 2) initially, which later progress to a grayish or Figure 2. Eutetranychus feeding damage on silvery appearance of the Ptychosperma palm. Photos courtesy of leaves. At times, the leaves http://www.pestalert.org/viewArchPestAlert.cfm?rid=62 appear to be covered in a

129 Eutetranychus orientalis Secondary Pest of Corn Arthropods Citrus brown mite Mite

layer of fine dust. When damaged, the younger, tender leaves show margins that are twisted upwards. Usually, little webbing is produced but can occur. In heavier infestations, the mites feed and oviposit over the whole upper surface of the leaf. Very heavy infestations on citrus cause leaf fall and die-back of branches, which may result in defoliated trees. Lower populations in dry areas can produce the same effect.

Survey CAPS-Approved Method: Visual survey is the method to survey for E. orientalis.

Literature-Based Methods: Visual survey: The presence of E. orientalis can be detected by discoloration of the host leaves and pale-yellow streaks along the midribs and veins. Eggs, immature stages, and adults may be observed visually on the upper leaf surface. Adult females are larger than the males. They are oval and flattened and are often pale brown through brownish- green to dark green. Webbing is possible (often dust colored), providing protection for the eggs. The spread of the mite is windborne, and new infestations commonly occur at the field perimeters. Field perimeters should, therefore, be scouted, especially field perimeters facing prevailing winds. Studies indicate that alfalfa plays a role in dispersing tetranychid mites to other crops (Osman, 1976). Fields near alfalfa should be targeted for survey. Shake leaves above white paper or cloth, and use a hand lens to observe mites.

Surveys should be focused where the greatest risk for establishment occurs. A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that most states in the United States are at low to moderate risk for E. orientalis establishment based on climate and host availability. Florida, however, has a moderate to high risk for establishment of this mite. Establishment of E. orientalis is unlikely in portions of Colorado, Idaho, Indiana, Iowa, Kansas, Louisiana, Minnesota, Montana, Nebraska, New Mexico, North Dakota, Oklahoma, Oregon, South Dakota, Texas, Utah, Washington, and Wyoming.

Hall (1992) discusses sampling strategies for spider mites in orange groves. The author’s sampling method consisted of examining 16 leaves per tree, 5 trees within a small area of trees, and 3 areas per block. The leaves were collected by gently pulling four leaves from each of the north, east, south, and west sides of a tree. The leaves from each side of the tree were placed into separate plastic bags. The bags were placed in a cold ice chest, taken to the laboratory, and examined under a microscope to count the number of spider mites present per leaf (both surfaces).

Gilstrap and Browing (1983) recommend using a liquid sampling procedure for leaf collecting of mites, where leaves are placed in a jar filled with 0.5% liquid dishwashing soap and 0.5% standard bleach (5% NaCl) (each % by volume) in a solvent of distilled water. The liquid soap is used to break up surface tension; while the bleach is used to dissolve any webbing. The author showed that the liquid sampling procedure collected more mites than more mites than the ‘normal procedure’. In the ‘normal procedure’, leaves are placed in a paper bag and a mite brushing machine is used to dislodge mites from the samples when processed the next day. Dhorria et al. (1982) collected forty

130 Eutetranychus orientalis Secondary Pest of Corn Arthropods Citrus brown mite Mite

random leaves (10 leaves/tree) from each variety at different heights and all sides of the plants to assess mite resistance. A mite brushing machine was used to dislodge the mites from the leaves on to counting disks.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of E. orientalis is by morphological identification. The mite can only be identified by examination of the adult male.

Literature-Based Methods: According to a NAPPO pest alert, the only form of E. orientalis that can be identified is the adult male. Conflicting information states that identification of E. orientalis requires examination of cleared and mounted female specimens by transmitted light microscopy.

Mite experts agree that though it may be possible to identify a specimen with a slide mounted female, one can never be 100% sure without a male for confirmation. E. orientalis can be easily mistaken for the Texas citrus mite (E. banksii). Similarity of the female E. orientalis with other tetranychid mites such as the two-spotted mite () can make identification difficult.

References Avidov, Z. and Harper, I. 1969. Plant Pests of Israel. Israel Universities, Jerusalem.

Dhorria, M.S., Sandhu, G.S., Khangura, J.S., and Dhatt, A.S. 1982. Incidence of citrus mite, Eutetranychus orientalis (Klein) on different varieties of almond in Ludhiana. Punjab Hortic. J. 22(1/2): 110-112.

Dhooria, M.S. and Butani, D.K. 1984. Citrus mite, Eutetranychus orientalis (Klein) and its control. Pesticides 18(10): 35-38.

Gilstrap, F.E. and Browing, H.W. 1983. Sampling predaceous mites associated with citrus. PR Tex. Agric. Exp. Stn. 4149.

Hall, D.G. 1992. Sampling citrus and red mites and Texas citrus mites in young orange trees. Proc. Annu. Meet. Fla. State Hort. Soc. 105: 42-46.

Lal, L. 1977. Studies on the biology of the mite Eutetranychus orientalis (Klein) (Tetranychidae: Acarina). Entomol. 2(1): 53-57.

Meyer, M.K.P.S. 1974. A revision of the Tetranychidae of Africa (Acari) with a key to genera of the world. Entomology Memoir, Department of Agricultural Technical Services, Republic of South Africa No. 36.

Osman, A.A. 1976 (publ. 1980). The role of alfalfa in dispersing tetranychid mites to other crops. Bull. Soc. Ent. Egypte 60: 279-283.

Smith-Meyer, M.K.P. 1981. Mite pests of crops in southern Africa. Science Bulletin, Department of Agriculture and Fisheries, Republic of South Africa, (No. 397): 65-67.

131 Metamasius spp. Secondary Pest of Corn Arthropods Metamasius weevils Weevil

Metamasius spp.

Scientific Name Metamasius hemipterus

Note: There are approximately 100 Neotropical species in this genus. For this data sheet, information is at genus level unless otherwise noted. M. hemipterus is the only reported pest of corn. Vaurie (1966) gives information on 57 Metamasius spp. including taxonomy, anatomy, ecology, sexual dimorphism and distribution.

Synonyms: Calandra hemipterus sericeus, Calandra sacchari, hemipterus, Curculio rufofasciatus, Curculio variegatus, Rhynchophorus hemipterus carbonarius, Sphenophorus ambiguus, Sphenophorus decoratus, Sphenophorus hemipterus, Sphenophorus inscripta, Sphenophorus sacchari, and Sphenophorus sexguttatus.

Common Names West Indian cane weevil, rotten cane stalk borer, rotten sugarcane weevil, silky cane weevil, weevil borer, and West Indian sugarcane borer.

Type of Pest Weevil

Taxonomic Position Class: Insecta, Order: Coleoptera, Family: Curculionidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 (as Metamasius spp.)

Pest Description The following information is from Vaurie (1966) unless noted otherwise.

General (all species in genus): Metamasius spp. are associated with orchids, bromeliads, palms, sugarcane, bananas and cacti (Vaurie, 1966). Adults may be found in the juicy interior of cane or bananas lying on the ground or in or under the sheaths of palms on the ground. Many species have been collected at very high altitudes in the mountains; others, such as those in bananas or cane, at sea level, along rivers, and in valleys (Vaurie, 1966).

Species of the genus Metamasius are generally medium to large (from about 10 to 20 mm in length), elongate, torpedo-shaped, or spindle-shaped (fusiform), black in ground color, but usually banded, spotted, or streaked with bright or dull red, orange, or yellow, the colors are interchangeable with the black in many species; a few species are gray, with black velvety spots. In contrast to many weevils, these species have no scales or dorsal hairs, although several have a rather tomentose covering or coating. They are

132 Metamasius spp. Secondary Pest of Corn Arthropods Metamasius weevils Weevil

diurnal, and good fliers, except for three species with reduced wings (M. cornurostris, M. fahraei, and M. foveolatus). All species seem to pupate in the host plant and construct a fibrous cocoon (O’Brien and Thomas, 1990).

Metamasius (From CABI, 2007) Eggs: The egg is yellowish cream, approximately 1.7 mm long, ovoid and semitransparent. Eggs are often laid in cracks or damaged areas of the host plants.

Larvae: The larvae are white and robust with a width of 3.2-4.5 mm. Thoracic and abdominal sternites are yellow in color; while the head is brown with paler stripes on the dorsal side. Body length is 15-17 mm. Three dorsal folds are present on the abdominal segments; while the 9th abdominal segment is either smoothly rounded or transverse. Abdominal segments 1-8 have distinct spiracles. The larvae are described by Cotton (1924) and by Anderson (1948). They provided a generic key to the larvae of Rhynchophorinae, including Metamasius. Figure 1. Dorsal and side view of M. hemipterus. Photos courtesy of Pupae: The pupa is elongate, narrow and Natasha Wright. contracted both anteriorly and posteriorly and www.forestryimages.org is approximately 14.5 mm in length. Five pairs of functional abdominal segments are visible from above.

Adults: Length of adults is around 9-14 mm with color varying from red to orange and black all over the body (Fig. 1) (Weissling and Giblin-Davis, 2007). A detailed description of the adult stage can be found in Vaurie (1966).

Some authors believe that M. hemipterus contains three subspecies; M. hemipterus hemipterus, M. hemipterus sericeus, and M. hemipterus carbonarius (Vaurie, 1966). Differences are found in the color patter, of the elytra, pronotum, or venter but not in form (Vaurie, 1966).

Symptoms/Signs Metamasius spp. are generally secondary pests that are found in decaying or rotting trunks or stems; however, they can attack healthy plants (Vaurie, 1966). Some species attack bromeliad or orchid leaf bases (Vaurie, 1966). Most of the time, the plants or plant parts are already in bad condition but further damage is caused by the weevil galleries (Vaurie, 1966).

133 Metamasius spp. Secondary Pest of Corn Arthropods Metamasius weevils Weevil

M. hemipterus usually attacks damaged or unhealthy host plants, but it has caused losses in bananas, pineapple, and sugarcane in the Caribbean (Woodruff and Baranowski, 1985; O’Brien and Thomas, 1990). M. hemipterus sericeus is an important pest of palms, sugarcane, pineapple, and bananas in the West Indies, Mexico, Central and South America (Giblin-Davis et al., 1996). M. hemipterus larvae can bore into sugarcane and banana stems and occasionally sheaths (Vaurie, 1966). Corn is considered a minor host for this pest. Specific symptoms in corn are not available at this time.

The plant is weakened as M. hemipterus sericeus larvae bore into the stems and petioles, providing a pathway for penetration by fungi or other pests (Weissling et al., 2003). As soon as larvae development begins, the cane stalks begin to rot, giving off a distinctive acetic acid odor (Wolcott, 1948). In sugarcane, growth slows, plants turn yellow and stalks become riddled with large galleries (Wyniger, 1962). This weevil has already caused some economic loss in Florida on sugarcane cultivar CP85-1382 (Sosa et al., 1997). In banana, plant growth slows, leaves wilt and wither, young plants will turn yellow and collapse and pseudostems, which are heavily mined, will often break (Wyniger, 1962). In palm, larval tunneling through the stem, petioles and leaves of palms usually results in production of a gummy, amber-colored exudate (Giblin-Davis et al. 1996). Infestations can often go undetected in trees, such as Canary Island date palm, causing an increase in weevil population. Host palm damage can be severe in some instances but infestations are rarely lethal. In Colombia, however, this weevil is associated with and is a possible vector of red ring disease caused by Bursaphelenchus cocophilus (a ) in palm trees (Ramirez-Lucas et al., 1996).

Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Surveys should be focused where the greatest risk for establishment occurs. A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that most states in the United States are at low to moderate risk for Metamasius spp. based strictly on availability of hosts. Metamasius hemipterus (resembling subspecies sericeus) is established in parts of Florida. Giblin-Davis et al. (1994) found that M. hemipterus is established in Dade, Broward and Palm Beach Counties in Florida.

Trapping: Because weevils are attracted to damaged and rotting plant materials, surveying for this genus may be conducted through trapping using host plant material. Giblin-Davis et al. (1994, 1996) used a semiochemical-based lethal pitfall trap to catch M. hemipterus sericeus. The trap consisted of two translucent white, high density polyethylene (HDPE) plastic tapered containers; one 4.9 and the other 0.9 liter in size. Two drainage holes in the sides, 6 cm from the bottom, allowed for rain overflow. The smaller container had a screened lid (8.5-cm screened opening), to retain the sugarcane and prevent weevils from interacting with it when inverted, and was suspended with a vinyl-coated wire inside the larger container. Each assembled trap was buried to within about 2.5 cm of the trap opening in the shade of a mature banana plant. The insect can be killed by adding soapy water in the larger container.

134 Metamasius spp. Secondary Pest of Corn Arthropods Metamasius weevils Weevil

Giblin-Davis et al. (1996) also evaluated kariomones released as sugarcane ferments using two devices: 1) 1 ml of a test chemical in 2-ml Gold Brand ampule with a 4-mm diameter opening and 2) 4 or 8 ml of the test chemical in a 14.8-ml vial without a top (11.4-mm-diam. opening). Release devices were placed in a 120-ml HDPE specimen container suspended from inside of the 0.9 liter test stimulus container by wires. Giblin- Davis et al. (1996) determined that the most cost-effective semiochemical-based bait was a combination of sugarcane (250 g), ethyl acetate (400-800 mg/day), and metalure (3 mg/day). If needed, the sugarcane component can be replaced with 45 g of molasses and 158 ml of water. Trap placement and color of trap body did not seem to affect efficacy of trap (Giblin-Davis et al., 1996).

Aggregation pheromones produced by M. hemipterus may also be used for survey trapping. Perez et al. (1997) found that the aggregation pheromone for M. hemipterus was a mixture of 4-methyl-5-nonanol and 2-methyl-4-heptanol, both of which should be used (termed metalure).

Trapping for both M. hemipterus and Rhynchophorus palmarum (palm weevil) can be accomplished with the same trap by using a combination lure (Chinchilla et al., 1996; and Alpizar et al., 2002).

Not recommended: Visual survey may be used in conjunction with trapping but should not be used alone for early detection surveys because host symptoms may not be distinct enough to determine presence.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: The genus Metamasius can be distinguished from other similar genera by looking at morphological characteristics. Vaurie (1966) has a key to help distinguish species of Metamasius, as well as a key to distinguish between the three subspecies, M. hemipterus carbonarius, M. hemipterus hemipterus, and M. hemipterus sericeus. The weevil is easily distinguished from other Florida species by the color (red to yellow and black) and size (9-14 mm long). Similar species include the banana root borer (Cosmopolites sordidus) and the yucca weevil (Scyphophorus acupunctatus), but both are uniformly black (Woodruff and Baranowski, 1985).

Metamasius species can be taxonomically difficult to distinguish because of the large number of species in the genus (O’Brien and Thomas, 1990). M. hemipterus is similar to both M. benoisti and M. ensirostris (Vaurie, 1966). M benoisti differs in having a rostrum which is longer and significantly bent, while in males, the median lobe of the aedeagus has a slight knob or projection apically. M. ensirostris differs from M. hemipterus in lacking any emargination or knobs on the scutellum and the males having front tibiae dotted with punctures, leaving no free space, and the last segment of the abdomen with a distinct oval patch of long erect hairs (not scattered, reclining hairs); female differing by lacking pale, tomentose basal patch on rostrum. The only native

135 Metamasius spp. Secondary Pest of Corn Arthropods Metamasius weevils Weevil

Metamasius in the United States is M. mosier, which lives in bromeliads. It is red and black but there are two round spots on the elytra and it is only about 6-9 mm long (Woodruff and Baranowski, 1985).

References Alpizar, D., Fallas, M., Oehlschlager, A.C., Gonzalez, L.M., Chinchilla, C.M., and Bulgarelli, J. 2002. Pheromone mass trapping of the West Indian sugarcane weevil and the American palm weevil (Coleoptera: Curculionidae) in Palmito palm. Florida Entomologist 85(3): 426-430.

Anderson, W.H. 1948. Larvae of some genera of Calendrinae (=Rhynochophrinae) and Stromboscerinae. Annals of the Entomological Society of America 41: 413-437.

CABI. 2007. Crop Protection Compendium. Wallingford, UK: CAB International. http://www.cabicompendium.org/cpc.

Chinchilla, C., Oeshlschlager, C., and Bulgarelli, J. 1996. A pheromone based trapping system for Phynchophorus palmarum and Metamasius hemipterus. ASD Oil Palm Papers no. 12: 11-17.

Cotton, R.T. 1924. A contribution toward the classification of the weevil larvae of the subfamily Calendrinae occurring in North America 66(5): 1-11.

Giblin-Davis, R.M., Pena, J. E., and Duncan, R.E. 1994. Lethal pitfall trap for evaluation of semiochemical-mediated attraction of Metamasius hemipterus sericeus (Coleoptera: Curculionidae). Florida Entomologist 77(2): 247-255.

Giblin-Davis, R.M,, Pena, J.E., Oehlschlager, A.C., and Perez, A.L. 1996. Optimization of semiochemical-based trapping of Metamasius hemipterus sericeus (Olivier) (Coleoptera: Curculionidae). Journal of Chemical Ecology 22(8): 1389-1410.

O’Brien, C.W. and Thomas, M.C. 1990. The species of Metamasius in Florida (Coleoptera: Curculionidae). Florida Department of Agriculture and Consumer Services, Division of Plant Industry, Entomology Circular 330.

Perez, A.L., Campos, Y., Chinchilla, C.M., Oehlschlager, A.C., Gries, G., Gries, R., Giblin-Davis, R.M., Castrillo, G., Pena, J.E., Duncan, R.E., Gonzalez, G., Pierce Jr., H.D., McDonald, R., and Andrade, R. 1997. Aggregation pheromones and host kairomones of West Indian sugarcane weevil, Metamasius hemipterus sericeus. Journal of Chemical Ecology 23(4): 869-888.

Ramirez-Lucas, P., Rochat, D., and Zagatti, P. 1996. Field trapping of Metamasius hemipterus with synthetic aggregation pheromone. Entomologia Experimentalis et Applicata 80: 453-460.

Sosa, O., Shine, J., and Tai, P.Y.P. 1997. West Indian cane weevil (Coleoptera: Curculionidae): a new pest of sugarcane in Florida. J. Econ. Entomol. 90: 634-638.

Vaurie, P. 1966. A revision of the neotropical genus Metamasius (Coleoptera, Curculionidae, Rhynchophorinae) species groups I and II. Bulletin of the American Museum of Natural History 131(3): 132 pp.

Weissling, T., Giblin-Davis, R., Center, B., Heath, R., and Pena, J. 2003. Oviposition by Metamasius hemipterus sericeus (Coleoptera: Dryophthoridae: Rhynchophorinae). Florida Entomologist 86(2): 174- 177.

Weissling, T. and Giblin-Davis, R., 2007. Silky cane weevil, Metamasius hemipterus sericeus (Oliver) (Insecta: Coleoptera: Dryophthoridae). University of Florida IFAS Extension. EENY-053. 6 pp.

136 Metamasius spp. Secondary Pest of Corn Arthropods Metamasius weevils Weevil

Wolcott, G. N. 1948. The Insects of Puerto Rico: Coleoptera. Journal of Agriculture of the University of Puerto Rico, 32: 225-416.

Woodruff, R. E. and Baranowski, R. M. 1985. Metamasius hemipterus (Linnaeus) recently established in Florida. Florida Department of Agriculture, Division of Plant Industry, Entomology Circular 272: 1-4.

Wyniger, R. 1962. Pests of crops in warm climates and their control. Acta Tropica Supplementum 7. Basel, p. 1-555.

137 Oxycarenus hyalinipennis Secondary Pest of Corn Arthropods Cottonseed bug True bug

Oxycarenus hyalinipennis

Scientific Name Oxycarenus hyalinipennis Costa

Synonyms: Aphanus hyalinipennis and Aphanus tardus var. hyalinipennis

Common Name(s) Cotton seed bug, cotton stainer, dusty cotton stainer, dusky cotton bug, dusky cottonseed bug, and Egyptian cotton seed bug

Type of Pest Bug

Taxonomic Position Class: Insecta, Order: Hemiptera, Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description Eggs: Oval 0.28 x 0.95 mm, longitudinally striated, pale yellow becoming pink. Eggs are laid singly or in groups of 2-4 or rarely more (Hammed et al., 1982). The eggs that are laid in groups are stuck to each other side-by- Figure 1. Adult O. hyalinipennis. Photo side by sticky material. courtesy of Georg Goergen/IITA Insect Museum, Cotonou, Benin. CABI (2007). Nymphs: Head and thorax brownish- olivaceous, abdomen pinkish. Fifth instar darker brown on head and thorax, wingpads distinct, extending to at least third abdominal segment. The nymph undergoes five ecdyses before attaining maturity.

Adults: (Fig. 1, 2) Newly emerged individuals are pale pink, but rapidly turn black. Length of male about 3.8 mm; female 4.3 mm. Male abdomen terminates in round lobe, while the female’s is truncate. The insects have three tarsal joints and a pair of ocelli. Second antennal segments are usually in part pale yellow. Hemelytra hyaline and usually whitish; clavus, base of corium, and costal vein more opaque than rest. Setae of 3 different types: More or less erect, stiff setae, which are blunt at tip and terminate in 4- 7 small teeth; normal, straight, tapering setae; and very thin, curved, flat-lying setae (USDA, 1983).

138 Oxycarenus hyalinipennis Secondary Pest of Corn Arthropods Cottonseed bug True bug

Oxycarenus hyalinipennis begins feeding, mating, and egg laying when the seeds of its host become available. Resting adults leave their shelters, move to young cotton plants, and wait for the bolls to ripen. Females lay eggs in the lint of the open bolls. Adults and nymphs generally feed on the seeds of plants in the family Malvaceae. The last generation undergoes aestivation until seed material is available the next growing season (NPAG, 2003).

Symptoms/Signs Oxycarenus hyalinipennis is a seed feeder, with primary hosts occurring within the Malvaceae family, specifically Gossypium spp. (cotton). Currently there are 40 hosts reported in the literature from the Malvales order on which O. hyalinipennis is capable of reproduction. O hyalinipennis has been reported on several other hosts, including corn, in 11 different families. The ability of O. hyalinipennis to reproduce on these hosts and the level of feeding damage, however, is not known on these additional hosts (Holtz, 2006).

When hosts are unavailable, O. hyalinipennis will migrate to ‘shelters’ on hosts and non-hosts. Shelters may be comprised of trunks, the underside of leaves, the interior of legume pods, dried flower heads of weeds, crevices along roots in grasses, under sheath leaves of corn and sugar cane, telephone poles, wooden posts, crevices in barb wire, old paper wasp nests, and other sites. Figure 2. Adult O. hyalinipennis. On cotton, this lygaeid infests the seed in the field as the Photo courtesy of bolls open. Weight loss in cottonseed, decreased Natasha Wright. germination, and reduced oil quality of the seed is www.forestryimage observed. The lint, where bugs have been present is s.org stained pinkish, sometimes with a trace of green, and contaminated with crushed fragments of the insect. This is due to the bugs being crushed during ginning. Cotton seeds appear undamaged on the outside; internally, the embryos are shriveled and discolored (USDA, 1983).

Survey CAPS-Approved Method: Visual survey is the method to survey for O. hyalinipennis.

Literature-Based Methods: O. hyalinipennis has been intercepted a few times each year at U.S. ports of entry. All interceptions occurred at airports, mostly in baggage; no interceptions were recorded from preferred malvaceous hosts. These interceptions point to the risk of O. hyalinipennis moving on commodities that are not its reproductive hosts (NPAG, 2003). O. hyalinipennis was recently found in Puerto Rico and Florida, but its current distribution is unknown.

139 Oxycarenus hyalinipennis Secondary Pest of Corn Arthropods Cottonseed bug True bug

Visual survey: Visual inspection is the only survey method available at this time. Samy (1969) observed adult clusters on leaves of mango, guava, and citrus. For cotton, cotton bolls can be tapped or torn open and examined for evidence of O. hyalinipennis. Additionally, sweep netting of weeds between cotton rows or field edges is recommended. Adults prefer crevices in such resting sites as tree trunks, undersides of leaves on trees, pods of legumes, dried flower heads, roots of grasses, under sheath leaves of corn and sugarcane, telephone poles or wooden posts, old nests of Polistes spp. (paper wasps), and crevices between strands of barbed wire (Kirkpatrick, 1923).

Surveys should be focused where the greatest risk for establishment occurs. According to Holtz (2006), O. hyalinipennis could potentially complete 4-7 generations per year in all areas where U.S. cotton is produced. Based on a probability map, California, Arizona, and Texas may be most vulnerable to O. hyalinipennis. A recent risk analysis by USDA-APHIS-PPQ-CPHST, however, indicates that most states in the United States are at risk for O. hyalinipennis establishment based on climate and host availability. Areas of Alabama, Arkansas, Arizona, California, Delaware, Illinois, Indiana, Iowa, Florida, Georgia, Kansas, Kentucky, Louisiana, Maryland, Massachusetts, Mississippi, Missouri, Nebraska, New York, North Carolina, Ohio, Oklahoma, Pennsylvania, South Carolina, Tennessee, Texas, Utah, Virginia, West Virginia, and Wisconsin have the highest levels of risk for establishment of O. hyalinipennis. Surveys should pay particular attention to Florida and states along the Gulf Coast, as O. hyalinipennis is present in the West Indies (Slater and Baranowski, 1994) and the Bahamas (Randall Smith and Brambila, 2008).

In O. hyalinipennis, the metathoracic glands appear similar in males and females, but a day or so after emergence the tubular glands of both sexes undergo a dramatic change from synthesizing aliphatics to the synthesis of sesquiterpenes, principally (Z,E)-α- farnesene (Olagbemiro and Staddon, 1983; Knight et al., 1984). Although many explanations for this phenomenon have been proposed, it appears that the metathoracic scent glands may have evolved sexual roles in this lygaeid species.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of O. hyalinipennis is by morphological identification.

Literature-Based Methods: The key diagnostic involves morphological examination of adults. A screening aid is available for O. hyalinipennis at http://caps.ceris.purdue.edu/webfm_send/529.

References Avidov, Z. and Harpaz I. 1969. Plant Pests of Israel. Jerusalem, Israel University Press.

CABI. 2007. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Holtz, T. 2006. Qualitative analysis of potential consequences associated with the introduction of the cottonseed bug (Oxycarenus hyalinipennis) into the United States. USDA-APHIS.

140 Oxycarenus hyalinipennis Secondary Pest of Corn Arthropods Cottonseed bug True bug

Kirkpatrick, T.W. (1923). The Egyptian cottonseed bug (Oxycarenus hyalinipennis (Costa). Its bionomics, damage, and suggestions for remedial measures. Bull. Minist. Agric. Egypt Tech. Sci. Serv. 35: 28-98.

Knight, D.W., Rossiter, M., and Staddon, B.W. 1984. (Z,E)-α-Farnesene: major component of secretion of metathoracic scent gland of cotton seed bug, Oxycarenus hyalinipennis (Costa) (: Lygaeidae). Journal of Chemical Ecology 10: 641-649.

New Pest Advisory Group (NPAG). 2003. Oxycarenus hyalinipennis (Costa): Cotton Seed Bug. USDA- APHIS-PPQ-CPHST.

Olagbemiro, T. and Staddon, B.W. 1983. Isoprenoids from metathoracic scent gland of cotton seed bug, Oxycarenus hyalinipennis (Costa) (Heteroptera: Lygaeidae). Journal of Chemical Ecology 9: 1397-1412.

Randall Smith, T. and Brambila, J. 2008. A major pest of cotton, Oxycarenus hyalinipennis (Heteroptera: ) in the Bahamas. Florida Entomologist 91(3): 479-482.

Samy, O. 1969. A revision of the African species of Oxycarenus (Hemiptera:Lygaeidae). Royal Entomol. Soc. London Trans. 121(4): 79-165.

Slater, J.A, and Baranowski, R.M. 1994. The occurrence of Oxycarenus hyalinipennis (Costa) (Hemiptera: Lygaeidae) in the West Indies and new Lygaeidae records for the Turks and Caicos Ilsnada of Providenciales and Nort Caicos. Florida Entomologist 77(4): 495-497.

USDA. 1983. Pest not known to occur in the United States or of limited distribution, No. 38: Cottonseed bug.

141 Planococcus minor Secondary Pest of Corn Arthropods Passionvine Mealybug

Planococcus minor

Scientific Name Planococcus minor Maskell

Synonyms: Planococcus pacificus, Pseudococcus minor, Dactylopius calceolariae minor, Planococcus psidii, and Pseudococcus calceolariae minor.

Common Name(s) Passionvine mealybug and Pacific mealybug A

Type of Pest Mealybug

Taxonomic Position Class: Insecta, Order: Homoptera, Family: Pseudococcidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description Planococcus minor is a small sucking B insect with a cottony appearance. . Females are oval, 1.3 to 3.2 mm long. The insect body is distinctly segmented, yellow to pink in color, and covered with powdery wax, with the appearance of "having been rolled in flour" (Fig. 1A). The margin of the body has a complete series of 18 pairs of cerarii, each cerarius with 2 conical setae (except for preocular cerarii which may have 1 or 3 setae). Legs are elongate.

It is assumed that this species is identical in appearance to P. citri (Fig. 1B) as follows: body oval; slightly rounded in Figure 1: Planococcus minor (A) and P. lateral view; body yellow when newly citri (B). Photos courtesy of Jeel Miles molted, pink or orange-brown when fully (www.invasive.org) and J. V. French. mature; legs brown-red; mealy wax covering body, not thick enough to hide body color; with dorsomedial bare area on dorsum forming central longitudinal stripe

142 Planococcus minor Secondary Pest of Corn Arthropods Passionvine mealybug Mealybug

(more obvious than on P. ficus); ovisac ventral only, may be 2 times longer than body when fully formed; with 17 or 18 lateral wax filaments, most relatively short, often slightly curved, posterior pair slightly longer, filaments anterior of posterior pair small, posterior pair about 1/8 length of body. Primarily occurring on foliage of host. Oviparous, eggs yellow. Surface of lateral filaments rough (Rung et al., 2007).

Mealybugs produce honeydew, which is a liquid rich in sugar. Ants like to feed on honeydew and some ants will, therefore, protect the by chasing away predators and parasitoids. The ants also carry mealybugs around and thus contribute to their distribution. P. citri was reported as a virus vector in cocoa, banana, and grape, but whether P. minor can serve as a vector is unknown (Jones and Lockhart, 1993; Canaleiro and Segura, 1997).

Symptoms/Signs Mealybugs have piercing-sucking mouthparts. Planococcus minor is a phloem feeder, and in general this may cause reduced yield, reduced plant or fruit quality, stunting, wilting, discoloration, and defoliation. Indirect or secondary damage is caused by sooty mold growth on honeydew produced by the mealybug.

Survey CAPS-Approved Method: Visual survey is the method to survey for P. minor

Literature-Based Methods: (From Venette and Davis, 2004) Visual survey: In the United States, surveys for mealybugs other than P. minor require “time-consuming and often laborious examination of plant material for the presence of live mealybugs” (Millar et al., 2002). No simple, alternative techniques are available (Millar et al., 2002). The same holds true for P. minor surveys in other parts of the world. In India, a regional survey for scales and mealybugs, including P. minor, was based on visually examining 25 branches or leaves on each of 15 plants collected from each of 3 field sites in 162 locations (25 x 15 x 3 x 162 = 182,250 leaves examined).

Researchers also depend on visual inspections to assess densities of P. minor. In a study of P. minor population dynamics, populations of the mealybug were evaluated by visual inspection of citrus leaves, specifically 10 to 15 leaves from 10 randomly selected plants (Bhuiya et al., 2000). Reddy et al. (1997) followed a similar protocol for coffee.

Surveys should be focused in areas most at risk for establishment of P. minor. The host range of P. minor includes a wide range of plants grown in the United States, so this insect appears capable of establishing populations that mirror the distribution of P. citri. P. citri is present in the southern states and has been reported as far north as Ohio, Kansas, and Massachusetts. Venette and Davis (2004) estimate that approximately 52% of the continental United States would have a suitable climate for P. minor. A recent host analysis by USDA-APHIS-PPQ-CPHST indicates that portions of Arkansas, Illinois, Indiana, Iowa, Minnesota, Missouri, Nebraska, Ohio, South Dakota, have the greatest risk for P. minor establishment based on host availability within the continental United States.

143 Planococcus minor Secondary Pest of Corn Arthropods Passionvine mealybug Mealybug

Trapping: A sex pheromone has been identified for P. minor. Ho et al. (2007) identified the sex pheromone as (E)-2-isopropyl-5-methyl-2,4-hexadienyl acetate. The (Z) isomer was found to be highly antagonistic. Millar (2008) describes the short and completely stereospecific synthesis of (E)-2-isopropyl-5-methyl-2,4-hexadienyl acetate. The availability of this pheromone is not known at this time.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of P. minor is by morphological or molecular identification. Adult females can be identified by a qualified taxonomist using a matrix of morphological characters; immatures can be easily confused with other Planococcus species and other mealybug genera. For molecular analysis, maintain some specimens in 95-100% alcohol for DNA analyses. Final identification will be based on morphological identification of adult female, and for new state records, followed by molecular analysis for confirmation.

Literature-Based Methods: (from Venette and Davis, 2004) Infestations reduce the vigor and growth of foliage plants, which reduces the beauty of the plant and affects marketability (Hamlen, 1975). Mealybugs are a quarantine problem on exported foliage and flowers. This is due to the fact that species cannot be accurately identified outside of the lab, so inspectors/surveyors should treat all specimens as unknown species. There are a large number of endemic species of mealybugs in the United States and identifications need to be made by a recognized taxonomic authority.

Planococcus species are not easily distinguishable from one another, especially when immature. A level of complexity is added with variable morphological characters in some species; distinguishing morphological characters can change depending on environmental conditions, such as temperature. Distinguishable morphological features of closely related mealybug species are described by Cox (1981, 1983, and 1989). A Lucid tool for scale insects has been recently developed, which contains a tool on mealybugs (see http://www.sel.barc.usda.gov/ScaleKeys/ScaleInsectsHome/ScaleInsectsMealybugs.ht ml).

Planococcus citri and P. minor have been taxonomically confused and routinely misidentified as adults are similar in appearance and share similar hosts and geographic range (Williams, 1985; Cox, 1989; Williams and Granara de Willink, 1992). Adults (females) can be identified based upon close examination of morphological characters by a taxonomist. PPQ initiated a project to develop molecular diagnostics to separate P. citri from P. minor. A PCR-RFLP technique was developed to distinguish P. citri, P. minor, and a genetically distinct Planoccocus that is morphologically identical to P. citri from Hawaii (Rung et al., 2008, 2009).

References Bhuiya, B.A., Chowdhury, S.H., Kabir, S.M.H., Austin, A.D., and Dowton, M. 2000. Natural population of Aenasius advena Compere (Chalcidoidea: Encyrtidae) and its host preference in Bangladesh, pp. 417- 420, In: Fourth International Conference, held in Canberra, Australia. January 1999.

144 Planococcus minor Secondary Pest of Corn Arthropods Passionvine mealybug Mealybug

Canaleiro, C. and Segura, A. 1997. Field transmission of grapevine leafroll associated virus 3 (GLRaV- 3) by the mealybug Planococcus citri. Plant Dis. 283-287.

Cox, J.M. 1981. Identification of Planococcus citri (Homoptera: Pseudococcidae) and the description of a new species. Systematic Entomology 6: 47-53.

Cox, J.M. 1983. An experimental study of morphological variation in mealybugs (Homoptera: Coccoidea: Pseudococcidae). Systematic Entomology 8: 361-382.

Cox, J.M. 1989. The mealybug genus Planococcus (Homoptera: Pseudococcidae). Bulletin of the British Museum (Natural History) 58(1): 1-78.

Hamlen, R.A. 1975. Insect growth regulator control of longtailed mealybug, hemispherical scale, and Phenacoccus solani on ornamental foliage plants. J. Econ. Entomol. 68 (2): 223-226.

Ho, H.-Y., Hung, C.-C., Chuang, T.-H., and Wang, W.-L. 2007. Identification and synthesis of the sex pheromone of the passionvine mealybug, Planococcus minor (Maskell). J. Chem. Ecol. 33: 1989-1996.

Jones, D.R. and Lockhart, B.E.L. 1993. Banana streak disease. Musa fact sheet No.1. International Network for Improvement of Banana and Plantain. France: Montpellier.

Millar, J.G. 2008. Stereospecific synthesis of the sex pheromone of the passionvine mealybug, Planococcus minor. Tetrahedron Letters 49: 315-317.

Reddy, K.B., Bhat, P.K., and Naidu, R. 1997. Suppression of mealybugs and green scale infesting coffee with natural enemies in Karnataka (1997). Pest Management and Economic Zoology 5(2): 119- 121.

Rung (Baptista), A., Venable, G.L., Miller, D.R., Gill, R. J., Williams, D. J. 2007. Scale Insects: Identification tools, images, and diagnosis information for species of quarantine significance. Systemic Entomology Laboratory, USDA; Center for Plant Health Science and Technology, APHIS, USDA; National Identification Services, APHIS, USDA. http://www.sel.barc.usda.gov/ScaleKeys/ScaleInsectsHome/ScaleInsectsMealybugs.html.

Rung, A., Scheffer, S.J., Evans, G., and Miller, D. 2008. Molecular identification of two closely related species of mealybugs of the genus Planoccocus (Homoptera : Pseudococcidae). Ann. Entomol. Soc. Am. 101(3) : 525-532.

Rung, A., Miller, R.D., and Scheffer, S.J. 2009. Polymerase chain reaction-restriction fragment length polymorphism method to distinguish three mealybug groups within the Planococcus citri-P. minor species complex (Hemiptera : Coccoidea : Pseduococcidae). J. Econ. Entomol. 102(1): 8-12.

Venette, R.C. and Davis, E.E.. 2004. Mini Risk Assessment – Passionvine mealybug: Planococcus minor (Maskell)[Pseudococcidae: Hemiptera]. Cooperative Agricultural Pest Survey, Animal and Plant Health Inspection Service, US Department of Agriculture. Available on line at: http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/pminorpra.pdf . Accessed 19 June 2007.

Williams, D.J. 1985. Australian mealybugs. British Museum (Natural History), London.

Williams, D.J. and Granara de Willink, M.C. 1992. Mealybugs of Central and South America. CAB International, Wallingford.

145 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

Rhabdoscelus obscurus

Scientific Name Rhabdoscelus obscurus (Boisduval, 1835)

Synonyms: Calandra obscura, Rhabdocnemis beccarii, Rhabdocnemis fausti, Rhabdocnemis interruptecostata, Rhabdocnemis interruptocostatus, Rhabdocnemis maculata, Rhabdocnemis nudicollis, Rhabdocnemis obscura, Rhabdocnemis obscurus, Rhabdocnemis promissus, Rhabdoscelis obscura, Rhabdoscelus maculatus, Rhabdoscelus obscurus, Sphenophorus obscurus, Sphenophorus beccarii, Sphenophorus insularis, Sphenophorus interruptecostatus, Sphenophorus nudicollis, Sphenophorus obscura, Sphenophorus obscurus, Sphenophorus promissus, Sphenophorus sulcipes, and Sphenophorus tincturatus

Common Names Sugarcane weevil borer, New Guinea cane weevil borer, beetle borer, cane weevil borer, New Guinea sugarcane weevil, cane weevil borer, and Hawaiian sugarcane borer

Type of Pest Weevil

Taxonomic Position Class: Insecta, Order: Coleoptera, Family: Dryophthoridae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description Rhabdoscelus obscurus is native to New Guinea and surrounding islands but has spread to almost all of the sugarcane growing areas in the Pacific. This pest is present in Hawaii but exotic to the contiguous United States. R. obscurus is considered a significant pest of sugarcane, palms, and banana and a secondary pest of corn.

Figure 1: Rhabdoscelus obscurus. Eggs: The eggs of R. obscurus are 1-2 mm Photo courtesy of Anthony O'Toole. in length, ivory white in color and slightly http://www.ento.csiro.au/aicn/name_s/ curved (USDA, 1967). Eggs change from a b_3568.htm translucent white with thick chorion to an

146 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

opaque white as they age; incubation period can range from 3-7 days (Napompeth et al., 1972). Eggs are laid singly into small holes or split tissues (Agnew, 1997)

Larvae: The mature larvae are approximately 15 mm in length with a hard brown head and whitish, almost transparent, body (USDA, 1967). The larvae are fleshy with two or more folds on each dorsal segment and are legless oval-shaped grubs (Napompeth et al., 1972). Both the head and mandibles are highly sclerotized; while the cervical shield is less sclerotized (Napompeth et al., 1972). The larva is covered sparsely with fine, stiff hairs (USDA, 1967). Figure 2. R. obscurus (left) compared with The last few abdominal segments Cosmoplites sordidus (right), the banana have longer stiff hairs that may help weevil. with larval movement within the http://www.ctahr.hawaii.edu/nelsons/banana/. tunnel (Napompeth et al., 1972).

Both the fifth and sixth abdominal segments are markedly expanded on the ventral side (Zimmerman, 1968). A plug of plant fibers usually blocks the entrance to their tunnel (Schreiner, 2000). Napompeth et al. (1972) stated that there were six larval instars, not including the prepupal stage, and the mean larval period of the pest lasted 54.3 ± 3.7 days.

Pupae: When R. obscurus is ready to pupate, the weevil constructs a large cocoon made from host plant fibers. In southeastern Polynesia, this is the only known weevil to construct such a cocoon and can thus be used to identify the infestation (Zimmerman, 1968).

According to Napompeth et al. (1972), after passing the 6th stadium, the full-grown larva transforms to a prepupa. Its general body shape differs from that of the 6th instar larva by the absence of the posterior enlargement. Pupation usually takes place within the cocoon, which is found in the tunnel made by the larva. It usually takes the prepupa 24 to 48 hours to transform into the yellowish white exarate pupa. Pupation takes place within a spirally woven fibrous cocoon. The frass, masticated and left in the tunnel, is also utilized in making the cocoon. However, the adult does not emerge from the cocoon immediately but remains inactive within it for a considerable period. Upon emergence from the cocoon, most of the adults are light in color while some are dark.

Adult: The adult (Figs. 1, 2) is rather large, ranging from 12-14 mm in length. The body of R. obscurus is primarily reddish to reddish brown, while the head is darker. The

147 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

pronotum has a medium black stripe that extends from the apex to base with “less distinct black marks on middle of elytra, sides of thorax and undersides of body” (USDA, 1967). The distal pilose section of the last segment on the antennae is wedge-shaped.

According to Napompeth et al. (1972), the oblong body with a protruding rostrum and well developed prothorax are typical of the curculionid subfamily Calandrinae. The anterior end of the rostrum bears the sclerotized mandibles. The dorsal body coloration is predominantly brown with a lighter shade of brown on the prothorax. The coloration of the elytra varies considerably, but generally has lateral and central dark brown patches. The elytra are well developed with longitudinal grooves or striae. The hind wings are membranous and strongly developed. The tarsal formula is 4:4:4. In the case of R. obscurus, two distinct morphological characteristics are present and they may be employed in distinguishing the sexes. The rostrum of the male is shorter, less curved, and more robust than that of the female. It is also ventrally serrated with a double row of highly sclerotized tubercles, varying in number from 5 to 8 pairs. Another morphological difference between the sexes may be found in the last abdominal tergite or the pygidium which usually protrudes slightly beyond the tip of the elytra in both sexes. Variation in color patterns of the adults of R. obscurus was first observed by Muir and Swezey (1916). They reported that the median dorsal marking on the pronotum and those on the elytra varied in size and shape considerably among specimens collected from different localities in the Pacific.

Most adult activity is at dusk, with beetles able to fly more than 500 meters. In lab experiments, longevity of adults can be high at around 160 days for both sexes (Napompeth et al., 1972).

Symptoms/Signs Although corn is listed as a host by Napompeth et al. (1972) and Zimmerman (1968), specific symptoms for corn are not given. Corn is considered a secondary and infrequent host.

Sugarcane: R. obscurus is a serious pest of sugarcane and is also one of the most important sugarcane pests in Hawaii. Fortunately, the tachinid parasite, Lixophaga sphenophori, has greatly helped to reduce the once annual losses of over half a million dollars in the state. However, this parasite has failed to control the pest in Fiji where sugarcane losses are severe (USDA, 1967).

Damage is usually associated with cane damaged through other means, such as rat damage, cane knife cuts, splits in stalks, and other stem borers. Damage is initiated at approximately four months after planting and is more severe on ratoon cane (regrowth after harvest) due to reinfestation by the weevil that remains in the stubble after harvest. Newly tunneled sugarcane stalks show an accumulation of frass in tunnels and reddening of the surrounding tissues. External signs of infestation are tiny exit or breathing holes 4-6 mm in diameter, also called windows or ‘windowing’ to regulate atmospheric conditions within the tunnel, and frass exuding at internodes.

148 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

Stalks can look healthy if the weevil attacks the bottom section of sugarcane when it is still soft and young, but the injuries encourage secondary attack by microorganisms such as red rot pathogen, Colletotrichum falcatum, to which most of the losses in sugarcane farming is attributed (Githure, n.d.; University of Queensland, n.d.). During cyclonic winds, infested stems can become heavily damaged through splitting and twisting (Agnew, 1997).

Palm: Adult females bore into the outer layer of stems and leaf bases of maturing palm trees and lay their eggs in small cavities. Eggs hatch into larvae (grubs), which develop inside the trunks resulting in exudation of pinkish sap. Larval damage has been observed from just above and adjacent to the root mass to two meters or more above the ground. The grubs then pupate in a cocoon of fibers inside the trunk. In young palms, the larvae mine the central portion of the stem, destroying the plants. Damage extends up and down the stem for a number of centimeters from the initial point of entry.

In older palms, R. obscurus mine the thicker leaf bases, as well as a short distance into the trunk. Older palms are disfigured by the emergence holes made by the weevils, and also by trunk splitting, rendering them unfit for sale. Heavy infestations may weaken the trunk sufficiently for the tree to collapse, with damage occurring mostly up to 1 meter above the ground. Jelly-like exudates form holes in leaf bases and/or stems may be observed. Signs of infestation include pin holes all over the trunk 2-3 feet above the ground. During heavy infestation a large number of grubs feed inside the palm, tunneling through and destroying the tissues. This leads to secondary infection by pathogens, resulting in weakening and falling over of the palms (Githure, n.d.). The “windowing” which occurs in sugar cane was not observed (Halfpapp and Storey, 1991). Trees can collapse and die with heavy infestations (Lake, 1998).

Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Trapping: Pheromone traps containing host material are used as a type of population control where the pest is prevalent. Giblin-Davis et al. (1996) determined that approximately 3 mg per day of synthetic pheromone with insecticide-treated plant host tissue was highly attractive as bait for palm weevils, including R. obscurus.

Chang and Curtis (1972) reported that male R. obscurus produce an aggregation pheromone that is attractive to both male and female weevils, but did not make an effort to identify the compound. Furthermore, the authors found that split-cane traps baited with mated or virgin male R. obscurus were more attractive than traps with or without female weevils. The aggregation pheromone of Hawaiian R. obscurus was later identified as 2-methyl-4-octanol; while the pheromone compounds of Australian R. obscurus are 2-methyl-4-octanol, (E2)-6-methyl-2-hepten-4-ol (rhynchophorol) and 2- methyl-4-heptanol (Giblin-Davis et al., 2000). However, 2-methyl-4-heptanol has not shown any noticeable behavioral effect on R. obscurus (Giblin-Davis et al., 2000). In previous studies, the addition of ethyl acetate and cut sugarcane to pheromone traps

149 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

significantly increased trap catches when added to pheromone lures (Muniappan et al., 2004; Reddy et al., 2005). The lures are reported to be commercially available (Muniappan et al., 2004).

Sallam et al. (2004) used split-cane traps, bundles of 6-8 split lengths of sugarcane, wrapped in black plastic with the ends left open to evaluate adult borer population fluctuations. Sallam et al. (2007) evaluated several types of traps when surveying for R. obscurus and found that the ‘water trap’ was the most effective at catching adults. This trap is described as 20 cm-diameter pot with a plastic bag inserted to hold water with pheromone lures and cane pieces held together in a plastic container suspended over the water from a square of wire mesh. This trap can attract borers from adjacent fields (Sallam et al., 2007).

Muniappan et al. (2004) and Reddy et al. (2005) used plastic bucket traps baited with the pheromone lure in combination with a food volatile compound (ethyl acetate) and cut sugarcane. The trap (Fig. 3) consisted of 19.0-1 white plastic- tapered containers (37 cm height x 30 cm inner diameter base). Two holes (17.5 cm long and 7.5 cm wide) were cut on opposite sides of the container to allow weevil entry into the trap. Twenty drainage holes, each 3 mm in diameter, were made in the base. Each assembled trap was placed at the base of a mature coconut tree in the field and strapped securely against it. Such a set- up helped the weevils walk into the trap. At each location, inter-trap distance was set at 100 meters. The pheromone lure was sealed in a polymer membrane release device optimized for the Australian population of R. obscurus [(E2)-6- Figure 3. Diagram of the methyl-2-hepten-4-ol and 2-methyl-4-heptanol) Reddy et al. (2005) trap and was suspended halfway inside the trap with a showing the various holes wire. Release devices for ethyl acetate lures were and bait placement. hung in the trap. The pheromone and ethyl acetate were changed at 4-month intervals. Fresh sugarcane sections were15 cm long and split in the middle along their length. The cut sugarcane was placed directly in the bucket trap and replaced weekly.

Surveys should occur in areas most at risk for establishment of R. obscurus. According to a recent host analysis by USDA-APHIS-PPQ-CPHST, portions of Arkansas, California, Florida, Georgia, Idaho, Illinois, Indiana, Iowa, Kansas, Kentucky, Louisiana, Maryland, Michigan, Minnesota, Mississippi, Missouri, Nebraska, New York, North Carolina, North Dakota, Ohio, Pennsylvania, South Carolina, South Dakota, Tennessee, Texas, and Wisconsin are at moderate risk for establishment of R. obscurus based on host availability alone.

150 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

Visual survey: Napompeth et al. (1972) states that sampling for the pest can be done effectively by checking for feeding scars on the leaf sheaths of cane. However, visual survey alone may not be adequate to detect the pest at low population densities.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: This pest has high variability in its general appearance, including size, color, and pattern, which can cause confusion when identifying (Zimmerman, 1968), as evident by its numerous synonyms over the years.

This pest can be identified through morphological characteristics. Key characteristics include: mesocoxae separated by more than the width of a mesocoxa, elytra spotted or blotched, pile raised in irregular vittae and pustules, and moderately large elytral strial punctures (Zimmerman, 1993). There are two characteristics that can be used to separate the male and female. The ventral side of the rostrum is smooth in females but roughened in males (Zimmerman, 1993). Also the last abdominal tergite (pygidium) is blunt in males and somewhat pointed in females (Napompeth et al., 1972).

R. obscurus resembles other sugarcane attacking weevils in the Americas and palm attacking weevils in Southeast Asia. Zimmerman (1968) provides a key to adults and larvae at the genus level. A key distinguishing R. obscurus from R. interstitialis, the two species present in Australia, is provided in Zimmerman (1993).

References Agnew, J.R. 1997. Australian sugarcane pests. Bureau of Sugar Experiment Stations, Indooroopilly, AU.

Chang, V.C.S. and Curtis, G.A. 1972. Pheromone production by the New Guinea sugarcane weevil. Environmental Entomology 1: 476-481.

Giblin-Davis, R.M., Oehlschlager, A.C., Perez, A., Gries, G., Gries, R., Weissling, T.J., Chinchilla, C.M., Pena, J.E., Hallett, R.H., Pierce Jr. H.D., and Gonzalez, L.M. 1996. Chemical and behavioral ecology of palm weevils (Curculionidae: Rhynchophorinae). Florida Entomologist 79(2): 153-167.

Giblin-Davis, R.M., Gries, R., Crespi, B., Robertson, L.N. Hara, A.H., Gries, G., O’Brien, C.W., and Pierce Jr., H.D. 2000. Aggregation pheromones of two geographical isolates of the New Guinea sugarcane weevil, Rhabdoscelus obscurus. Journal of Chemical Ecology 26(12): 2763-2780.

Githure, C. n.d. Rhabdoscelus obscurus. EcoPort Foundation. Retrieved on November 6, 2009 from: http://ecoport.org/ep?Arthropod=20025&entityType=AR****&entityDisplayCategory=full.

Halfpapp, K.H. and Storey, R.I. 1991. Cane Weevil Borer, Rhabdoscelus obscurus (Coleoptera: Curculionidae), in Palms in Northern Queensland, Australia. Principes 35: 199-207.

Lake, J. 1998. Getting control of weevil borers and leaf beetles in palms. The Nursery Papers 1998(02): 1-4.

Muir, F., and Swezey, OH. 1916. The cane borer in Hawaii and its control by natural enemies. Hawaiian Sugar Planters’ Assoc. Exp. Sta. Entomol. Ser. Bull. No. 13 102 pp.

151 Rhabdoscelus obscurus Secondary Pest of Corn Arthropods Sugarcane weevil Weevil

Muniappan, R., Bamba, J., Cruz, J., and Reddy, G.V.P. 2004. Field response of Guam populations of the New Guinea sugarcane weevil, Rhabdoscelus obscurus (Boisduval) (Coleoptera: Curculionidae), to aggregation pheromones and food Volatiles. Micronesica 37(1): 57-68.

Napompeth, B., Nishida, T., and Mitchell, W.C. 1972. Biology and rearing methods of the New Guinea sugarcane weevil Rhabdoscelus obscurus. Technical Bulletin 85.

Reddy, G.V.P., Cruz, Z.T., Bamba, J., and Muniappan, R. 2005. Development of a semiochemical- based trapping method for the New Guinea sugarcane weevil, Rhabdoscelus obscurus in Guam. Journal of Applied Entomology 129(2): 65-69.

Sallam, M.N., McAvoy, C.A., Puglisi, G.D., and Hopkins, A.M. 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.

Sallam, M.N., Peck, D.R., McAvoy, C.A., and Donald, D.A. 2007. Pheromone trapping of the sugarcane weevil borer, Rhabdoscelus obscurus (Boisduval) (Coleoptera: Curculionidae): an evaluation of trap design and placement in the field. Australian Journal of Entomology 46(3): 217-223.

Schreiner, I. 2000. New Guinea Sugarcane Weevil (Rhabdoscelus obscurus Boisduval). Agricultural Pests of the Pacific: Agricultural Development in the American Pacific.

USDA (United States Department of Agriculture). 1967. Insects not known to occur in the Continental United States: New Guinea sugarcane weevil (Rhabdoscelus obscurus (Boisduval)). Coop. Econ. Ins. Rpt. 17: 749-750.

University of Queensland. n.d. Zoology and Entomology: Rhabdoscelus obscurus. Retrieved on November 6, 2009 from: https://www.gpdd.info/references/137_xx_nd_University_Queensland_15804.pdf.

Zimmerman, E.C. 1968. Rhynchophorinae of southeastern Polynesia. Pacific Insects 10: 47-77.

Zimmerman, E.C. 1993. Australian weevils (Coleoptera: Curculionoidea). Volume III. Nanophyidae, Rhynchophoridae, Erirhinidae, Curculionidae: Amycterinae, literature consulted. CSIRO Australia and Entomological Society of America Publishing.

152 Stenchaetothrips biformis Secondary Pest of Corn Arthropods Rice Thrips

Stenchaetothrips biformis

Scientific name Stenchaetothrips biformis Bagnall

Synonyms Bagnallia biformis, Bagnallia oryza, Baliothrips biformis, Baliothrips holorphnus, Baliothrips oryzae, Chloethrips blandus, Chloethrips oryzae, Plesiothrips oh, Stenchaetothrips blandus, Stenchaetothrips dobrogensis, Stenchaetothrips dobrogensis, Stenchaetothrips oryzae, Thrips biformis, Thrips blandus, Thrips dobrogensis, Thrips holorphnus, and Thrips oryzae

Common names Rice thrips, rice leaf thrips, Oriental rice thrips, and paddy thrips

Type of pest Thrips

Taxonomic position Class: Insecta, Order: Thysanoptera, Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description Eggs: Eggs are tiny and kidney- shaped (0.25 mm long and 0.10 mm wide) and oviposited singly in tissues of the youngest leaves on the side facing the stem. New eggs are transparent and turn pale- yellow as they mature. Eggs hatch after three days.

Larvae: Neonate larvae are almost transparent. After the first molt, they turn yellowish-white (Fig. 1). The legs, head and antennae of the Figure 1. Stenchaetothrips biformis larvae. second-instar are slightly darker Photo courtesy of Rice IPM. than the first-instar larvae. Once http://www.pestalert.org/PhotoDetail.cfm?Re they begin feeding, larvae are most cordID=78. likely found on the basal parts of the leaf sheath and within very young unopened and/or more mature rolled leaves. Larvae stay on the plant in which they hatched and have three instars which, in total,

153 Stenchaetothrips biformis Secondary Pest of Corn Arthropods Rice thrips Thrips

last 6-8 days.

Prepupae: The prepupae are brown and usually found in groups with larvae and adults. Four pointed processes are found on the hind margin of the ninth tergite.

Pupae: The pupa has long wing pads reaching over two-thirds the length of the abdomen. It has four long processes on the hind margin of the ninth tergite. Pupation occurs inside rolled leaves. Prepupal and pupal stages collectively last 3-4 days.

Adults: Adult thrips are dark-brown, slender insects, about 1.25-1.35 (female) and 1 mm (male) long, with the eyes very slightly projecting (Fig. 2) (Bagnall, 1913; Williams, Figure 2. Stenchaetothrips 1916). Their antennae are 7-segmented. Adults biformis female adult with wings. can either be winged or apterous (wingless). If Note: the 7-segmented present, wings are long, narrow, fringed with antennae. Photo courtesy of fine hairs, and folded along the body when at Laurence Mound (ANIC, rest. Hindwings are paler in coloration, with CSRIO) via PADIL. distinct longitudinal vein reaching the wing tip. Adults fly during the day, and are not attracted to light. An incomplete row of short, pointed teeth is found on the hind margin of the eighth tergite. The ninth abdominal segment is very long compared to other abdominal segments.

Adult thrips may live up to 20 days and females may lay about 25 eggs in her lifetime (Nugaliyadde and Heinrichs, 1984b; Dale, 1994). While larvae stay on one individual plant during development, adults have the ability to migrate and are very mobile (Medina and Saxena, 1988).

Symptoms/Signs Stenchaetothrips biformis is considered an ‘occasional’ pest of rice, particularly during dry periods and when multiple crops of rice are planted per year, in part because of its ability to rapidly increase in population size due to its short life Figure 3. Patchy yellowing of a rice field cycle (Dale, 1994). In the vast majority of due to feeding by S. biformis. Photo outbreaks on rice, the young plants are courtesy of University of Queensland, Australia.

154 Stenchaetothrips biformis Secondary Pest of Corn Arthropods Rice thrips Thrips

attacked at the third through fifth leaf stage. It is also during this time that adult fecundity is at its highest (Nugaliyadde and Heinrichs, 1984a). Plants can generally recover from damage if water becomes available (Shepard et al., 1995). While S. biformis is a pest on rice, it is a generalist feeder on many members of the Poaceae family and has been observed on Zea Figure 4. Top: A rice leaf rolled due to S. mays (Takahashi, 1936), biformis infestation. Bottom: Unrolled rice leaf sugarcane, wild grasses, and showing damage from S. biformis feeding. Photo grassy weeds. These hosts are courtesy of University of Queensland, Australia. reported to be common reservoirs for this pest in cropping systems. Specific symptoms on corn are not available.

S. biformis larvae and adults have mouthparts that lacerate leaf tissue then puncture it to suck out the leaf sap (Dale, 1994). They feed on the growing tips of rice seedlings. Damaged leaves start showing symptoms when the tips begin to dry, then proceed to yellow or silvery streaks and translucent marks; sometimes this looks much like plants under water or nutritional stress (Figs. 3,4) (Chand and Shaw, 1975; Calora and Ferino, 1968). Both the larva and adult roll the leaf longitudinally to form a protective chamber (Fig. 4, top portion). Other symptoms include lower leaf die-off, stunting, wilting and/or scorching. In severe infestations, seedlings may be killed. Infestations can be seen in patches in fields as sickly, blighted and faded seedlings (Fig. 3) (Reyes and Rillon, 1994).

Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Survey is conducted using visual inspection of leaves for thrips injury and thrips presence. Thrips may be found in rolled up leaves. S. biformis is found on many different grasses, especially near streams, ponds and other bodies of water. Overwintering sites near water with hosts should be surveyed as well (Anon, 1976).

One sampling protocol suggests ‘wetting your hand with water and passing it over the foliage in 12 places in the nursery’; if 60 or more total thrips are captured in 12 hand passes, treatment is recommended (Tamil Nadu Ag. University, 2005).

Surveys should be focused on areas with the highest risk for establishment of S. biformis. According to a recent host analysis by USDA-APHIS-PPQ-CPHST, portions of Arkansas, Colorado, Delaware, Georgia, Idaho, Illinois, Indiana, Iowa, Kansas,

155 Stenchaetothrips biformis Secondary Pest of Corn Arthropods Rice thrips Thrips

Kentucky, Louisiana, Maryland, Michigan, Minnesota, Mississippi, Nebraska, New York, North Carolina, North Dakota, Pennsylvania, Ohio, South Carolina, South Dakota, Tennessee, Texas, and Wisconsin have a moderate level of risk for this pest based on host presence. The Sacramento Valley of California is also at the same risk level.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Stenchaetothrips species differ from other thrips genera commonly found on grasses in having 7-segmented antennae and the pronotum lacking long setae at the anterior margin (Fig. 2).

Because the symptoms of thrips infestation are similar to other problems (nutritional deficiencies, water stress), close examination of the young leaves are necessary. Inward leaf-rolling is an important characteristic of S. biformis.

In North America, at least three other thrips species may be confused with this pest. In contrast to the 7-segmented antennae for S. biformis, Chirothrips spp. have 8- segmented antennae with the second segment prolonged laterally as a tooth (Fig. 5, top), Anaphothrips obscurus has 9-segmented antennae (Fig. 5, bottom), and Frankliniella tenuicornis has 8-segmented antennae, the pronotum with long setae at the anterior margin, and the forewing with two complete rows of setae (Fig. 6) (CABI 2007).

Figure 6. Frankliniella tenuicornis with 8- segmented antennae, pronotum with long setae at the anterior margin and Figure 5. Top: Chirothrips spp. adult the forewing with two with 8-segmented antennae and with complete rows of setae. the tooth-like second segment. Photo courtesy of Jack T. Bottom: Anaphothrips obscurus with Reed, Mississippi State 9-segmented antennae. Photos Universit. courtesy of John W. Dooley, USDA-156 www.bugwood.org APHIS-PPQ. www.bugwood.org.

Stenchaetothrips biformis Secondary Pest of Corn Arthropods Rice thrips Thrips

References Anon. 1976. Further studies on the paddy thrips in Fukien Province. Acta Entomologica Sinica 19(4): 393-400.

Bagnall, R.S. 1913. Further notes on new and rare British Thysanoptera (Terebranchia) with descriptions of new species. Journal of Economic Biology 8: 231-240.

CABI. 2007. Crop Protection Compendium. Commonwealth Agricultural Bureau. http://www.cabicompendium.org.

Calora, F.B. and Ferino, M.P. 1968. Seasonal fluctuation of stemborers, thrips and leaf folders of rice in the Philippines. Philippine Entomologist 1(2): 149-160.

Chand, P. and Shaw, S.P. 1975. Incidence of thrips on paddy in Ranchi. Oryza 12(1): 51.

Dale, D. 1994. Insect pests of the rice plant – Their biology and ecology. In: Heinrichs EA, ed. Biology and Management of Rice Insects. Wiley Eastern Limited and New Age International 430-432.

Medina, E.B. and Saxena, R.C. 1988. Resistance of Oryza spp. to rice thrips, Stenchaetothrips biformis (Bagnall) (Thysanoptera: Thripidae). IRRI Seminar. Manila, Philippines.

Nugaliyadde, L. and Heinrichs, E.A. 1984a. Resistance of Oryza spp. to thrips, Stenchaetothrips biformis (Bagnall) (Thysanoptera: Thripidae). Crop Protection 3(3): 305-313.

Nugaliyadde, L. and Heinrichs, E.A. 1984b. Biology of rice thrips, Stenchaetothrips biformis (Bagnall) (Thysanoptera: Thripidae) and a greenhouse rearing technique. Journal of Economic Entomology 77(5): 1171-1175.

Reyes, C.P. and Rillon, G.S. 1994. Survey of thrips fauna and their predators on irrigated, transplanted rice. Philippine Entomologist 9(3): 268-285.

Shepard, B.M., Barrion, A.T., and Litsinger, J.A. 1995. Rice-feeding insects of tropical Asia. Los Banos, Laguna, Philippines: International Rice Research Institute.

Takahashi, R. 1936. Thysanpotera of Formosa. Philippine Journal of Science 60(4): 427-458.

Tamil Nadu Agricultural University. 2005. Crop Protection Guide, pg. 34.

Williams, C.B. 1916. Thrips oryzae, sp. Nov., injurious to rice in India. Bulletin of Entomological Research 6: 354-357.

157 Achatina fulica Secondary Pest of Corn Mollusk Giant African Snail Snail Mollusks

Primary Pests of Corn (Full Pest Datasheet)

None at this time

Secondary Pests of Corn (Truncated Pest Datasheet)

Achatina fulica

Scientific Name Achatina fulica Bowdich

Synonyms: Lissachatina fulica

Common Names Giant African snail, African giant snail, and Kalutara snail

Type of Pest Mollusk

Taxonomic Position Class: , Order: , Family: Achatinidae

Reason for inclusion in manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description Achatina fulica is distinctive in appearance and is readily identified by its large size and relatively long, narrow, conical shell (Fig. 1, 2).

Eggs: Elyptical, about 4 mm by 5 mm in diameter, usually pale yellow, laid in clutches of 100-400 (Fig. 3) (USDA, 1982).

Juveniles: Similar to adults, but have a Figure 1. The large size of the giant thinner, more brittle, translucent shell. African snail. Photo courtesy of USDA- Upon emergence, the juvenile shell is APHIS.

158 Achatina fulica Secondary Pest of Corn Mollusk Giant African Snail Snail

approximately 4 mm long (Denmark and Poucher, 1969; USDA-APHIS, 2005). Increase at a rate of 10 mm per month for first four months. The coloration is similar to adults. The columella is truncated.

Adults: Columella abruptly truncate (Burch, 1960). Columella and the parietal callus are white or bluish-white with no trace of pink (Bequaert, 1950). Shell size may be up to 8 inches (203 mm) in length and almost 5 inches (127 mm) in maximum diameter (Bequaert, 1950). Shell has seven to nine whorls and rarely as many as ten whorls (Bequaert, 1950). Shell color is reddish-brown with light yellowish, vertical (axial) streaks; or, light coffee colored. Protoconch is not bulbous. Body coloration can be either mottled brown or more rarely a pale cream color. The truncated columella is evident throughout the lifespan of the snail. The columella is generally Figure 2. Adult giant African snail. Photo concave. Snails with a lesser courtesy of Matt Ciomperlik, USDA-APHIS- concaved columella tend to be PPQ somewhat twisted. Snails with a broader shell tend to have a more concave columella (Bequaert, 1950). In calcium-rich areas, the shells of the adults tend to be thicker and opaque (USDA-APHIS, 2005).

The giant African snail, Achatina fulica, is a polyphagous pest. This species is one of the most serious land snail pests known, reported to consume all growth stages of vegetables, cover crops, garden flowers, herbaceous ornamentals, and damaging many fruit and ornamental trees (USDA, 1982). Its preferred food is decayed vegetation and animal matter, lichens, algae and fungi. The bark of relatively large trees such as citrus, papaya, rubber and cocoa is also subject to attack. Poaceous crops (sugarcane, maize, rice) suffer little or no damage from this species. There are reports of A. fulica feeding on more than 500 species of plants (CABI, 2007).

A large infestation presents a nuisance problem Figure 3. Giant African snail with slime trails, excretions, and odors of decay eggs. Courtesy of USDA APHIS. when they die in large numbers. A. fulica has been shown to be a health hazard by transmitting the rat lungworm, Angiostrongylus

159 Achatina fulica Secondary Pest of Corn Mollusk Giant African Snail Snail

cantonensis, which causes eosinophicillic meningoencepahlitis in humans. A. fulica has also been implicated in transmitting the following plant pathogens: Phytophthora palmivora on commercial pepper, coconut, betel pepper, papaya, and vanda orchid; Phytophthora colocasiae on taro; and Phytophthora parasitica on eggplant and tangerine (USDA, 1982).

A. fulica, believed to be originally from East Africa, has become established throughout the Indo-Pacific Basin, including the Hawaiian Islands. This mollusk has also been introduced to the Caribbean islands of Martinique and Guadeloupe. Recently, the snails were detected on Saint Lucia and Barbados. Although many introductions are accidental via cargo or ships, some introductions were purposeful. The market for this snail species as food is expanding. In Africa and Asia, the medicinal properties of these snails are also being investigated. The U.S. Department of Agriculture has recently discovered and confiscated illegal giant African land snails from commercial pet stores, schools, and one private breeder. These snails were being used for science lessons in schools by teachers who were unaware of the risks associated with the snails and the illegality of possessing them. In 1966, a Miami boy smuggled three giant African land snails into the country. His grandmother eventually released them into a garden, and in seven years, there were more than 18,000 of them. The Florida state eradication effort took 10 years at a cost of $1 million.

Symptoms/Signs Information specific to corn is not available. A. fulica is easily seen due to its large size, and attacked plants exhibit extensive rasping and defoliation. The weight of the number of snails on a plant can break the stems of some host species. A. fulica can also be detected by signs of ribbon-like excrement, and slime trails on plants and buildings.

Ornamentals of a number of varieties and vegetables at all stages of development are eaten by A. fulica. Cuttings and seedlings are the preferred food items. Young snails up to about 4 months feed almost exclusively on young shoots and succulent leaves. The bark of relatively large trees such as citrus, papaya, rubber and cocoa is also subject to attack. In these plants, damage is caused by complete consumption or removal of bark. The papaya appears to be the only fruit that is seriously damaged by A. fulica, largely as a result of its preference for fallen and decaying fruit (CABI, 2007).

Survey CAPS-Approved Method: Visual survey is the method to survey for A. fulica.

Literature-Based Methods: Visual survey: The most effective method of survey for mollusks is through visual searching methods. Baited traps have been used in the past but are not effective for tropical species, such as the achatinids. A. fulica is a large and conspicuous crop pest that hides during the day. Surveys are best carried out at night using a flashlight, or in the morning or evenings following a rain event.

160 Achatina fulica Secondary Pest of Corn Mollusk Giant African Snail Snail

Surveys should occur in areas that are at greatest risk for establishment of A. fulica. A recent risk analysis by USDA-APHIS-PPQ-CPHST shows that portions of Alabama, Arizona, Arkansas, California, Florida, Georgia, Louisiana, Mississippi, New Mexico, North Carolina, Oklahoma, South Carolina, Texas, and Virginia are at low to moderate risk from A. fulica. Risk of A. fulica establishment is either low or unlikely in other parts of the continental United States based on climate and host availability. Detailed survey information is available in the New Pest Response Guidelines available at http://www.aphis.usda.gov/import_export/plants/manuals/emergency/downloads/nprg_g as.pdf.

When using visual inspection methods for detection surveys an amount of bias and variation in sampling intensity is possible. In an effort to standardize sampling efforts, and thus approach a higher level of confidence in results, the following factors should be considered:

Seasonality: Conduct detection surveys on an ongoing basis, with repeated visits at the beginning, during, and/or just after the rainy season. Keep in mind that Achatina fulica remains active at a range of 9-29°C (48-84 °F). Achatina fulica begins hibernating at 2°C (35°F), and begins aestivation at 30°C (86 °F).

Time of Sampling: Plan surveys for early morning and overcast days. Achatinids are active on warm nights, early mornings, and overcast and rainy days. To maintain a consistent sampling time, conduct surveys in the early morning. On overcast days, conduct additional surveys throughout the day.

Micro Habitats: During the day, find snails in the following moist micro habitats: near heavily vegetated areas; under or near rocks and boulders; under discarded wooden boards and planks, fallen trees, logs, and branches; in damp leaf litter, compost piles, and rubbish heaps; under flower pots and planters; on rock walls, cement pilings, broken concrete, or grave markers; in gardens and fields where plants have been damaged by feeding snails and ; and at the base of the plants, under leaves, or in the “heart” of compact plants, such as lettuce or cabbage.

Evidence: While conducting a survey, look for the following clues that suggest the presence of snails: chewing damage to plants, eggs, juveniles and adults, empty snail shells, mucus and slime trails, large, ribbon-like feces, and an increase in rat population densities in an area.

Trapping: Use traps to supplement a visual inspection, if time and resources allow. Use commercial brands of bait to attract snails; however, due to the slow- acting effects of the molluscicide, these baits alone are not effective in trapping snails.

Note: Serious diseases are associated with the consumption and improper handling of certain mollusks (snails and slugs). Of particular concern, many mollusk species serve as intermediate hosts of nematodes and trematodes. While most cases of human

161 Achatina fulica Secondary Pest of Corn Mollusk Giant African Snail Snail

infections result from consumption of raw or partially cooked snail meat, government inspectors, officers and field surveyors are at-risk due to the handling of live snail, samples, and potential exposure to mucus secretions. Wear neoprene gloves when handling mollusks and wash hands thoroughly after any mollusk survey or inspection activities.

Key Diagnostics/Identification CAPS-Approved Method: Confirmation of A. fulica is by morphological identification. Identification should be verified by a malacologist at National Identification Services.

Literature-Based Methods: A. fulica is distinctive in appearance and is readily identified by its large size and relatively long, narrow, conical shell. Reaching a length of up to 20 cm the shell is more commonly in the range of 5 to 10 cm. See identification section in USDA-APHIS (2005).

References

Bequaert, J.C. 1950. Studies in the Achatinidae, a group of African Snails. Bulletin of the Museum of Comparative Zoology at Harvard College 105: 1-216.

Burch, J.B. 1960. Some snails and slugs of quarantine significance to the United States. ARS 82-1. Washington, DC: USDA–PPQ–Agricultural Research Service.

CABI. 2007. Crop Protection Compendium Wallingford, UK: CAB International. www.cabicompendium.org/cpc.

Denmark, H.A. and Poucher C. 1969. Giant African Snail in Florida. Leaflet no. 4. Gainesville: Florida Department of Agriculture and Consumer Services, Division of Plant Industry.

USDA. 1982. Pests not known to occur in the United States or of limited distribution, No. 22: Giant African Snail. USDA-APHIS-PPQ.

USDA-APHIS. 2005. New Pest Response Guidelines. Giant African Snails: Snail Pests in the Family Achatinidae. USDA–APHIS–PPQ–Emergency and Domestic Programs–Emergency Planning, Riverdale, Maryland.

162 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug

Arion vulgaris (A. lusitanicus)

Scientific Name Arion vulgaris Moquin-Tandon

Taxonomic note: In European malacological literature, A. vulgaris is known as A. lusitanicus (Anderson, 2005; Quinteiro et al., 2005). Some believe that A. lusitanicus should refer to an endemic Arion species in the Iberian Peninsula and A. vulgaris should refer to the widespread synanthropic pest (Anderson, 2005; Quinteiro et al., 2005).

Synonyms:

Common Names Iberian slug, killer slug, Lusitanian slug, murder slug, and Spanish slug

Type of Pest Slug

Taxonomic Position Class: , Order: , Family:

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description Arion vulgaris is polyphagous, feeding on a range of crop species, dumped plant material and carcasses (Whittenberg, 2005). Kozlowski (2005) found that this pest fed on at least 103 different species. Throughout most of Europe, this slug is a serious invasive pest in agricultural and forest settings (Barr et al., 2009). It is considered one of the most important slug pests in Europe due to damage to horticultural plants and cultivated agricultural crops (Rabitsch, 2006). It can damage both cultivated and uncultivated plants and can be a pest in field grown strawberries and vegetables Figure 1. A. vulgaris juveniles. Photo courtesy (Weidema, 2006). Damage to many of Inger Weidema. agricultural host crops, like oilseed rape, is usually restricted to the first few meters along the crop border (Frank, 1998a,b),

163 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug

usually when they border unmanaged field margins and wildflower strips (Frank 1998b; Friedeli and Frank, 1998). However, this slug is a bigger pest of home gardens and horticultural plants (Speiser et al., 2001a).

Unintentional introductions can occur through humans in the plant trade. Movement of potted plants and soil can transport eggs and small slugs (Weidema, 2006). This pest began spreading through human-made habitats but now is increasingly invading (semi) natural habitats while outcompeting indigenous slugs (Rabitsch and Essl, 2002). Figure 2. A. vulgaris adult. Photo The most commonly spread life stages are courtesy of Lubos Kolouch, Czechia. eggs and small juveniles. A negative impact www.bugwood.org. on , as well as local ecosystems, can occur due to competition, introgression, and outbreeding depression (Grimm and Paill, 2001). Populations of native slugs can be displaced due to A. vulgaris’ large size and high population densities (Rabitsch, 2006).

Eggs: The eggs of A. vulgaris are white, slightly transparent and have a soft shell with a diameter of approximately 2 mm. The eggs are clustered in groups of 20-200 in soil cavities (Briner and Frank, 1998b; CABI, 2007). A. vulgaris eggs are similar to other slug eggs (CABI, 2007).

Juveniles: Juveniles can have darker bands along their sides (Weidema, 2006). The body color of juvenile A. vulgaris (referred to as lusitanicus by this source) is light or dark brown (Fig. 1), often yellowish, greenish, grayish, or reddish (CABI, 2007). Juveniles up to approximately 2 cm have two dark, lateral bands. Later in the life cycle, these bands are gradually lost. Hatchlings are approximately 5 mm long when stretched out (CABI, 2007).

Adults: A. vulgaris adults (Fig. 2, 3) range in size from 6-12 cm and weigh around 5-15 grams (CABI, 2007). A. vulgaris sizes can vary between autumn and spring hatched specimens (CABI, 2007). Color can be variable ranging from dark brown, red or yellow, but is most commonly brown (Weidema, 2006). Unlike juveniles, adults usually do not have bands and are more uniform in color (Weidema, 2006). The respiratory hole () is found on the right side of the slug on the front half of the mantle (Whittenberg, 2005). The foot of A. vulgaris has a dark brown tread and looks like it has been sewn onto the body (Weidema, 2006).

Biology and Ecology A. vulgaris is active in spring and summer, even when moisture levels are low (Speiser et al., 2001a). A. vulgaris prefers moist habitats (deciduous forests, grasslands, parks and gardens) and is found mainly in cultural habitats (Weidema, 2006). Its native

164 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug

habitat is broadleaved deciduous woodland, but it can invade “regular or recently cultivated agricultural, horticultural and domestic habitats” (Rabitsch, 2006). Briner and Frank (1998a) states that the pest prefers open areas. To avoid desiccation, slugs can burrow deep into soil in dry weather (NPAG, 1999; van Dinther, 1973).

Figure 3. A. vulgaris adults. Photo courtesy of Lubos Kolouch, Czechia. www.bugwood.org. A. vulgaris’ life cycle is approximately one year with adults dying in autumn after reproduction (Weidema, 2006). Mating usually occurs in spring with 1-2 generations per year (Rabitsch, 2006). This pest is hermaphroditic so all individuals can lay eggs; outcrossing is most common (Foltz et al., 1982). Individuals can lay approximately 400 eggs in batches of 20-30 in soil crevices or compost heaps (Anonymous, 1999). Eggs are laid in late summer and in the autumn (NPAG, 1999). Eggs incubate from 3.5 to 5 weeks before young slugs hatch (Weidema, 2006). A. vulgaris can either overwinter in either the egg stage or as juveniles (CABI, 2007).

Kozlowski (2001) states that A. vulgaris has two distinct phases, an activity phase composed of crawling, feeding and copulation and a resting phase. The behavior is determined by various atmospheric factors including light intensity, rainfall and presence of dew (Kozlowski, 2001). The activity phase is interrupted by short resting periods in which the slug remains motionless, but is not numb as in the resting phase (Kozlowski, 2001).

When resting, the body of A. vulgaris is contracted with tentacles retracted. A. vulgaris also contracts when disturbed. When moving, A. vulgaris secretes a thick, shiny mucus trail (CABI, 2007). Smaller A. vulgaris spend the day in the soil; while larger specimens are found in dense vegetation or under other debris (CABI, 2007). A. vulgaris can be active during the day if the weather is cloudy and wet or air humidity and ground moisture is high (Kozlowski, 2001). Although newly sown fields are not ideal habitats, A. vulgaris can cause extensive damage if they immigrate into them from neighboring

165 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug habitats. They may migrate several meters into the field feeding during the night and return to their neighboring habitat in the morning (CABI, 2007).

Symptoms/Signs A. vulgaris can cause extensive damage to a variety of vegetables, ornamentals, and herbs by defoliating or completely devouring plants. They can also damage leaves, flowers and fruit by feeding holes (CABI, 2007). According to CABI (2007) different symptoms can be found on different crops. Symptoms by affected plant part are: visible frass on fruits or pods, leaves and/or stems; abnormal shape in fruits or pods; and external feeding signs on fruits and/or pods, leaves, roots and/or stems.

When determining if A. vulgaris has caused damage, look for obvious, shiny mucus trails. Field damage only occurs in the first few meters of the field edge and is usually close to dense, undisturbed vegetation. Large plant parts can be completely eaten but no belowground damage will occur (CABI, 2007).

This pest can transmit plant pathogens (Rabitsch, 2006) and can also harbor several parasites including (Guihon and Cens, 1973), Choanotaenia crassiscolex (Jourdane, 1972) and Choanotaenia estravarensis (Jourdane, 1972). This pest can transmit tapeworm species to livestock, and feces of A. vulgaris have been found to contain Alternaria species, Fusarium species, and Phytophthora species (Godan, 1983).

Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: In 1998, this pest was found in the United States at Cornell University (New York) (listed as A. lusitanicus) (NPAG, 1999). The current distribution of this mollusk in the United States is unknown.

The following detection survey guidelines are taken from the New Pest Response Guidelines (NPRG) (2008): “Temperate Terrestrial Gastropods”. In general, these methods are followed when a pest is known to occur.

Visual survey: The survey site should be a high risk area such as any direct points of introduction (rail yards, container yards, etc.) or known habitats (moist vegetated areas). Once the site has been identified, perform a visual inspection. This is the most effective slug and snail survey method.

Two types of visual surveys can be performed (line and plot). Line surveys are transects across a target property that allows the surveyor the flexibility to choose inspection points likely to shelter the pest. “To conduct a line survey, examine microhabitats that include vegetation, duff, and structures that might serve as diurnal or seasonal refuge sites for the pest. A minimum of one hour should be spent surveying per two-acre site. Plot surveys are small, defined areas used to conduct detailed,

166 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug

standardized subsampling throughout a target site. This method is effective for detecting immature and minute pest species. To begin surveying, randomly toss the template into the habitat. Stand outside of the plot while examining rocks, boards, litter, vegetation, and other structures within the plot. Look under leaves, duff, and at the base of plants. Spend a standardized minimum amount of time surveying each plot. Repeat four times per acre of target property. If the target property is small (less than one acre), conduct one plot sample and then line survey the entire property.”

A. vulgaris is nocturnal and easier to survey when it rains or when it is twilight (Weidema, 2006). It becomes more active as temperatures decline, so survey should occur during morning and/or evening when dew falls (Weidema, 2006). It should be noted that this pest prefers sown to naturally occurring wildflowers (Briner and Frank, 1998a).

Trapping: Trapping may be used as a supplement to visual surveying. Various methods for trapping slugs have been suggested and range from homemade tins with beer to ingeniously constructed, expensive devices (Speiser et al., 2001a, Hagnell et al., 2006). Beer can serve as an attractant for adults (Speiser et al., 2001b).

There are two main types of traps that can be used to help survey for slugs. Platform traps are artificial diurnal refuges for the pest. Platform traps are square cardboard or wood sheets, placed directly on the ground. Since the adults of this pest are rather large, the platform can be elevated slightly off the ground (NPRG, 2008). Baited traps include a food attractant inside a container that is set into the ground. Feeding attractants can include bran, molluscicide, beer or other preferred food sources.

Surveys should occur in areas that are at greatest risk for establishment of A. vulgaris. A recent host analysis by USDA-APHIS-PPQ-CPHST shows that portions of Arkansas, California, Colorado, Georgia, Idaho, Indiana, Illinois, Iowa, Kansas, Kentucky, Louisiana, Maryland, Michigan, Minnesota, Mississippi, Missouri, Nebraska, New York, North Carolina, North Dakota, Ohio, Pennsylvania, South Carolina, South Dakota, Tennessee, Texas, Washington, and Wisconsin are at moderate risk from A. vulgaris.

For more information on survey information, please refer to the New Pest Response Guidelines (2008).

Key Diagnostics CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: In Arionidae, the pneumostome (or respiration hole) is located in the anterior half on the mantle, which has a granular structure. No kneel is present in Arionidae (CABI, 20007). Identification may be achieved through dissection and genitalia examination (Noble, 1992; Hagnell et al., 2006; Barr et al., 2009).

Arion vulgaris can be confused with both A. ater and A. rufus and make up the ater- rufus-vulgaris complex (ARVC) (Barr et al., 2009). Distinguishing between these

167 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug

different slugs can become more difficult considering that they can interbreed to form hybrids between ARVC species (Schander and von Proschwitz, 2003; Hagnell et al., 2006). Identification can only be certain through dissection of the genitalia or molecular analysis (Barr et al., 2009).

Striped immatures may be mistaken for Arion subfuscus (NPRG, 2008).

A. vulgaris can also be confused with three other slug families, Agriolimacidae, Limacidae and Milacidae (CABI, 2007). Careful examination of morphological features can help distinguish between the two families.

References Anderson, R. 2005. An annotated list of the non-marine mollusca of Britain and Ireland. Journal of Conchology 38(6): 607-637.

Anonymous. 1999. Iberisk Skovsnegl, Grønviden, Havebrug 124.

Barr, N.B., Cook, A., Elder, P., Molongoski, J., Prasher, D., and Robinson, D.G. 2009. Application of a DNA barcode using the 16S rRNA gene to diagnose pest Arion species in the USA. Journal of Molluscan Studies 75: 187-191.

Briner, T. and Frank, T. 1998a. The palatability of 78 wildflower strip plants to the slug Arion lusitanicus. Annals of Applied Biology 133:123-133.

Briner, T. and Frank, T., 1998b. Egg laying activity of the slug Arion lusitanicus Mabille in Switzerland. Journal of Conchology 36: 9-15.

CABI. 2007. Crop Protection Compendium [CD]. Wallingford, UK: CAB International. http://www.cabicompendium.org/cpc.

Foltz, D.W., Ochman, H., Jones, J. S., Evangelisti, S.M., and Selander, R.K. 1982. Genetic population structure and breeding systems in arionid slugs (Mollusca: Pulmonata). Biological Journal of the Linnean Society 17: 225-241.

Frank, T. 1998a. Slug damage and number of slugs (Gastropoda: Pulmonata) in winter wheat in fields with sown wildflower strips. Journal of Molluscan Studies. 64: 319-328.

Frank, T. 1998b. Slug damage and numbers of slugs in oilseed rape bordering on grass strips. Journal of Molluscan Studies 64: 461-466.

Frideli, J. and Frank, T. 1998. Reduced applications of metaldehyde pellets for reliable control of the slug pests Arion lusitanicus and Deroceras reticulatum in oilseed rape adjacent to sown wildflower strips. Journal of Applied Ecology 35: 504-513.

Godan, D. 1983. Pest Slugs and Snails: Biology and Control. Berlin: Springer-Verlag. 445 pp.

Grimm, B. and Paill, W. 2001. Spatial distribution and home-range of the pest slug Arion lusitanicus (Mollusca: Pulmonata). Acta Oecologica, 22: 219-227.

Guilhon, J. and Cens, B. 1973. Angiostrongylus vasorum (Baillet, 1866) Etude biologique et morphologique. Annales de Parasitologie (Paris) 48(4): 567-596.

168 Arion vulgaris Secondary Pest of Corn Mollusk Iberian slug Slug

Hagnell, J., Schander, C., Nilsson, M., Ragnarsson, J., Valstar, H., Wollkopf, A.M., von Proschwitz, T. 2006. How to trap a slug: commercial versus homemade slug traps. Crop Protection, 25: 212-215.

Jourdane, J. 1972. Etude experimentale du cycle biologique de deux especes de Choanotaenia intextinaux des Soricidae. Zeitschrift fur Parasitenkunde 38: 333-343.

Kozlowski, J. 2001. Daily activity of Arion lusitanicus Mabille, 1868 (Gastropoda: Pulmonata: Arionidae). Journal of Plant Protection Research 41: 279-287.

Kozlowski, J. 2005. Host plants and harmfulness of the Arion lusitanicus Mabille, 1868 slug. Journal of Plant Protection Research 45(3): 221-233.

Noble, L.R. 1992. Differentiation of large arionid slugs (Mollusca, Pulmonata) using ligula morphology. Zoologica Scripta 21(3): 255-263.

NPAG. 1999. NPAG data: Arion lusitanicus a European slug. United States Department of Agriculture, Animal and Plant Health Inspection Service, Plant Protection and Quarantine, Center for Plant Health Science and Technology, Pest Epidemiology and Risk Assessment Laboratory.

NPRG. 2008. Temperate Terrestrial Gastropods. United States Department of Agriculture, Animal and Plant Health Inspection Service and Cooperating State Departments of Agriculture.

Quinteiro, J., Rodriguez-Castro, J., Castillejo, J., Inglesias-Pineiro, J., and Rey-Mendez, M. 2005. Phylogeny of slug species of the genus Arion: evidence of monophyly of Iberian endemics and of the existence of relict species in Pyrenean refuges. Journal of Zoological Systematics and Evolutionary Research 43(2): 139-148.

Rabitsch, W. 2006. Arion vulgaris. DAISIE.

Rabitsch, W. and Essl. F. 2002. Neobiota in Osterreich. Umweltbundesamt. Federal Environment Agency, Austria.

Schander, C. and von Proschwitz, T. 2003. Hybridization in Arionids: the rise of a super slug? BCPC Symposium Proceedings 80: 221-226.

Speiser, B., Zaller, J.G., Neudecker, A. 2001a. Size-specific susceptibility of the pest slugs Deroceras reticulatum and Arion lusitanicus to the nematode biocontrol agent Phasmarhabditis hermaphrodita. Biocontrol 46:311-320.

Speiser, B., Glen, D., Piggott, S., Ester, A., Davies, K., Castillejo, J., Coupland, J. 2001b. Slug damage and control of slugs in horticultural crops. EU funded research project brochure. van Dinther, J. 1973. Mollusks and their control. World Crops 25(6): 282-286.

Weidema, I. 2006. NOBANIS – Invasive Alien Species Fact Sheet –Arion lusitanicus. – From: Online Database of the North European and Baltic Network on Invasive Alien Species – NOBANIS www.nobanis.org.

Wittenberg, R. (ed.) 2005. An inventory of alien species and their threat to biodiversity and economy in Switzerland. CABI Bioscience Switzerland Centre report to the Swiss Agency for Environment, Forests and Landscape. 240-241.

169 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode Nematodes

Primary Pests of Corn (Full Pest Datasheet)

Punctodera chalcoensis

Scientific Name Punctodera chalcoenis Stone, Moss, and Mulvey

Synonyms: Heterodera punctata

Common Name(s) Mexican corn cyst nematode

Type of Pest Nematode

Taxonomic Position Class: , Order: , Family:

Reason for Inclusion in Manual National cyst nematode survey

Pest Description Cyst nematodes that are now considered Punctodera have been recognized for 60 years, primarily on the basis of a cyst without a cone but with both a vulval circumfenestra and an anal fenestra. Females and cysts of Punctodera spp. are globose to ovoid tapering to a protruding neck (Hesling, 1978). Subsurface punctations of the female cuticle, while indicated by the genus name, also occur in other heteoderines, although their intensity and occurrence in parallel rows is striking in Punctodera.

From Baldwin and Mundo-Ocampo (1991): The Mexican corn cyst nematode, Punctodera chalcoensis, was first observed by Vasquez (1976) in corn fields in Huamantla, Tlaxcala, Mexico in the late 1950’s and was considered Heterodera punctata (=Punctodera punctata). In the early 1960’s, a cyst nematode was observed attacking corn in the valley of Mexico at Chalco and subsequently in the states of Puebla and Tlaxcala, which was also identified as H. punctata (Sosa-Moss and Gonzalez, 1973; Vazquez, 1976). Sosa-Moss (1965) believed that the Chalco population of the Mexican corn cyst nematode, unlike H. punctata, was limited to corn as a host. In 1976, a new genus, Punctodera, was proposed to accommodate P. punctata and a new species from Canada, P. matadorensis (Mulvey and Stone, 1976). Meanwhile, morphological differences were noted when comparing the Mexican corn

170 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

cyst nematode with H. punctata populations from other countries (Sosa-Moss, 1965; Villanueva, 1974) and with P. matadorensis (Stone et al. 1976). These observations led to description of the Mexican corn cyst nematode as a distinct species, P. chalcoensis, with the type locality in Chalco, Mexico and the corn as the type host (Stone et al., 1976).

From Stone et al. (1976): Note: Measurements are from cysts collected from field soil; field specimens were smaller than the females cultured in the glasshouse.

Eggs: Length = 114.4 + 3.6 µm; width = 43.1 + 1.7 µm; length/width = 2.7 + 0.1 µm. Second stage juvenile folded four times within the egg.

Second-stage juveniles: (Fig. 1) Typical heteroderid second-stage juvenile with vermiform body adopting a slight ventral curve on heat relaxation; tail tapering to a slender point. Cuticular annulations distinct; four incisors in lateral field reducing to three anteriorly and posteriorly, aerolated intermittently throughout. Cuticle thicker for first eight annules. Head offset, apparently three-five, usually four head annules when seen by light microscopy. SEM observation Figure 1. Second-stage juvenile. Top: Shape and shows usually three, the extra annule size of stylet knob. Bottom. Long hyaline tail seen in light microscopy being part of section. Photos courtesy of Janet A. Rowe. the lips. Oral disc distinct, elongated www.cabicompendium.org dorso-ventrally to about 1.5 times its width and surrounded by distinct lateral lips bearing the amphid apertures; the two components of each pair of submedian lips are fused into single distinct arcs. No indication of cephalic or labial papillae. Moderately heavy hexaradiate head skeleton. Cephalids at level of second and seventh-eigth body annules. Well developed stylet with massive basal knobs rounded posteriorly, flat, to shallowly concave anteriorly. Anterior (prorhabdial) portion of stylet less than half stylet length. Median oesophageal bulb well developed occupying full width of body cavity and often with a somewhat rectangular shape. Oesophageal gland lobes with ventral overlap, usually extending back about one third of body length but sometimes considerably shorter. No distinct oesophageal-intestinal valve observed. Hemizonid one-two annules anterior to

171 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

excretory pore; hemizonion not observed. Genital primordium lying about 60% of body length from anterior end. Phasmids situated two-thirds along tail length; small refractive bodies sometimes present in matrix of hyaline terminus.

Unfixed juveniles: Length = 542 + 26 µm; maximum body width = 21.6 + 0.7 µm; stylet length = 24.4 + 0.2 µm; stylet base to dorsal oesophageal gland duct junction = 5.5 + 0.8 µm; head tip to median bulb valve = 70.4 + 6.3 µm; head tip to excretory pore = 110.1 + 7.9 µm; head tip to end of oesophageal glands lobe = 176.0 + 15.1 µm; tail length = 66.2 + 3.5 µm; hyaline terminus length = 38.6 + 3.4 µm; hyaline terminus length/stylet length = 1.6 + 0.1.

Fixed juveniles: Length = 533 + 29 µm; maximum body width = 20.1 + 0.6 µm; stylet length = 24.7 + 0.6 µm; stylet base to dorsal oesophageal gland duct junction = 5.1 + 0.7 µm; head tip to median bulb valve = 73.7 + 2.7 µm; head tip to excretory pore = 107.8 + 4.9 µm; tail length= 63.2 + 3.3 µm; hyaline terminus length = 38.2 + 3.1 µm; hyaline terminus length/stylet length = 1.5 + 0.1.

Female: Mature females white in color (Fig. 2), spherical or subspherical with projecting neck containing an oesophagus; length excluding neck/width ratio close to one. Head with one or two prominent annules. Stylet slender with rounded basal knobs. Oesophagus strongly developed with massive circular median bulb with prominent valve; oesophageal glands lobe with distinct dorsal and subventral gland cells. Excretory pore at base of neck. Two large ovaries filling enlarged body cavity, which in mature females is occupied by eggs. Cuticle greatly Figure 2. P. punctodera females (white) and cyst thickened except in head region, (brown) on corn roots. Photo courtesy of Janet A. covered with a rugose or lace-like Rowe. www.cabicompendium.org pattern of shallow ridges, with a

sub-surface pattern of rows of fine refractive spots (the “punctations”).

Vulva a short transverse slit situated at the opposite pole of the body to the neck, lying centrally in a circular zone lacking ridges and punctations, with the cuticle of reduced thickness (the vulval fenestra). Vulval slit on sight circular prominence. The vulval fenestra appears more transparent than the surrounding body wall and is itself surrounded by a narrow zone of less thick cuticle. Anus, a transverse slit smaller than the vulva and lying ‘dorsal’ to the vulva within a thin-walled circular zone (the anal

172 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode fenestra). The anal fenestra is similar in size and appearance to the vulval fenestra but lacks the surrounding clear zone; the anus does not lie centrally in the fenestra but towards the ventral side. Clusters of elongate bodies are associated with the anal and vulval apertures. The size of the vulval aperature precludes extrusion of eggs but a very small gelatinous matrix (egg sac) was observed on some specimens and may have been lost from other specimens in extraction. A thick white sub-crystalline layer is typically present consisting of polygonal plates and resembling that described from P. punctata.

Length excluding neck = 473 + 105 µm; width = 429 + 120 µm, neck length = 142 + 24 µm; length excluding neck/width ratio =1.14 + 0.18; stylet length = 25.8 + 0.9 µm, stylet base to dorsal oesophageal gland duct junction = 5.7 + 0.9 µm; length of median oesophageal bulb = 25.7 + 1.9 µm; width of median oesophageal bulb = 24.9 + 2.4 µm; head tip to median oesophageal bulb valve = 77.4 + 8.4 µm; head tip to excretory pore = 131.0 + 0.6 µm.

Perineal portions of females: Vulval fenestra length = 30.7 + 7.0 µm; width 32.1 + 7.0 µm; length of vulval slit = 4.0 + 0.5 µm; anal fenestra length = 29.5 + 5.7 µm; width = 31.1 + 5.3 µm; length of anal slit = 3.0 + 1.0 µm; distance between fenestrae = 67.7 + 14.5 µm.

Cysts: Cyst shape as that of female, color pale to dark brown (Fig. 2), darkening with age. New cysts often retain the subcrystalline layer. In old cysts, the thin walls of the vulval and anal fenestrae are lost; younger cysts show incomplete fenestration. Some specimens have small scattered bullae in the perineal region or closely sited in an area just below the vulval fenestra but they are lacking from many cysts. New cysts typically contain from 200-400 embryonated eggs.

Cysts entire, length excluding neck = 510 µm; maximum width = 470 µm; neck length = 88 µm; length excluding neck/width ratio =1.1. Dry cyst length excluding neck = 441 + 69 µm; width = 416 + 61 µm; neck length = 95 + 26 µm; length excluding neck/width ratio= 1.06 +0.10.

Perineal portions of cysts: Vulval fenestra length = 18.1 + 2.7 µm; width 19.8 + 2.9 µm; diameter of clear zone surrounding vulval fenestra ~ 30 µm; length of vulval slit = 4.2 + 0.4 µm; anal fenestra length = 21.1 + 3.4 µm; width = 22.4 + 3.1 µm; length of anal slit = 2.8 + 0.4 µm; distance between fenestre = 142.3 + 8.8 µm.

Males: Typical heteroderid male morphology. Body vermiform, heat relaxed specimens with strong ventral curvature and tail frequently twisted through 180°. Head offset with 5-7 annules as seen by light microscopy. Tail bluntly rounded less than one quarter of body width long. Cuticle with regular annulations, four lateral incisures, areolated, lateral field terminating on tail. Heavy hexaradiate cephalic skeleton. Cephalids at level of second and eighth body annules. Stylet well developed with shallow basal knobs, flat to slightly concave anteriorly. Median oesophageal bulb ellipsoidal, not filling body cavity; oesophageal glands lobe overlapping intestine ventrally. Hemizonid extending over two

173 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

annules, one annule anterior to excretory pore, hemizonion not observed. Single gonad extending for about half body length, less in small specimens. Paired spicules flask- shaped proximally and tapering distally, curved ventrally with single points, single un- ornamented gubernaculums. Phasmids not observed.

Length = 985 + 69 µm; body width = 25.8 + 1.7 µm; stylet length = 26.8 + 0.8 µm; stylet base to dorsal oesophageal gland duct junction = 4.0 + 0.6 µm; head tip to oesophageal median bulb valve = 82.7 + 3.7 µm; head tip to excretory pore = 131.3 + 7.7 µm; tail length = 2.8 + 1.4 µm; spicule length (across chord) = 31.9 + 1.8 µm; gubernaculum length 7.3 + 0.8 µm; overall gonad length 476 + 107 µm; G=48.

Biology and Ecology This cyst forming nematode has sedentary endoparasitic habits. Cysts are persistent (>10 years) tanned sacs derived from the female body and contain eggs. Cysts of this species are subspherical or ovoid, lack posterior protuberance, have large vulval and anal fenestrae and small scattered or absent bullae(Sosa Moss, 1987). Second-stage juveniles emerge from cysts, penetrate host roots and establish a specialized feeding site (syncytium) in the stele. They develop becoming swollen females, which retain the eggs, rupture the root cortex and protrude from the root Figure 3. P. punctodera damage to corn. Photo surface. At the end of the courtesy of Laurence I. Miller, Virginia Tech. reproductive phase, females University. www.forestryimages.org die and become dark brown cysts (Society of Nematologists, n.d.).

Several authors concluded that the life cycle of P. chalcoensis spans the growing season of corn, so that only one generation occurs per year and that the cysts must remain in the soil during the winter to initiate the next cycle (Baldwin and Mundo- Ocampo, 1991). Others have reported a typical heteroderine life cycle of about 30 days (Baldwin and Mundo-Ocampo, 1991). Apparently fresh eggs may hatch readily, but once diapause is established within cysts, only a low percentage of hatch occurs each season. Villanueva (1974) reported that root exudates of most plants induce egg hatching of the Mexican population, but that exudates from corn roots resulted in a significantly greater degree of hatching.

174 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

The environmental requirements of P. chalcoensis are unknown, although the distribution at high elevations (above 2000 meters) in Mexico, and its absence in corn fields in warmer, subtropical regions suggests that it does not tolerate continuous, warm, humid conditions. The nematode is most abundant in sandy soils (Stone et al., 1976).

Symptoms/Signs P. chalcoensis is highly damaging to corn in Mexico, causing severe yellowing, stunting (Fig. 3), and even death of young seedlings. Pale stripes on the leaves can be observed. Plants without supplementary fertilization showed more damage and less tolerance to the nematode. Heavily attacked plants have stunted root systems and many short laterals, giving a bottle brush effect and the aerial parts of the plants appear unthrifty.

Later sown corn suffers greater damage than early sown corn because the peak emergence of infective juveniles of P. chalcoensis occur at the same time as seed germination (Sosa-Moss, 1987). Early sown corn has already developed a good root system before the rains provide sufficient moisture to stimulate hatching of juveniles.

White females can be observed on roots of corn (Fig. 2).

Pest Importance P. chalcoensis is a serious pest of corn in Mexico. When associated with fungi, P. chalcoensis can cause yield suppression of 90% (Sosa-Moss, 1987). The plants that remain are so stunted and yellow that no straw can be harvested.

Known Hosts Zea mays (corn) and Zea mexicana (teosinte, wild maize).

Graminaceae tested and proved to be non-hosts were: Avena sativa (oats), A. fatua (wild oat), Triticum aestivum (wheat), Secale cereale (rye), triticale, Sorghum vulgare (sorghum), Agropyron spp. (crested wheatgrass), Bromus spp. (brome), Dactylis spp. (orchard grass), Elymus spp. (wild rye), Festuca spp. (fescue), Lolium spp. (ryegrass), and Phleium spp. (timothy grass).

Known Vectors (or associated organisms) P. chalcoensis is not a known vector, but it attacks the roots of the plant, damaging its nutrition and allowing other pathogens (fungi and viruses) access to the plant.

Known Distribution North America: Mexico

Potential Distribution within the United States Biological information for this pest is not currently available to determine the potential distribution of this nematode within the United States. Once a cyst nematode is introduced into a country, however, it is very difficult to minimize spread without

175 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode extensive quarantine measures and expensive eradication strategies. Cyst nematodes are efficiently disseminated by all means of soil movement, including minute amounts of soil that contaminate equipment, by and plant products, and by soil that is moved by water and wind.

A recent host analysis by USDA-APHIS-PPQ-CPHST shows that portions of Arkansas, Idaho, Illinois, Indiana, Iowa, Kansas, Kentucky, Louisiana, Maryland, Michigan, Minnesota, Mississippi, Missouri, Nebraska, New York, North Dakota, Pennsylvania, Ohio, South Dakota, Tennessee, Texas, and Wisconsin have the greatest risk based on host availability. Corn cultivars grown in the United States are susceptible to P. chalcoensis (Mundo et al., 1987).

Survey

CAPS-Approved Method: Use either soil sampling or collection of host roots or of a combination of both methods.

Soil sample: send sample to a nematology diagnostic lab where nematodes will be extracted from the soil and identified (preferred method). Sieving, Fenwick-can, and modified Fenwick-can are the best techniques for extraction of cysts and juveniles.

Collect host roots: Send sample to nematology diagnostic lab where nematodes will be extracted and identified.

Literature-Based Methods: Soil sampling: The soil around corn plants with cysts should be sampled. Cyst nematodes are then extracted from soil, identified, and counted. A composite soil sample is collected from a sampling unit such as an entire field, a specific area of a field, or an experimental plot. Multiple (15 to 20) cores of soil are collected from the upper 8 to 12 inches of soil and combined for each composite sample. A subsample is removed from the larger sample and air-dried.

Cysts can be extracted from the subsample using a Fenwick can method (20 mesh screen over a 60 mesh screen), a modified Baermann funnel technique, or a modified Fenwick can elutriation method (Fig. 4a) with further separation of cysts from the plant debris by flotation in an ethanol and glycerin solution (Fig. 4b) (Caswell et al., 1985; Ingham, 1994). Cysts are then picked from the remaining debris and identified. Cysts can be crushed to determine the number of eggs plus juveniles, which can then be adjusted to reflect the nematode density per pound of oven-dry soil.

Survey for P. punctodera should crossover easily and inexpensively if a state already has equipment to survey for potato cyst nematodes, because the Fenwick can method used to survey for Globodera pallida and G. rostochiensis should also work for the extraction of P. punctodera cysts.

176 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

A

B

Figure 4. A) Modified Fenwick can used to separate nematode cysts and organic debris from soil sample. B) Ethanol-glycerine flotation apparatus for separation of nematode cysts from sample organic matter. Reproduced from Caswell et al., 1985.

There are a variety of additional methods available for cyst extraction from soil. Soil can also be processed using Cobb’s decanting and sieving technique to determine the presence of males, white females, and cysts. Subbotin et al. (2003) used sieving- decanting and centrifugation-flotation methods to isolate cysts from soil. Holgado et al. (2004) air dried soil samples, passed the samples through a 5 mm sieve, and extracted cysts using a fluidizing column. Abidou et al. (2005) processed soil samples through a Kort elutriator.

Motile nematodes (e.g. juveniles) can be extracted using the Whitehead tray method (Smiley et al., 2007).

Key Diagnostics CAPS-Approved Method: Confirmation of P. chalcoenisis is by morphological identification. Characteristics of second-stage juveniles, females, and cysts can be used to differentiate from other Punctodera species.

Literature-Based Methods: Most diagnoses are made via morphological characteristics. The Society of Nematologists provide an illustrated key to selected genera and species of cyst forming Heteroderidae in the Western Hemisphere (based on cysts and second-stage larvae) at http://nematode.unl.edu/cystkey.htm.

Stone et al. (1976) provide a morphological key to the three Punctodera spp.:

1. Cyst pear-shaped, second-stage juvenile 350-470 µm …………Punctodera punctata

- Cyst spherical or sub-spherical shaped, second stage juvenile >500 µm ………….2

177 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

2. Massive bullae between vulval and anal fenestrae, second stage juvenile with oesophageal gland lobe extending to about 50% of body length and stylet knob strongly concave anteriorly, host grasses ………………………Punctodera matadorensis

- Bullae small, scattered or absent, second-stage juvenile with oesophageal gland lobe extending to about 30% of body length and stylet knobs flattish to slightly concave anteriorly, host maize ……………………………………………….Punctodera chalcoenisis

Sabo et al (2002), however, sequenced the internal transcribed spacer (ITS) 1, 5.8S ribosomal RNA gene, and ITS2 of P. chalcoensis, which may provide for molecular identification.

Easily Confused Pests Punctodera chalcoensis can be confused with the two other Punctodera spp. (P. punctata and P. matadorensis), although neither occur on corn. P. punctata is distributed worldwide on turf, wheat, and weed grasses, and P. matadorensis is apparently limited to weed grasses at the type location in Canada. P. matadorensis was, however, recently detected in North Dakota in February, 2010. P. chalcoensis and P. matadorensis can be separated from P. punctata by the spherical to subspherical shape of the females and cysts in the former versus more elongate, pear-shaped females and cysts in the latter (Baldwin and Mundo-Ocampo, 1991). Cysts of P. chalcoensis differ from those of P. matadorensis by small, scattered bullae or no bullae versus massive and consistently present bullae in P. matadorensis. P. chalcoensis also differs slightly by the shape of the stylet knobs of the second-stage juvenile, which are flat to slightly concave anteriorly in P. chalcoensis versus strongly concave and anchor- shaped in P. matadorensis and rounded in P. punctata. The esophagus of the second- stage juveniles of P. chalcoensis and P. punctata occupies about 30% of the body length, whereas in P. matadorensis it includes about 50% of the body length (Baldwin and Mundo-Ocampo, 1991).

In Mexico, corn and potatoes are sometimes grown in the same areas. Globodera cysts sometime occur in the same soil sample. Globodera cysts are similar in size, shape, and color. The cyst nematode Heterodera zeae is present in India and North America on corn. Heterodera spp. have lemon-shaped cysts, so confusion should not arise between the two genera (CABI, 2007).

References Abidou, H., Valette, S., Gautheire, J.P., Rivoal, R., El-Ahmed, A., and Yahyaoui, A. 2005. Molecular polymorphism and morphometrics of species of Heterodera avenae group in Syria and Turkey. Journal of Nematology 37(2): 146-154.

Baldwin, J.G. and Mundo-Ocampo, M. 1991. , cyst- and non-cyst forming nematodes. In: Manual of Agricultural Nematology. W.R. Nickle (ed). Pp. 275-362. New York: Marcel Dekker.

CABI. 2007. Crop Protection Compendium [CD]. Wallingford, UK: CAB International. t http://www.cabicompendium.org/cpc.

178 Punctodera chalcoensis Primary Pest of Corn Nematode Mexican corn cyst nematode Cyst nematode

Caswell, E.P., Thomason, I.J., and McKinney, H.E. 1985. Extraction of cysts and eggs of Heterodera schachtii from soil with an assessment of extraction efficiency. Journal of Nematology 17: 337-340.

Hesling, J.J. 1978. Cyst nematodes: Morphology and identification of Heterodera, Globodera, and Punctodera. Tech. Bull. G B Minist. Agric. Fish Food 7: 125-155.

Holgado, R., Rowe, J.A., and Magnusson, C. 2004. Morphology of cysts and second stage juveniles of Heterodera filipjevi (Madzhidov, 1981) Stelter, 1984 from Norway. Journal of nematode morphology and systematics 7(1): 77-84.

Ingham, R.E. 1994. Nematodes. Pages 459-490. In R.W. Weaver et al. (ed). Methods of soil analysis, Part 2. American Society of Agronomy. Madison, WI.

Mulvey, R.H., and Stone, A.R. 1976. Description of Punctodera matadorensis n. gen., n. sp. (Nematoda: Hetoroderidae) from Saskatechewan with lists of species and generic diagnosis of Globodera (n. rank), Heterodera, and Sarisodera. Can. J. Zool. 54: 772-785.

Mundo, M., Baldwin, J.G., and Jeromino, R.J. 1987. Distribution, morphological variation, and host range of Punctodera chalcoensis. J. Nematology 19: 545.

Sabo, A., Reis, L.G.L., Krall, E., Mundo-Ocampo, M., and Ferris, V.R. 2002. Phylogenetic relationships of a distinct species of Globodera from Portugal and two Punctodera species. Journal of Nematology 34(3): 263-266.

Smiley, R., Sheedy, J., Pinkerton, J., Easley, S., Thompson, A., and Yan, G. 2007. Cereal cyst nematode: distribution, yield reduction, and crop management strategies. Oregon State University. Pages 15-29. http://extension.oregonstate.edu/catalog/html/sr/sr1074-e/05.pdf.

Society of Nematologists. n.d. Punctodera chalcoensis. Exotic Nematode Plant Pests of Agricultural and Environmental Significance to the United States. Assessed 2/11/10. http://nematode.unl.edu/pest29.htm.

Sosa Moss, C. 1965. Recheches sur Heterodera punctata Thorne, parasite du maiss au Mexique Comptes rendus du huiteme. Symposium International de Nematologie, Antibes, E.J. Brill, p. 54.

Sosa-Moss, C. 1987. Cyst nematodes of Mexico, Central America, and South America. Nematologica Mediterranea 15: 1-12.

Sosa Moss, C. and Gonzales, C.P. 1973. Respuesta de maiz chalqueno ferilizado y no fertilizado a 4 differentes niveles de Heterodera punctata raza Mexicana (Nematoda: Heterodidae). Nematropica 3: 13- 14.

Stone, A.R., Sosa Moss, C., and Mulvey, R.H. 1976. Punctodera chalcoensis n. sp. (Nematoda: Heteroderidae) a cyst nematode from Mexico parasitizing Zea mays. Nematologica 22: 381-389.

Subbotin, S.A., Sturhan, D., Rumpenhorst, H.J., and Moens, M. 2003. Molecular and morphological characterization of the Heterodera avenae species complex (Tylenchida: Heteroderidae). Nematology 5(4): 515-538.

Vasquez, J.T. 1976. Infesaciones de nematodos fitoparasitos como factor limitante en la produccion de maiz en el altiplano mexicano. Produccion del Departamento Mexicano. CODAGEM, pp. 79.

Villnueva, R.M. de J. 1974. Comparacion morfometrica entre uno poblacion inglesa y una Mexicana de Heterodera punctata Thorne (nematode Heteroderodae), influencia de exudados radiculares en la poblacion Mexicana. Tesis professional, I.P.N. Mexico

179 Meloidogyne fallax Secondary Pest of Corn Nematode False Columbia root-knot nematode Root-knot nematode Secondary Pests of Corn (Truncated Pest Datasheet)

Meloidogyne fallax

Scientific Name Meloidogyne fallax Karssen

Synonyms: Meloidogyne chitwoodi B-type

Common Name(s) False Columbia root-knot nematode

Type of Pest Nematode

Taxonomic Position Class: Secernentea, Order: Tylenchida, Family: Heteroderidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description Meloidogyne fallax was named for its morphological similarity to M. chitwoodi and was initially considered a new race of M. chitwoodi. M. fallax is a significant pest primarily of potato in Europe. The nematode occurs in the Netherlands, Belgium and Germany; outside of Europe, it has been reported from Australasia and South Africa.

Eggs: Length 89.7-103.6 μm; width 34.1-44.2 μm

Second-stage juveniles (J2): Vermiform, motile; 381-435.2 µm long and 13-16.4 µm wide; tail length is 46-55.6 µm with a smooth hyaline tail and broadly rounded tail terminus.

Females: pear-shaped, pearly-white and sedentary; 404-720.3 µm long and 256-464 µm wide. The stylet is dorsally curved, 13.9-15.2 μm long, with rounded to ovoid stylet knobs, slightly sloping posteriorly.

Males: Vermiform, motile; 736-1520 µm in length and 27-44 µm in width with a slight taper at each end. The stylet is 18.9-20.9 μm long, with large rounded knobs, set off from the shaft. The tail measures 7.6-12.1 µm long and is twisted. Cuticular annules are distinct.

180 Meloidogyne fallax Secondary Pest of Corn Nematode False Columbia root-knot nematode Root-knot nematode

For detailed descriptions of each life stage see Karssen, 1996; Davis and Venette, 2004.

Symptoms/Signs Corn is a reported host of M. fallax by Waeyenberge and Moens, 2001. The Society of Nematologists (n.d), however, suggest that corn is not a host of M. fallax. Little information exists on the pest importance to corn and symptoms specific to corn.

On potato, symptoms are very similar to those caused by M. chitwoodi (CABI, 2007). Aboveground symptoms are often not obvious but may consist of varying degrees of stunting, lack of vigor, and a tendency to wilt under moisture stress, all leading to reduced yield. Infested potato tubers may have small raised swellings on their surface above the developing nematodes (Fig. 1), as compared with the large easily noticeable galls of other Meloidogyne species. A number of galls may be concentrated on one area of the tuber or single galls may be scattered near eyes or lesions. Internal tissue below the gall is necrotic and brownish. Adult Figure 1. Symptoms of M. fallax on potato. Photos females are visible just below the courtesy of EPPO. surface; when alive they appear as glistening, white, pear-shaped bodies surrounded by a brownish layer of host tissue. As the female matures and dies, the egg-filled sac becomes dark-brown with age. Potato roots may also be infected, but this is difficult to detect without a magnifying lens, as little or no galling occurs, even in heavy infestations. The spherical bodies of females may protrude from the surface of small rootlets surrounded posteriorly by a large egg-filled sac, which becomes dark- brown with age.

181 Meloidogyne fallax Secondary Pest of Corn Nematode False Columbia root-knot nematode Root-knot nematode

Survey

CAPS-Approved Method: Use either soil sampling or visual sampling of tubers or of a combination of both methods.

1. Soil sample: send sample to nematology diagnostic lab where nematodes will be extracted from the soil and identified.

The presence of M. fallax in infested soil can be determined by sampling and extraction of the second-stage juveniles, using a standard nematode extraction procedure for free-living nematodes of this size or techniques used for root-knot nematodes (Baermann trays, Baermann trays with elutriation or sieving, centrifugal flotation, flotation-sieving, semiautomatic elutriation and Cobb's decanting and sieving).

2. Visual survey: collect symptomatic tubers.

Literature-Based Methods: Surveys should occur in areas with higher risk of M. fallax establishment. A recent host map developed by USDA-APHIS-PPQ-CPHST (Fig. 2) shows much of the country is at low to moderate risk for establishment based on host availability.

Detection and inspection methods are similar to those for M. chitwoodi. The presence of M. fallax in infested soil can be determined by sampling and extraction of the second- stage juveniles using a standard nematode extraction procedure for free-living nematodes of this size. External symptoms on tubers are obvious in the case of heavy infestations, but where nematode numbers are low or in the early stages of infection, such symptoms are not obvious. Clearing and staining of the tissues can show the presence of nematodes (Hooper, 1986) but this can be a laborious procedure. Storage of lightly infested tubers may lead to the development of obvious external symptoms.

Vovlas and Inserra (1996) outline general considerations for conducting a survey for a new Meloidogyne spp. in citrus orchards. In general, they recommend sampling root tissues to inspect for the presence of galled roots. They also note that soil samples may detect Meloidogyne spp., but these individuals may not be of particular concern. Many native or naturalized Meloidogyne spp. parasitize a number of weed hosts. Thus, careful examination of individuals will be necessary to confirm species identity. Samples of soil or host roots must be collected with the purpose of obtaining males, juveniles, or female nematodes within root tissues. Samples must then be processed to separate nematodes from soil and debris. Finally, nematodes must be prepared either for identification using morphological (e.g., perineal patterns) or molecular techniques.

Root-knot nematodes are extracted from soil using a variety of techniques. Six methods (and subtle variations thereof) are particularly common: Baermann trays; Baermann trays with elutriation or sieving; centrifugal flotation; flotation-sieving; semiautomatic elutriation; and Cobb’s decanting and sieving. These methods are described in detail by

182 Meloidogyne fallax Secondary Pest of Corn Nematode False Columbia root-knot nematode Root-knot nematode

Barker (1985). The efficiency of nematode extraction is influenced by the amount of soil that is processed at one time. Extraction efficiencies are greatest when 100 g to 450 g of soil are processed. Extraction efficiencies for Meloidogyne spp. are frequently low and can vary between 13 and 45% (Davis and Venette, 2004).

Key Diagnostics CAPS-Approved Method: Confirmation of M. fallax is by morphological identification.

Literature-Based Methods: Meloidogyne fallax may occur in mixed or adjacent populations with M. chitwoodi, though it is reportedly rare for the two nematodes to share the same host. Unless very closely examined, Meloidogyne fallax, M. chitwoodi, and M. hapla may be easily confused (Karssen, 1996). Before M. fallax was first recorded in New Zealand, it had been misidentified as M. incognita (Marshall et al., 2001). M. fallax closely resembles M. chitwoodi; it can be distinguished by greater female and male stylet length, absence of small, irregular outlined male and female stylet knobs, male labial disc elevated, longer juvenile body-, tail-, and hyaline tail length, different hyaline tail shape, and hemizonoid position. It differs from M. hapla by the absence of fine, smooth striae, rounded and flattened dorsal arch and tail area punctations in the female perineal pattern, broader J2 tail and tail terminus with distinct hyaline part, shorter female and male stylet length, and the absence of small rounded stylet knobs (Davis and Venette, 2004).

Advances in molecular techniques have improved diagnoses among morphologically similar nematodes. Common molecular techniques to identify M. fallax include isozyme electrophoresis (Esbenshade and Triantaphyllou, 1987), protein patterns (Tastet et al. 1999), restriction fragment length polymorphism (PCR-RFLP) of ribosomal DNA (Peterson et al., 1997; Zijlstra, 1997a,b; Castagnone et al., 1999; Castagnone, 2000; Wishart et al., 2002), sequence characterized amplified region (SCAR) (Zijlstra, 2000, Adam et al., 2007), random amplified polymorphic DNA (RAPD) (Adam et al., 2007) and most recently real-time polymerase chain reaction (PCR) (Zijlstra and Hoof, 2006).

Petersen and Vrain (1996) and Wishart et al. (2002) developed PCR primers to identify M. chitwoodi, M. hapla, and M. fallax. Zijlstra (1997) uses a multiplex PCR to distinguish M. hapla, M. chitwoodi, M. fallax, and M. incognita, and Zijlstra (2000) uses SCAR-PCR to identify M. hapla, M. chitwoodi, and M. fallax. Adam et al. (2007) developed a molecular diagnostic key to identify juveniles of seven species, including M. incognita, M. javanica, M. arenaria, M. mayaguensis, M. hapla, M. chitwoodi, and M. fallax. Peterson et al. (1997) used a multiplex PCR to distinguish the juveniles and eggs of the same seven species. Fourie et al. (2001) distinguished the same species except M. mayaguensis. A real-time PCR for the simultaneous detection of M. chitwoodi and M. fallax has been developed (Zijlstra and van Hoof, 2006).

183 Meloidogyne fallax Secondary Pest of Corn Nematode False Columbia root-knot nematode Root-knot nematode

References Adam, M.A.M., Phillips, M.S., and Blok, V.C. 2007. Molecular diagnostic key for identification of single juveniles of seven common and economically important species of root-knot nematode (Meloidogyne spp.) Plant Pathology 56: 190-197.

Barker, K.R. 1985. Nematode extraction and bioassays, pp. 19-35. In K.R. Barker, C.C. Carter and J. N. Sasser [eds.], An advanced treatise on Meloidogyne, Vol II. Methodology. North Carolina State University Graphics, Raleigh.

CABI. 2007. Crop Protection Compendium Wallingford, UK: CAB International. http://www.cabi.org/compendia/cpc/.

Castagnone-Sereno, P., Leroy, F., Bongiovanni, M., Zijlstra, C., and Abad, P. 1999. Specific diagnosis of two root-knot nematodes, Meloidogyne chitwoodi and M. fallax, with satellite DNA probes. Phytopathology 89: 380-384.

Castagnone-Sereno, P. 2000. Use of satellite DNA for specific diagnosis of the quarantine root-knot nematodes Meloidogyne chitwoodi and M. fallax. EPPO Bulletin 30: 581-584.

Davis, E.E. and Venette, R.C. 2004. Mini Risk Assessment - False Columbia root-knot nematode: Meloidogyne fallax Karssen [Nematoda: Heteroderidae]. Assessed 2/11/10. http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/mfallaxpra.pdf.

Esbenshade, P.R. and Triantaphyllou, A.C. 1987. Enzymatic relationships and evolution in the genus Meloidogyne (Nematoda: Tylenchida). Journal of Nematology 19: 8-18.

Fourie, H., Zijlstra, C., and McDonald, A.H. 2001. Identification of root-knot nematode species occurring in South Africa using the SCAR-PCR technique. Nematology 3(7): 675-680.

Hooper, D.J. 1986. Preserving and staining nematodes in plant tissues. In: Southey JF, ed. Laboratory methods for work with plant and soil nematodes. London, UK: HMSO.

Karssen, G. 1996. Description of Meloidogyne fallax n. sp. (Nematoda : Heteroderidae), a root-knot nematode from The Netherlands. Fundamental and Applied Nematology 19: 593-599.

Marshall, J.W., Zijlstra, C., and Knight, K.W.L. 2001. First record of Meloidogyne fallax in New Zealand. Australasian Plant Pathology 30: 283-284.

Petersen, D.J. and Vrain, T.C. 1996. Rapid identification of Meloidogyne chitwoodi, M. hapla, and M. fallax using PCR primers to amplify their ribosomal intergenic spacer. Fundamental and Applied Nematology 19(6): 601-605.

Petersen, D.J., Zijlstra, C., Wishart, J., Blok, V., and Vrain, T.C. 1997. Specific probes efficiently distinguish root-knot nematode species using signature sequences in the ribosomal intergenic spacer. Fundamental and Applied Nematology 20: 619-626.

Society of Nematologists. n.d. Meloidogyne fallax. Exotic Nematode Plant Pests of Agricultural and Environmental Significance to the United States. http://nematode.unl.edu/pest39.htm.

Tastet, C., Bossis, M., Gauthier, J.P., Renault, L., and Mugniery, D. 1999. Meloidogyne chitwoodi and M. fallax protein variation assessed by two-dimensional electrophoregram computed analysis. Nematology 1: 301-314.

184 Meloidogyne fallax Secondary Pest of Corn Nematode False Columbia root-knot nematode Root-knot nematode

Vovlas, N. and Inserra, R.N. 1996. Distribution and parasitism of root-knot nematodes on Citrus, Nematology Circular No. 217. Florida Department of Agriculture & Consumer Services Division of Plant Industry.

Waeyenberge, L. and Moena, M. 2001. Meloidogyne chitwoodi and M. fallax in Belgium. Nematol. Medit. 91-97.

Wishart, J., Phillips, M.S., and Blok, M.C. 2002. Ribosomal intergenic spacer: a polymerase chain reaction diagnostic for Meloidogyne chitwoodi, M. fallax, and M. hapla. Phytopathology 92: 884-892.

Zijlstra, C. 1997a. A fast PCR assay to identify Meloidogyne hapla, M. chitwoodi, and M. fallax, and to sensitively differentiate them from each other and from M. incognita in mixtures. Fundamental and Applied Nematology 20: 505-511.

Zijlstra, C. 1997b. A reliable, precise method to differentiate species of root-knot nematodes in mixtures on the basis of ITS-RFLPs. Fundamental and Applied Nematology 20: 59-63.

Zijlstra, C. 2000. Identification of Meloidogyne chitwoodi, M. fallax and M. hapla based on SCAR-PCR: a powerful way of enabling reliable identification of populations or individuals that share common traits. European Journal of Plant Pathology 106: 283-290.

Zijlstra, C. and Hoof, R.A. 2006. A multiplex real-time polymerase chain reaction (Taqman) assay for the simultaneous detection of Meloidogyne chitwoodi and M. fallax. Phytopathology 96: 1255-1262.

185 Cochliobolus pallescens Primary Pest of Corn Plant Pathogens

Primary Pests of Corn (Full Pest Datasheet)

Cochliobolus pallescens

Scientific Name Cochliobolus pallescens (Tsuda & Ueyama) Sivanesan pallescens Boedijin (Anamorph)

Synonyms: Pseudocochliobolus pallescens, Curvularia leonensis

Common Name(s) Curvularia leaf spot, leafspot of cereals, black point of wheat, leaf spot of rubber, ear rot of barley, rot of garlic

Type of Pest Fungal

Taxonomic Position Class: Ascomycetes, Order: , Family:

Reason for Inclusion in Manual Requested by the CAPS community – Trade issue – A concern of international trading partners (e.g., New Zealand).

Pest Description The fungus Cochliobolus is the teleomorph (sexual stage) of Bipolaris and Curvularia spp., the causal agents of disease in a wide variety of economically important crop species and weeds. In general, the teleomorphic stage of this fungus is extremely rare in nature and thus the anamorphic stage (asexual) usually is the cause of infection in fields. Species of Curvularia are saprophytes or plant pathogens, occur mostly in tropical and subtropical areas, and are isolated from soil, air, organic matter, plant, and even animals. Some species are known cellulase producers.

Unless specific information is available for Cochliobolus pallescens, most information in this datasheet will cover the anamorph Curvularia pallescens. Curvularia pallescens is characterized primarily by 3-septate conidia with very pale cells (Tsuda and Ueyama, 1983). Cochliobolus pallescens is heterothallic, and the segregation ratio of the two mating types in conidial isolates is about 1:1.

186 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

Ascomata: Pseudothecia superficial, globose to subglobose, black, 250-750 µm wide, with protruding ostiolar breaks, developing on a columnar to flat stromata, firmly adhering to the substrate at the base; ostiolar beak 190-690 µm long with a hyaline apex. Asci almost cylindrical with a short stalk, usually 8-spored, 140-215 x 12-19 µm, among filamentous pseudoparaphyses (Tsuda and Ueyama, 1983; Sivanesan, 1990).

Ascospores: Flagelliform or filiform, colorless, tapering towards both ends, 6-13-septate, 125-215 x 2.5-6.3 µm, parallel or coiled in a certain portion in the (Tsuda and Ueyama, 1983; Sivanesan, 1990).

Hyphae: Septate, hyaline, branched, 2.71-4.11 µm (Rath and Dhal, 1972).

Conidiophores: Arising singly or in groups, terminally and laterally on hyphae, simple or rarely branched, straight or flexuous, sometimes geniculate near the apex, brown to dark brown, smooth, multiseptate, variable in length, up to 6 µm thick (Tsuda and Ueyama, 1983; Sivanesan, 1990).

Conidia: Acropleurogenous, straight or slightly-to-markedly curved to hook-shaped, pale to somewhat pigmented, almost concolorous, 3-distoseptate, smooth 17-32 x 7-12.5 µm (Sivanesan, 1990). Tsuda and Ueyama (1983) give the size of conidia as 15-32 x 5- 12.5 µm. Conidial colonies on natural substrata effuse, brown or grayish brown, hairy, in culture grayish brown, dark brown, cottony or velvety, usually with very pale layer of hyphae bearing conidiophores (Tsuda and Ueyama, 1983; Sivanesan, 1990).

Pereira Freire et al. (1998) redescribe Curvularia pallescens with information from scanning electron microscopy. The authors also describe culture variability using different growth media.

Biology and Ecology Curvularia pallescens grows within a temperature range of 15°C (59°F) to 35°C (95°F) with an optimum of 30°C (86°F) (Hasija, 1971). The fungus also grows at a pH range of 2.7 to 8.0 with an optimum of pH 5.4. Sporulation of Curvularia pallescens was optimum, however, at 15°C. On mycelial seeded plates, water agar was the best growth medium for sporulation. Temperatures of 15, 12 (53.6°F), and 18°C (64.4°F), in that order, supported the production of large numbers of . No sporulation was observed at 3 and 6°C (37.4 and 42.8°F) and very little occurred at 30°C (86°F) (Olufolaji, 1984). When spores were used to seed plates, however, the optimum temperature increased to 24°C (75.2°F) and malt extract and potato dextrose agars were superior to water agar. Viability of spores decreased with culture age and was totally lost after 10 months at 10°C (50°F) (Olufolaji, 1984). Olufolaji (1986) reports that optimum germination of spores occurs at 25-30°C (77-86°F) and 95-100% relative humidity.

Disease spread occurs via windborne conidia and in seed, as Curvularia pallescens is seed-borne.

187 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

Symptoms/Signs Corn: The primary symptoms of Curvularia pallescens on corn are straw-colored, circular to oval leaf spots (Fig. 1). A circular (sometimes oval) grayish spot is usually observed in the center of each lesion. Conidiophores are abundant in the grayish spot but may be absent from young lesions. Each lesion is delimited by a dark brown ring (border), which in turn is frequently surrounded by a yellow green, translucent halo. The veins of the leaf appear to restrict the lateral extent of the lesions. Lesions that extend laterally over two or even as many as four veins, however, are not uncommon. Lesions vary in size from less than 1 to 3 mm diameter in the circular type. Oval lesions vary from tiny specks to about 20 x 8 mm, the eventual size being determined by the corn variety. Sometimes many lesions coalesce and vary in size up to 60 x 12 mm (Mabadefe, 1969). Curvularia pallescens has been shown to cause white streaks on corn kernels when present with Fusarium moniliforme Figure 1. Curvularia leaf spot on corn. (Tripathi et al., 1977). Lesions can be Image courtesy of CIMMYT. found on the leaf sheath and husk leaves.

Canna: Quick blighting of petals and rotting of both open and unopened flowers (Rath and Dhal, 1972). Disease begins as minute discolored, usually yellow, irregular specks on petals. The spots gradually increase in size and were blighted. The central zone also turned darker in color. The affected buds did not open but rot. In advanced stages of disease, the pathogen invaded the gynoecium, the androecium, and the upper portion of the inflorescence axis. All the diseased flowers and buds dropped, leaving the decaying inflorescence axis.

Coriander: Root rot is observed with Curvularia pallescens. Roots become brownish to black and brittle; secondary root system is totally lacking. Basal portion of leaves become conspicuous giving a pale, ‘sick appearance’ to the plants. Severely infected plants show yellowing of tips of young leaves, which gradually spreads downward to the leaf blade. Ultimately the entire plant turns yellow and collapses due to rotting of the basal portion (Dwivedi et al., 1982).

188 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

Rubber: The disease is characterized by brownish-red spots with a yellow halo on leaves. Shrivelling of leaves was seen when leaves were infected at the ‘copper brown’ stage (Rajalakshmy, 1976).

Pearl millet: One of the fungi associated with a brown seed discoloration (Randhawa and Aulakh, 1984).

Wheat: Curvularia pallescens causes black point, defined as the discoloration of the embryo (germ) end and surrounding areas of the wheat kernel. The disease occurs any time from grain filling to near harvest (Hasija, 1963).

Pest Importance Curvularia pallescens is known to cause severe yield losses in maize in Nigeria and India, especially when combined with rust (Puccinia polysora) and blight (Helminthosporium maydis) (Oyeikan, 1975; Oyekan and Obajimi, 1977). In India under favorable environmental conditions, the disease is considered dangerous and chemical controls are employed (Grewal and Payak, 1976). Corn varieties evaluated for resistance in Nigeria were all proven to be susceptible to C. pallescens (Oyekan and Obajimi, 1977).

Curvularia pallescens is an opportunistic pathogen in individuals that are immune comprised. Curvularia pallescens has been reported causing a cutaneous infection in an immune compromised woman (Agrawal and Singh, 1995; Berg et al., 1995). Respiratory and cerebral disease has also been reported (Lampert et al., 1977).

Curvularia pallescens has been reported a mycoparasite of Botryosporium peristrophae in India (Chaurasia and Dayal, 1982). C. pallescens grew over and coiled the host hyphae and conidiophores. It also induced haustoria formation, penetration, lysis, and distortion of host hyphae. Chlamydospore formation inside the host hyphae was also observed.

Known Hosts Curvularia pallescens is polyphagous on many graminicolous and non-graminicolous hosts. Most references do not distinguish primary (major) from secondary (minor) hosts. The following is a list of reported hosts:

Hosts: Aeschynomene americana (shyleaf), Allium spp. (onion, leek), Alpinia zerumbet var. variegata (variegated shell ginger), Amaranthus gangeticus (Kahlalu amaranth), Arachis spp. (peanut), Axonopus compressus (broadleaf carpet grass), Bambusa tuldoides (), Bauhinia spp. (orchid tree), Borassus flabellifer (Palmyra palm), Bothriochloa glabra, Brachiaria mutica (para grass), Bougainvillaea spectabilis (great bougainvillaea), Brachiaria spp., Brassica spp. (mustards), Bursera simaruba (gumbo-limbo, West Indian birch), Cajanus cajan (pigeonpea), Calendula spp. (pot marigold), Calotropis spp. (milkweed), Canna indica (canna), Capsicum annuum (pepper), Carica papaya (papaya), Ceiba pentandra (kapok), Centrosema spp.

189 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

(butterfly peas), Chloris gayana (Rhodes grass), Cinnamomum spp. (cinnamon, camphor), Citrullus vulgaris (watermelon, vine), Citrus spp., Cocos nucifera (coconut), Colocasia esculenta (taro), Coriandrum sativum (coriander), Cupania macrophylla, Cymbopogon citratus (lemon grass), Cynodon dactylon (bermudagrass), Cyperus antillanus, Dactyloctenium aegyptium (Egyptian grass), Dahlia spp., Digitaria horizontalis (Jamaican crabgrass), Echinochloa spp. (barnyard grass, jungle rice), Elaeis guineensis (oil palm), Eleusine aegypti (star grass); Eleusine coracana (African/Indian finger millet), Eucalyptus spp., Fagopyrum spp. (buckwheat), Gaillardia spp. (blanket flowers), Gloriosa superba (glory lily), Glycine max (soybean), Gmelina arborea (gumhar), Hevea brasilensis (rubber), Hordeum spp. (barley), Hymenocallis littoralis (beach spider lily), arundiacea, Imperata cylindrica (cogongrass), Lens culinaris (lentil), Liquidambar macrophylla, Lycopersicon esculentum (tomato), Mangifera indica (mango), Mnesithea laevis, Musa spp. (banana, plantain), Oryza sativa (rice), Panicum spp. (switch grass), Paspalum spp. (knot grass), Pennisetum americanum (pearl millet), Pennisetum spp. (pearl millet), Peperomia spp. (radiator plant), Phaseolus vulgaris (bean), Pinus patula (Mexican weeping pine), Pinus spp., Poa spp. (bluegrass), Psophocarpus tetragonolobus (winged bean), Punica granatum (pomegranate), Quercus spp., Rhynchelytrum repens (rose natal grass), Ricinus communis (castorbean), Rottboellia exaltata (itchgrass), Saccharum spp. (sugarcane), Schizachyrium hirtiflorum (crimson bluestem), Senna spp., Setaria glauca (yellow foxtail), Setaria spp., Solanum melongena (eggplant), Solanum tuberosum (potato), Sorghum bicolor (sorghum), Sorghum halapense (Johnson grass), Sorghum spp., Sporobolus poiretti (smut grass), Stenotaphrum secundatum (St. Augustine grass), Stephania abyssinica, Stylosanthes guianensis (stylo), Triticum spp. (wheat), Vigna unguiculata (cowpea), Urena sinuata (bur mallow), Vicia faba (fava bean), and Zea mays (corn) (Subramanian, 1953; Hasija, 1963; Rath and Dhal, 1972; Mathur et al., 1973; Rajalakshmy, 1976; Lal and Tripathi, 1977; Tripathi et al., 1977; Dwivedi et al., 1982; Randhawa, 1984; Sivanesan, 1990; Rao et al., 1992; Amadi et al., 1996; Thaung, 2008; Farr and Rossman, 2010).

Known Vectors (or associated insects) Cochliobolus pallescens is not known to be a vector, is not known to be vectored by another organism, and does not have any associated organisms.

Known Distribution Africa: Egypt, Ethiopia, Ghana, Guinea, Kenya, Malawi, Nigeria, Sierra Leone, Somalia, South Africa, Swaziland, Sudan, Tanzania, Togo, Zambia, and Zimbabwe. Asia: Bangladesh, Brunei, Burma (Myanmar), China, India, Indonesia, Iran, Japan, Malaysia, Nepal, Pakistan, Philippines, Saudi Arabia, Singapore, Sri Lanka, and Thailand. Caribbean: Barbados, Cuba, Jamaica, West Indies, and Windward Islands Europe: Denmark. North America: Canada, Mexico, and United States of America. Oceania: American Samoa, Australia, Fiji, Guam, Marshall Islands, Papua New Guinea, and Solomon Islands. South America: Brazil, Colombia, Guyana, Peru, Uruguay, and Venezuela.

190 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

Potential Distribution within the United States Gulya et al. (1979) report recovery of Curvularia pallescens from corn with ear rot in Iowa, but it was considered as a miscellaneous species. The ‘miscellaneous species’ group were considered saprophytes, or at best, weak or secondary pathogens. Bean (1964) reports C. pallescens associated with leaf spot of blue grass in the Washington, D.C. area. The validity of these records, however, is not known at this time. Due to its extremely broad host range of the pathogen, pathogen spread would not be limited within the United States. A risk map is not currently available for this pathogen.

Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Visual survey: Conduct visual inspection for symptoms associated with Curvularia pallescens. Discolored seed may also be collected and assessed for the presence of the fungus.

To isolate Curvularia pallescens from infected plant material, lesions (usually with some of the uninfected tissue around them) are cut into pieces about 4-5 mm square. The squares are then surface sterilized and rinsed in sterile water. The squares were plated on agar (potato dextrose agar (PDA) or potato carrot agar (PCA)) with three pieces per Petri dish. Plates were incubated at room temperature (22-25°C, 72-77°F) for 5 days, by which time the colonies growing out of the leaf pieces will normally have begun to sporulate (Mabadefe, 1969).

Seeds have been assayed for internal fungi using 1) the agar plate method and 2) the blotter method (Amadi and Oso, 1996). In the agar plate method, seeds were surface sterilized with 0.5% NaOCl for 5 minutes and rinsed twice with sterile distilled water. Seeds were blotted dry between sterile Whatman No. 1 filter papers before plating on 5 mm diameter agar (PDA) disks. Inoculated plates were incubated at 27+1°C (81+1°F) under alternating light and dark regimes. In the blotter method, the same procedure was followed as in the agar plate method. Wet, sterile, Whatman No. 1 filter papers (9 cm) were used in this method in place of the PDA disks.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Identification is based on fungal morphology. In general, conidia are dark, 3-5 celled, end cells curved, more or less fusiform, with one of the central cells enlarged.

Easily Confused Pests There are a range of Curvularia spp. that cause leafspots on corn that could be confused with Curvularia pallescens. Most of these species are considered of minor

191 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

importance or do not occur in the United States (White, 1999). Southern corn leaf blight is caused by Cochliobolus heterostrophus but has a Bipolaris anamorph.

References Agrawal, A., and Singh, S.M. 1995. Two cases of cutaneous phaeohyphomycosis caused by Curvularia pallescens. Mycoses 38(7-8): 301-303.

Amadi, J.E., and Oso, B.A. 1996. Mycoflora of cowpea seeds (Vigna unguiculata) L. and their effects on seed nutrient content and germination. Nigerian Journal of Science 30: 63-69.

Bean, G.A. 1964. Prevalence of Curvularia pallescens and Helminthosporium spp. pathogenic on blue grass in Washington D.C., area. Phytopathology 54: 888

Berg, D., Garcia, J.A., Schell, W.A., Perfect, J.R., and Murray, J.C. 1995. Cutaneous infection caused by Curvularia pallescens: A case report and review of the spectrum of disease. Journal of the American Academy of Dermatology 32: 375-378.

Chaurasia, S.N.P., and Dayal, R. 1982. Curvularia pallescens – a destructive mycoparasite of Botryosporium peristrophae. Indian Phytopath. 35(4): 671-674.

Dwivedi, D.K., Shukla, D.N., and Bhargava, S.N. 1982. Two new root-rot disease of spices. Current Science 51(5): 243-244.

Farr, D.F., and Rossman, A.Y. Fungal Databases, Systematic Mycology and Microbiology Laboratory, ARS, USDA. http://nt.ars-grin.gov/fungaldatabases/.

Grewal, R.K., and Payak, M.M. 1976. Evaluation of systemic and non-systemic fungicides in vitro against Curvularia pallescens. Pesticides 10(4): 36-37.

Gulya, Jr., T.J., Martinson, C.A., and Tiffany, L.H. 1979. Ear-rotting fungi associated with opaque-2 maize. Plant Disease Reporter 63(5): 370-373.

Hasija, S.K. 1963. A new record of Curvularia pallescens Boed. on wheat grains. Indian Phytopathology 16: 375-377.

Hasija, S.K. 1971. Physiological studies of Curvularia pallescens. Nova Hedwigia 19(3/4): 551-558.

Lal, S., and Tripathi, H.S. 1977. Host range of Curvularia pallescens, the incitant of leaf spot of maize. Indian J. Mycol. Plant Path. 7(1): 92-93.

Lampert, R.P., Hutto, J.H., Donnelly, W.H., and Schulman, S.T. 1977. J. Pediatrics 91(4): 603-605

Mabadeje, S.A. 1969. Curvularia leaf spot of maize. Trans. Br. Mycol. Soc. 52(2): 267-271.

Mathur, S.K., Nath, R., and Mathur, S.B. 1973. Seed-borne fungi of pearl millet (Pennisetum typhoides) and their significance. Seed Sci. & Technol. 1: 811-820.

Olufolaji, D.B. 1984. Sporulation and growth of Curvularia pallescens as affected by media, temperature, and nitrogen sources. Phytopathology 74: 260-236.

Olufolaji, D.B. 1986. Optimum temperature and relative humidity for germination and germ tube growth of Curvularia pallescens on glass slides and maize leafs. Cryptogamie, Mycol. 7(2): 149-156.

Oyekan, P.O. 1975. The effect of Curvularia leaf spot on yield of maize. Nig. Soc. Plant Protection. Occassional Publication 1: 27 (Abstr.).

192 Cochliobolus pallescens Primary Pest of Corn Fungus Leaf spot

Oyekan, P.O., and Obajimi, A.O. 1977. Reaction of maize varieties to Curvularia leaf spot in south western Nigeria. Niger. J. Pl. Prot. 3: 116-188.

Pereira Freire, S.V., Mesquita Paiva, L., de Luna-Alves Lima, E.A., and Costa, Maia, L. 1998. Morphological, cytological, and cultural aspects of Curvularia pallescens. Reivista de Microbioloiga 29(3). Doi:10.1590/Sooo1-37419980003000010. http://www.scielo.br/scielo.php?script=sci_arttext&pid=S0001- 37141998000300010.

Rajalakshmy, V.K. 1976. Leaf spot disease of rubber caused by Curvularia pallescens Boedijin. Current Science 24: 530

Randhawa, H.S., and Aulakh, K.S. 1984. Mycoflora associated with discoloured and shriveled seeds of pearl millet. Indian Phytopath. 37(1): 119-122.

Rao, G.P., Singh, S.P., and Singh, M. 1992. Two new alternative hosts of Curvularia pallescens, the leaf spot causing fungus of sugarcane. Tropical Pest Management 38(2): 218.

Rath, G.C., and Dhal, N.K. 1972. Petal blight of Canna indica. Indian J. Mycol. Plant. Pathol. 19: 90-91.

Sivanesan, A. 1990. CMI Descriptions of Fungi and Bacteria No. 1003- Cochliobolus pallescens. Mycopathologia 111: 115-116.

Subramanian, C.V. 1953. Fungi Imperfecti from Madras – V. Curvularia. Proc. Indian Acad. Sci. 38B: 27- 39.

Thaung, M.M. 2008. A list of Hyphomycetes (and Agonomycetes) in Burma. Australasian Mycologist 27(3): 149-172.

Tripathi, H.S., Lal, S., and Agrawal, V.K. 1977. Note on the influence of fungicidal sprays on percent seed-borne incidence of Fusarium monilforme Sheld. and Curvularia pallescens Boed. In maize. Pantnagar J. Res. 2(1): 104-105.

Tsuda, M., and Ueyama, A. 1983. Pseudocochliobolus pallescens and variability of morphology. Mem. Coll. Agric., Kyotat Univ. 122: 85-91.

White, D.G. (ed.) Compendium of Corn Diseases, 3rd ed. APS Press, St. Paul, MN. 78 pp.

193 Harpophora maydis Primary Pest of Corn Fungus Late wilt

Harpophora maydis

Scientific Name Harpophora maydis (Samra, Sabet and Hingorani) Gams

Synonyms: Cephalosporium maydis and Acremonium maydis

Common Name(s) Late wilt of corn, ‘Shallal’ disease of maize, and black bundle disease

Type of Pest Fungal Pathogen

Taxonomic Position Class: Ascomycetes Order: Incertae sedis Family: Magnaporthaceae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 & 2010

Pest Description Taxonomic note: Late wilt is an important disease in Egypt and parts of India. It was first recorded in Egypt in 1960 and described as a new species, Cephalosporium maydis (Samra et al., 1962, 1963). The description was flawed in that it did not refer to a type specimen. Domsch and Gams (1972) suggested that the conidial state of C. maydis was a Phialophora (the anamorph of Gauemannomyces) and that spore production in C. maydis was typical of that genus (Ward and Bateman, 1999). Most members of the genus Cephalosporium were transferred to the genus Acremonium, a genus of hyaline hyphomycete with aculeate (spine-like) phialides unrelated to either Phialophora or Harpophora. Gams (2000) introduced Harpophora as a new genus (contains anamorphs of Gauemannomyces and Magnaporthe) that is distinct from Phialophora. Harpophora spp. are characterized by fast-growing, thin colonies with sickle-shaped conidia. Older hyphae are heavily pigmented, younger hyphae are nearly hyaline, and phialides are intermediate in pigmentation relative to the older and younger hyphae. When he introduced Harpophora, Gams (2000) also introduced the new combination Harpophora maydis (Samra, Sabet, and Hingorani) Gams as a replacement for Cephalosporium maydis.

Hyphae of H. maydis are hyaline, septate, branched, and decumbent. Conidiophores may develop terminally or laterally and contain five to eight conidia clustered in heads. Conidiophores are 30-250 µm long, but occasionally are 400 µm long. They are hyaline, mostly septate, branched, and straight. Conidia are produced successively and exogenously at apices of the conidiophore so that several spores collect in heads. Conidia are hyaline, straight, single-celled, oblong, and measure 3.6-14 x 3-3.6 µm

194 Harpophora maydis Primary Pest of Corn Fungus Late wilt

(avg. 7.2 x 3.5 µm). Conidia germinate rapidly by two polar germ tubes, but one and sometimes three germ tubes may be formed. Anastomosis of germ tubes in unusually frequent in this fungus. After about 3 weeks of growth on potato dextrose agar (PDA), conidia are difficult to find in culture. Small botryoid sclerotia-like bodies, consisting of a few thick-walled, dark colored cells, appear in old cultures (Samra et al., 1963).

The pathogen grew well and sporulated on oatmeal, prune, PDA, malt extract, and yeast extract-glucose agar. The minimum temperature for growth was 12°C (54°F), the optimum 25-30°C (77-86°F), and the maximum 30°C. No growth was observed at 8 and 38°C, (46 and 100°F ) even after 4 weeks, Spores appeared within 2 days after seeding on the media at 22-24°C (72-75°F) (Samra et al., 1963). The pathogen may be successfully isolated onto potato dextrose agar amended with 0.2% yeast extract and incubated at 29-30ºC (84-86ºF) (Michail et al., 1999). Young colonies are white, low growing and dense like felt, but may turn gray or black over time. Colony margins have a characteristic “rhizoid” appearance. Small sclerotia may develop in culture (Payak et al., 1970).

Biology and Ecology The fungus is soilborne and causes a vascular wilt disease that most commonly infects seedling plants and has no specific moisture requirements. Singh and Siradhana (1987a) found that three irrigations at an interval of eight hours after inoculation supported maximum disease. Infection in corn occurs through the roots or mesocotyl (Sabet et al., 1970b). Infections have been shown to occur through stalks in India.

As plants mature, fewer are infected, and they become immune about 50 days after planting. Initially, the fungus grows superficially on the roots, producing hyphae with short, thick-walled, swollen cells (Sabet et al., 1970b). After penetration, H. maydis colonizes xylem tissue and is rapidly translocated to the upper parts of the plant. When infections are severe, the fungus colonizes the kernels, resulting in seedborne dissemination and Figure 1. Rapid wilting of also causes seed rot and damping off (El-Shafey infected plant alongside and Claflin, 1999; Michail et al., 1999). No perfect resistant hybrids. Photo (sexual) state has been identified (Saleh and courtesy of H. Warren. Leslie, 2004).

H. maydis can remain viable in the soil for several years in the absence of a host. H. maydis has been shown to persist on corn stubble for 12-15 months (Sabet et al., 1970b; Singh and Siradhana, 1987b). Inoculum survival in soil is generally poor and restricted to the top 20 cm of soil. Although it is a weakly competitive saprophyte (Sabet

195 Harpophora maydis Primary Pest of Corn Fungus Late wilt

et al., 1970a), the production of sclerotia in infested host debris ensures its long-term survival. Lupine, an alternative host, can play a role in the parasitic survival of the pathogen. The pathogen is most common in hot and humid environments and in heavy- textured soils. Saturated soils lessen the incidence of H. maydis.

Dawood et al. (1979) noted that H. maydis produced no sclerotia on infected plants during its parasitic phase. It also did not form sclerotia on infected dead plants kept at room temperature for about six months. Sclerotia, however, were produced abundantly in pure cultures grown on enriched maize stalk pieces for 30-45 days at 30°C (86°F), followed by drying under electric air fan. Maximum number of sclerotia was produced at 30°C, followed by 35°C (95°F), and then 25°C (77°F). Considerably less sclerotia were produced at 20°C (68°F). The number of sclerotia decreased as the atmospheric humidity increased from 70 to 100%. Sclerotia also formed on stalk pieces of naturally infected, field grown plants when such pieces were buried in the soil. However, the viability of these sclerotia was rather low.

Figure 2. Leaf streaking of Harpophora infected plant (R) compared with healthy leaf Progressive development of (L). Photo from Sabet et al., Figure 3. yellow to reddish-brown streaks on H. 1966c. maydis infected lower stalks. Photo courtesy of H. Warren. The sclerotia freshly from stalk piece cultures or kept in a refrigerator germinated readily on antibiotic-containing Richards’ solution agar. Germination was visible with 48 hours of incubation at 30°C (86°F). Sabet (1984) developed a technique to produce uniform, abundant sclerotia on Farlene- glucose agar, where sclerotia were harvested after five weeks of incubation at 30°C.

196 Harpophora maydis Primary Pest of Corn Fungus Late wilt

Symptoms/Signs Corn: Root tips of infected corn plants are stained red during early stages of infection, but aboveground parts generally remain symptomless until tasseling when a rapid wilting (Fig. 1) of lower leaves progresses upward. The time of onset may extend from just prior to tasseling until shortly before maturity. The wilting progresses from the lower to the upper portions of the plant. Leaves become dull green, eventually lose color (Fig. 2), and become dry as through suffering form lack of water. Leaves assume a scorched appearance. Vascular bundles in the stalk become reddish brown and within a short period, lower internodes (area between nodes on the stalk) assume this color (Fig. 3) (Samra et al., 1963; El-Shafey and Claflin, 1999).

In advanced stages, lower portions of the stalk become dry, shrunken, and hollow (Fig. 4). Stalk symptoms may be modified depending on the extent of invasion of saprophytic organisms. Secondary infection by other organisms frequently progresses into stalk rot (soft and wet). According to Jain et al. (1974) a ‘sweetish smell’ often accompanies the wet rot. After the first wilt symptoms appear, progress of the disease is relatively rapid. Because of the delay in appearance of initial symptoms until about flowering, this Figure 4. Dry disease has been designated as ‘late wilt’ (Samra et al., hollow H. maydis 1963). Kernels that form may be poorly developed. Growers infected stalk. often only recognize stalk rot diseases in India during the Photo courtesy of final stages when the stalks begin to lodge (fall over), CIMMYT. especially when intensified by delayed harvest or wind damage (Jain et al., 1974).

Cotton: Reddish lesions and shallow cracks have been observed on cotton roots (Bahteem 185 cultivar) grown in inoculated soil. These lesions disappear as the cotton plants mature, and H. maydis has not been recovered from them (Sabet et al., 1966a). No aboveground effects are observed throughout the growth of the plants up to maturity. It is not known how important infections of H. maydis on cotton are to subsequent infections on corn.

Lupine: Wilt and root rot (Sabet et al., 1966b). H. maydis causes a significant damping- off and stunting of the widely cultivated Lupinus terminis in Egypt (Sahab et al., 1985).

Pest Importance Stalk and root rots are, in general, the most serious and widespread group of diseases affecting corn growing regions of the world (Sabet et al., 1966b). Harpophora maydis causes a severe stalkrot. In Egypt, 90% of the plants of susceptible cultivars may be infected. Yield losses of up to 40% have been reported (Jain et al., 1974; El-Shafey and

197 Harpophora maydis Primary Pest of Corn Fungus Late wilt

Claflin, 1999). The severity of the late wilt disease has diminished in most corn plantings due to the introduction of new resistant hybrids, which have replaced the local susceptible cultivars, through an active breeding program against late wilt disease by the Egyptian Ministry of Agriculture (Mostafa et al., 1996) and Ramana et al. (1997) and Satyanarayana (1995) in India. The extent of resistance or tolerance in corn lines adapted for the United States to late wilt, however, is not known since this disease is not commonly screened for in U.S. breeding programs (Bergstrom et al., 2008).

Although evidence exists that the H. maydis population in Egypt is clonal, at least four phylogenetic lineages are present (Zeller et al., 2000; Saleh et al., 2003). These lineages differ in their ability to colonize maize plants and their relative aggressiveness in single culture inoculations or both (Zeller et al., 2002). They also differed in mixed culture inoculations (El-Assiuty et al., 1999). Adequate understanding of where each lineage is located within a country and using all lineages to challenge host material during the development of resistant germplasm is needed to best deploy host resistance. For example, corn germplasm that is susceptible to lineage IV might be well suited for part of the country where this lineage is not present but not in parts of the country where it is present.

Known Hosts Zea mays (corn), Gossypium hirsutum (cotton), Lupinus terminus (lupine)

Known Vectors (or associated insects) H. maydis is not a known vector, but, it attacks the roots of corn and can allow other pathogens (fungi and viruses) access to the plant. Sabet et al. (1966a) showed that infection of cotton roots with H. maydis decreased the severity of cotton wilt, caused by Fusarium oxysporum.

Known Distribution H. maydis has been reported from Egypt, India, Hungary, Israel, Italy, Portugal, Spain (Samra et al., 1962, 1963; Payak et al., 1970; Pecsi and Nemeth, 1998; Bergstrom et al., 2008). There also are unconfirmed reports of the disease in Romania and Kenya which imply that some strain(s) of the pathogen are capable of surviving climates similar to U.S. corn production regions (Bergstrom et al., 2008).

Potential Distribution within the United States The pathogen is not currently known to exist in the United States, but poses a serious threat to corn production in this country. The organism can be easily moved in shipments that contain either infested soil or seed. Its ability to withstand high temperatures would allow it to survive in the southern United States. Growing conditions during May-June are most conducive. Based on climate models alone, the southern half of the country would be favorable for disease development. When the climatic model is combined with the geographic distribution of corn production in a recent risk analysis by USDA-APHIS-PPQ-CPHST, most of the continental United States is at moderate risk of H. maydis establishment. Areas of Arkansas, Illinois,

198 Harpophora maydis Primary Pest of Corn Fungus Late wilt

Indiana, Mississippi, Missouri, Tennessee, and Texas, have the highest risk for establishment of H. maydis.

Survey CAPS-Approved Method: Visual survey is the method to survey for H. maydis by collecting symptomatic plant material.

Literature-Based Methods: Visual survey: surveying for disease is difficult because symptoms cannot be identified until tassel emergence when host height makes viewing large numbers of plants difficult. At tasseling, fields should be monitored visually for characteristic symptoms. Symptom recognition is based on the dull green, desiccated (scorched) leaves, streaked and “collapsed” stalk, and discolored pith tissues. Identification may be complicated by the similarity of symptoms caused by other common problems, such as nitrogen deficiency, but which may occur over larger areas and following excessively wet weather when nitrogen sources may be leached beyond the reach of growing roots.

Key Diagnostics CAPS-Approved Method: Confirmation of H. maydis is by morphological identification. Pathogen may be identified morphologically by examination of the shape and size of conidia and conidiophores, color and type of colony, and temperature requirements.

Literature-Based Methods: Symptoms are not definitive and morphological and microscopic characteristics are still used to identify H. maydis (El-Shafey and Claflin, 1999). Isolates can differ in virulence and competitiveness (Zeller et al., 2002); thus, isolation, culture, direct microscopic evaluation, pathogenicity tests, or PCR are required for positive identification. Ward and Bateman (1999) use a pair of PCR primers that amplify a segment of the ribosomal gene locus from many members of the Gaeumannomyces- and Phialophora fungal pathogens from maize and other host plants. The PCR product from Harpophora maydis (=Cephalosporium maydis) can be distinguished from that of other members of the group on the basis of its unique size (490 bp) relative to that of other species. Species-specific PCR primers capable of distinguishing H. maydis from other species in the Gaeumannomyces-Harpophora complex have been developed and can be used for identification, but need to be validated for regulatory purposes (Saleh and Leslie, 2004; Zeller et el., 2000).

Successful isolation can usually be obtained by sterilizing the internode of symptomatic plants in 5% sodium hypochlorite (bleach), splitting them with a sterile knife, and placing a small piece of discolored vascular bundle on PDYA media (PDA + 0.2% yeast extract) (Samra and Sabet, 1966; Zeller et al., 2002). Single spore isolates can be obtained by dilution plating. Recovery of H. maydis, even from heavily infested material, is difficult due to its slow growth and to the relative abundance of other more rapidly growing fungi, most commonly Fusarium spp. (Saleh et al., 2003).

Infected seeds do not show discernible external symptoms and cannot be identified visually. H. maydis can be cultured from infected seed by soaking seeds in 1% sodium

199 Harpophora maydis Primary Pest of Corn Fungus Late wilt

hypochlorite for 3 minutes, plating on PDYA, incubating at 20°C (68°F) under 12 hour cycles of alternating near-ultraviolet light and darkness, and examining after 24 hours. Identification of cultures is accomplished by spore morphology and pathogenicity tests. The pathogen can be identified in tissue using PCR techniques that are not influenced by secondary fungal invaders (Saleh and Leslie, 2004).

Easily Confused Pests Late wilt does not occur in the United States and may not be readily recognized or distinguished initially from abiotic stresses without some training.

References Bergstrom,G., Leslie, J., Huber, D., Lipps, P., Warren, H., Esker, P., Grau, C., Botratynski, T., Bulluck, R., Floyd, J., Bennett, R., Bonde, M., Dunkle, L, Smith, K., Zeller, K., Cardwell, K., Daberkow, S., Bell, D., and Chandgoyal, T. 2008. Recovery plan for late wilt of corn caused by Harpophora maydis syn. Cephalosporium maydis. http://www.ars.usda.gov/SP2UserFiles/Place/00000000/opmp/Corn%20Late%20wilt%2081112.pdf.

Dawood, N.A., Sabet, K.K., and Sabet, K.A. 1979. Formation and survival of the sclerotia of Cephalosporium maydis. Agricultural Research Review 57(2): 185-199.

Domsch, K.H. and Gams, W. 1972. Fungi in agricultural soils. Halsted Press, New York.

El-Assuity, A.M., Ismael, A.M., Zeller, K.A., and Leslie, J.F. 1999. Relative colonization of greenhouse- grown maize by four lineages of Cephalosporium maydis from Egypt. Phytopathology 89(6): S23 (Abstr.)

El-Shafey, H.A. and Claflin, L.E. 1999. Late Wilt. Pp. 43-44. In: D. G. White (ed.) Compendium of Corn Diseases, 3rd ed. APS Press, St. Paul, MN. 78 pp.

Gams, W. 2000. Phialophora and some similar morphologically little-differentiated anamorphs of divergent ascomycetes. Stud Mycol. 45: 187-199.

Jain, K.L., Dange, S.R.S., and Kothari, K.L. 1974. Stalk rots of maize in Rajasthan. Farm and Factory 8(10): 20-23.

Michail, S.H., Abou-Elseoud, M.S., and Eldin, M.S.N. 1999. Seed health testing of corn for Cephalosporium maydis. Acta Phytopathologica Entomologica Hunarica 34(1/2): 35-41.

Mostafa, M.A.N., El-Aziz, A.A.A., Magoub, G.M.A., and El-Sherbiney, H.Y.S. 1996. Diallel analyses of grain yield and natural resistance to late wilt disease in newly developed inbred lines of maize. Bulletin of Faculty of Agriculture, University of Cairo 47: 393-403.

Payak, M.M., Lal, S., Lilaramani, J., and Renfro, B.L. 1970. Cephalosporium maydis – a new threat to maize in India. Indian Phytopathology 23(3): 562-569.

Pesci, S. and Nemeth, L. 1998. Appearance of Cephalosporium maydis Samra, Sabet, and Hingorani in Hungary. Facul. Lanbouw. En Toegepaste Biolog. Wetenschappen, Univ. Gent 63: 873-877.

Ramana, V.V., Reddy, V.K., Aeddy, S.M., and Reddy, K.J. 1997. Production of cellulases, hemicellulases, pectinases, proteinases, and lipases by Cephalosporium maydis isolated from Zea mays stalks, In: Reddy, S.M., Srivastava, H.P., Purchit, D.K., Ram Reddy, S. (eds.): Microbial Biotechnology. Jodphur, India. Scientific Publishers, pg. 187-192.

200 Harpophora maydis Primary Pest of Corn Fungus Late wilt

Sabet, K.K. 1984. A technique for sclerotial production by Cephalosporium maydis. Plant Prot. Bull. 32(4): 141-142.

Sabet, K.A., Samra, A.S., and Mansour, I.M. 1966a. Interaction between Fusarium oxysporum, F. vasifectum, and Cephalosporium maydis on cotton and maize. Ann. Appl. Biol. 58: 93-101.

Sabet, K.A., Samra, A.S., and Mansour, I.M. 1966b. Late wilt disease of maize and a study of the causal organism. Investigations of stalk rot disease of maize in U.A.R. Tech. Bull. Min. Agric. 1-1. 45 pp.

Sabet, K.A., Samra, A.S., and Abdel-Rahim, M.F. 1966c. Seed transmission of stalk-rot fungi and effect of seed transmission. pp 94-116. In: Samra, A.S. and Sabet, K.A. (eds). Investigations on Stalk-rot Disease of Maize in U.A.R. Ministry of Agriculture, Government Printing Offices, Cairo, Egypt.

Sabet, K.A., Samra, A.S., and Mansour, I.M. 1970a. Saprophytic behavior of Cephalosporium maydis and C. acremonium. Ann. Appl. Biol. 66: 265-271,

Sabet, K.A., Zaher, A.M., Samra, A.S., and Mansour, I.M. 1970b. Pathogenic behaviour of Cephalosporium maydis and C. acremonium. Ann. Appl. Biol. 66: 257-263.

Sahab, A.F., Osman, A.R., Soleman, N.K., and Mikhail, M.S. 1985. Studies on root-rot of lupin in Egypt and its control. Egypt. J. Phytopathol. 17:23-35.

Saleh, A.A. and Leslie, J.F. 2004. Cephalosporium maydis is a distinct species in the Gaeumannomyces-Harpophora species complex. Mycologia 96: 1294-1305.

Saleh, A.A., Zeller, K.A., Ismael, A.-S.M., Fahmy, Z.M., El-Assiuty, E.M., and Leslie, J.F. 2003. Amplified fragment length polymorphism in Cephalosporium maydis from Egypt. Phytopathology 93: 853- 859.

Samra, A.S. and Sabet, K.S. 1966. Investigations on stalk-rot disease of maize in U.A.R.: an introductory note. Min. Agric. Plant. Prot. Dep. Bull., U.A.R. pp. 1-7.

Samra, A.S., Sabet, K.A., and Hingorani, M.K. 1962. A new wilt of maize in Egypt. Plant Disease Reporter 46: 481-483.

Samra, A.S., Sabet, K.A., and Hingorani, M.K. 1963. Late wilt disease of maize caused by Cephalosporium maydis. Phytopathology 53:402-406.

Satyanarayana, E. 1995. Genetic studies of late wilt and turcicum leaf blight resistance in maize. Madras Agricultural Journal 82: 608-609.

Singh, S.D. and Siradhana, B.S. 1987a. Influence of some environmental conditions on the development of late wilt of maize induced by Cephalosporium maydis. Indian Journal of Mycology and Plant Pathology 17:1-5.

Singh, S.D. and Siradhana, B.S. 1987b. Survival of Cephalosporium maydis, incitant of late wilt of maize. Indian J. Mycol. Pl. Pathol. 17: 83-85.

Ward, E. and Bateman, G.L. 1999. Comparison of Gaeumannomcyes- and Phialophora-like fungal pathogens from maize and other plants using DNA methods. New Phytologist 141: 323-331.

Zeller, K.A., Jurgenson, J.E., El-Assiuty, E.M., and Leslie, J.F. 2000. Isozyme and Amplified Fragment Length Polymorphisms from Cephalosporium maydis in Egypt. Phytoparasitica 28(2): 121-130.

201 Harpophora maydis Primary Pest of Corn Fungus Late wilt

Zeller, K.A., Ismael, A.-S.M., Fahmy, A.M., and Bekheet, F.M. 2002. Relative competitiveness and virulence of four clonal lineages of Cephalosporium maydis from Egypt toward greenhouse-grown maize. Plant Disease 86: 373-378.

202 Peronosclerospora maydis Primary Pest of Corn Fungal-like Java downy mildew

Peronosclerospora maydis

Scientific Name Peronosclerospora maydis (Racib.) C.G. Shaw

Synonyms: Peronospora maydis and Sclerospora maydis

Common Name Java downy mildew, downy mildew of corn, and corn downy mildew

Type of Pest Fungal-like pathogen

Taxonomic Position Phylum: Oomycota, Class: Oomycetes, Order: Sclerosporales, Family: Sclerosporaceae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009

Pest Description Java downy mildew, caused by Peronosclerospora maydis, was discovered by Raciborski (1897) in Java, Indonesia in 1897 and has the distinction of being the first downy mildew disease reported on corn (Bonde, 1982). Initially it was misidentified as Sclerospora maydis. The disease was reported in India by Butler (1913) and has continued to spread.

Peronosclerospora maydis is an obligate parasite that will not grow on artificial media. The pathogen produces two kinds of hyphae: straight and sparsely branched, and lobed and irregularly branched. The mycelium has many haustoria with different forms (Semangoen, 1970). Clustered conidiophores arise from stomata and are dichotomously branched two to four times. The branches are robust and 150-550 µm long with basal cells 60-180 µm long. Conidia (17-23 x 27-39 µm) are hyaline and spherical to subspherical (Smith and Renfro, 1999). Semangoen (1970), however, indicated that the conidia are smaller (12-29 x 10-23 µm). Oospores have not been reported (Smith and Renfro, 1999).

Biology and Ecology Infected corn plants grown during the dry season are the primary source of inoculum in Indonesia. The fungus may also survive as mycelium in kernels, but this is thought to be a minor source of inoculum. Infection by conidia occurs through stomata of young plants, and lesions elongate toward the meristem, inducing systemic infection. P. maydis caused high levels of systemic infection from 8 to 36°C (46 to 97°F) (Bonde et al., 1992). If infection arises from seed, the cotyledonary leaf is always infected. Seed

203 Peronosclerospora maydis Primary Pest of Corn Fungal-like Java downy mildew

transmission occurs when freshly harvested seeds from diseased plants are used. No seed transmission has been detected from seeds dried prior to planting (Smith and Renfro, 1999).

Large numbers of conidia and conidiophores are produced on the upper leaf surface on wet leaves (dew for 5-6 hours) in the dark from 18-23°C (64-73°F) (the optimum temperatures for sporulation) (Inaba et al., 1980; Bonde et al., 1992). P. maydis has a very broad optimum temperature range for conidial germination (at least 10-30°C, 50- 86°F) and germ tube growth (18-30°C, 64-86°F) (Bonde et al., 1992). Conidia germinate under these conditions to produce germ tubes with free water being required (Semangoen, 1970; Smith and Renfro, 1999). Conidia formation is initiated from 12:00 AM to 1:00 AM, and the peak of conidial release is at 3:00 AM to 4:00 AM. Most conidia remain within a 16 meter radius of their source plant (Tantera, 1975), and Mikoshiba et al. (1977) concluded that secondary infection from a single disease cycle is limited to within a radius of about 42 meters from the inoculum source. Semangoen (1970) reported that conidia lost their ability to infect after 10-hour storage in saturated air in petri dishes, but at least a few conidia remained viable for 20 hours in saturated air on young corn leaves.

Symptoms/Signs White to yellow streaks, which become necrotic and brown, are the characteristic leaf symptoms of P. maydis. The fungus may become systemic, causing severe chlorosis in the upper leaves (Fig. 1.). Infected plants may be stunted and sterile and often lodge. Plants may develop multiple and deformed cobs, leaflike tassels and cobs, combined tassels and cobs, and either elongated or shortened stalks. Downy growth on the chlorotic streaks is common. Plants more than 4 weeks old are highly resistant to infection (Smith and Renfro, 1999).

Figure 1. Maize plants growing on the bunds of rice fields in West Java, showing typical chlorosis of systemic downy mildew. Photos courtesy of Rob Williams, CAB International. www.cabicompendium.org.

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Peronosclerospora maydis Primary Pest of Corn Fungal-like Java downy mildew

Pest Importance Downy mildew of sorghum, maize, and sugarcane are among the world’s most destructive diseases (Bonde et al., 1992). Heaviest losses are incurred when the disease becomes systemic (spreads throughout the plant host) (Kenneth, 1981). Java downy mildew is of great importance in Indonesia, where yield may be reduced by 40%. The most severe damage occurs when corn is planted late or the rainy season begins early, especially if the crop has been over fertilized with nitrogen or planted after corn or sugarcane (Smith and Renfro, 1999). Semangoen (1970) reported nearly 100% loss of late-planted maize in Java, Indonesia.

Known Hosts Saccharum spontaneum (wild sugarcane), Sorghum arundinaceum (wild sorghum), Sorghum plumosum (plume sorghum), Zea mays (corn), and Zea mexicana (teosinte). are reported hosts of P. maydis (Ramsey and Jones, 1988). Smith and Renfro (1999) indicate that Tripsacum, Euchlaena, and Pennisetum spp. are also hosts of P. maydis by inoculation.

Known Vectors (or associated insects) P. maydis is not a known vector and does not have any associated organisms.

Known Distribution Asia: China, India, Indonesia, Israel, Japan, and Thailand. Africa: Congo Democratic Republic. Caribbean: Jamaica. South America: Argentina. Oceania: Australia.

Reports from Africa are thought to be erroneous (Semangoen, 1970; Kenneth, 1976).

Potential Distribution within the United States This pathogen is expected to pose a relatively low threat to maize in the United States, because most of the crop is seasonally planted with periods of fallow between planting and harvest that will break the disease cycle. However, most of the corn growing regions of the United States have climatic conditions that would support plant infection, particularly during the months of May and June. A recent risk analysis by USDA- APHIS-PPQ-CPHST indicates that portions of Illinois, Indiana, Kentucky, and Ohio have the greatest risk for P. maydis establishment based on host availability and climate within the continental United States. The remaining states have low to moderate levels of risk for establishment of P. maydis.

Survey CAPS-Approved Method: Use visual survey, sentinel plots, spore trapping, or a combination of methods. For visual survey collect symptomatic plants. Spore traps, similar to those used in soybean rust monitoring, can be used to detect spores. Unsprayed, susceptible plants (sentinel plots) that are scouted regularly can also be used for early detection.

Literature-Based Methods:

205 Peronosclerospora maydis Primary Pest of Corn Fungal-like Java downy mildew

Visual survey: Surveys should occur in areas of the country that are at the greatest risk for establishment. Survey for Java downy mildew is conducted via visual survey of plants for symptoms of the disease (white to yellow streaks on leaves, stunted plants, sterile plants or those with multiple or deformed cobs) and signs of the pathogen (downy growth on underside of leaves consisting of conidia and conidiophores).

Trapping: Spore trapping using Burkhard spore traps and sentinel plots (unsprayed, susceptible plants that are scouted regularly) are suggested for early detection and have been employed for resistance screening for other exotic downy mildews (Cardwell et al., 1997; Expert Panel, 2006).

Key Diagnostics CAPS-Approved Method: Confirmation of P. maydis is by morphological identification. Pathogen may be identified morphologically by conidiophore structure and dimension and spore (conidia) shape and size. Isozyme comparisons have been used to identify Peronosclerospora spp., including P. maydis

Literature-Based Methods: Peronosclerospora spp. and other downy mildew genera (including Sclerospora and Sclerophthora) are primarily differentiated by pathogen morphology, including conidiophore structure and dimension and spore (conidia) shape and size. However, these characteristics can vary considerably under different culture conditions, at different developmental stages, and on different hosts. Identification of Peronosclerospora species often is difficult. They may be easily divided into three categories according to the shape of conidia: globose, ovoid to slightly elongate, and long or slipper-shaped, but within each group there are usually only minor morphological differences. Species are differentiated only by variations in the size and shape of their conidia and conidiophores, and sometimes differences in host ranges, presence or lack of oospores, and differences in morphology (Bonde et al., 1992). Characters for downy mildew pathogens are listed in Appendix A.

Isozyme comparisons have been used to identify Peronosclerospora spp., including P. maydis (Micales et al., 1988).

Some DNA-based approaches have been reported for other Peronosclerospora spp., which indicates that a molecular method for identification may be available in the future. Yao et al. (1991a) created a DNA clone that could be used as a species-specific hybridization probe for the detection and identification of P. sorghi. Yao et al. (1991b) then developed a DNA probe from a P. maydis genomic library to detect P. sacchari in maize leaves and seeds. Ladhalakshmi et al. (2009) used a DNA sequence characterized amplified region (SCAR) marker for identification of isolates of P. sorghi from maize using PCR. There has been some work on SSR markers for P. sorghi and related species that include this pathogen and will be helpful for identification and future PCR primer development (Perumal et al., 2008).

Easily Confused Pests

206 Peronosclerospora maydis Primary Pest of Corn Fungal-like Java downy mildew

Downy mildews on corn are caused by up to ten different species of Oomycete fungi in the genera Peronosclerospora, Scleropthora, and Sclerospora. Peronosclerospora maydis may be confused with other Peronosclerospora spp. occurring on corn and other downy mildew genera. Other indigenous downy mildews (e.g., P. sorghii) and physiological conditions (fertility, weather, etc.) can cause similar symptoms.

References Bonde, M.R. 1982. Epidemiology of downy mildew diseases of maize, sorghum, and pearl millet. Trop. Pest. Manage. 28(1): 49-60.

Bonde, M.R., Peterson, G.L., Kenneth, R.G., Vermeulen, H.D., Sumartini, and Bustaman, M. 1992. Effect of temperature on conidial germination and systemic infection of maize by Peronosclerospora species. Phytopathology 82(1): 104-109.

Butler, E.J. 1913. The downy mildew of maize (Sclerospora maydis). Mem. Dep. Agric. India Bot. Serv. 5: 275-280.

Cardwell, K.F., Kling, J.G., and Bock, C. 1997. Methods for screening maize against downy mildew Peronosclerospora sorghi. Plant Breeding 116: 221-226.

Expert Panel. 2006. Recovery plan for Philippine downy mildew and brown stripe downy mildew of corn caused by Peronosclerospora philippinensis and Sclerophthora rayssiae var. zeae, respectively.

http://www.ars.usda.gov/SP2UserFiles/Place/00000000/opmp/Corn%20Downy%20Mildew%2009H -18- 06.pdf.

Inaba, T., Hina, T., and Kajiwara, T. 1980. Appearance of sporulation ability after emergence of leaves infected with Java corn downy mildew fungus, Peronosclerospora maydis. Ann. Phytopath. Soc. Japan 46: 126-131.

Kenneth, R.G. 1976. The downy mildews of corn and other Gramineae in Africa and Israel, and the present state of knowledge and research. Kasetsart Journal 10: 148-159.

Kenneth, R.G. 1981. Downy mildew of Graminaceous crops. In The Downy Mildews. Spencer, D.M. (ed.). pgs. 367-394.

Ladhalakshmi, D., Vijayasamundeeswari, A., Paranidharan, V., Samiyappan, R., and Velazhahan, R. 2009. Molecular identification of isolates of Peronosclerospora sorghi from maize using PCR-based SCAR marker. World J. Microbiol. Biotechnol. 25: 2129-2135.

Micales, J.A., Bonde, M.R., and Peterson, G.L. 1988. Isozyme analysis and aminopeptidase activities within the genus Peronosclerospora. Phytopathology 78:1396-1402.

Mikoshiba, H., Mas Sudjudi, and Soadiarto, A. 1977. Dispersion of conidia of Sclerospora maydis in outbreaks of maize downy mildew disease in Indonesia. Japanese Agricultural Research Quarterly 11(3): 186-189.

Perumal, R., Nimmakayala, P. Erattaimuthu, S.R., No, E. –G., Reddy, U.K., Prom, L.K., Odovdy, G.N., Luster, D.G., and Magill, C.W. 2008. Simple sequence repeat markers useful for sorghum downy mildew (Peronosclerospora sorghi) and related species. BMC Genetics 9: 77.

Raciborski, M. 1897. Lijer, eine gefahrliche Maikrankheit. Ber. Dtsch. Bot. Ges. 15: 475.

207 Peronosclerospora maydis Primary Pest of Corn Fungal-like Java downy mildew

Ramsey, M.D., and Jones, D.R. 1988. Peronosclerospora maydis found on maize, sweetcorn, and plume sorghum in Far North Queensland. Plant Pathology 37: 581-587.

Semangoen, H. 1970. Studies on downy mildews of maize in Indonesia with special reference to the perennation of the fungus. Indian Phytopathology 23: 307-320.

Smith, D.R. and Renfro, B.L. 1999. Pg. 26, 28. In: D. G. White (ed.) Compendium of Corn Diseases, 3rd ed. APS Press, St. Paul, MN. 78 pp.

Tantera, D.M. 1975. Cultural practices to decrease losses due to corn downy mildew disease. Tropical Agricultural Research 8: 165-175.

Yao, C.L., Magill, C.W., and Frederickson, R.A. 1991a. Use of an A-T-rich DNA clone for identification and detection of Peronosclerospora sorghi. Applied and Environmental Microbiology 57(7): 2027-2032.

Yao, C.L., Magill, C.W., Frederickson, R.A., Bonde, M.R., Wang, Y., and Wu, P. 1991b. Detection and identification of Peronosclerospora sacchari in maize by DNA hybridization Phytopathology 81: 901-905.

208 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

Peronosclerospora philippinensis

Scientific Name Peronosclerospora philippinensis (W. Weston) C.G. Shaw

Synonyms: Peronosclerospora sacchari, Sclerospora indica, Sclerospora maydis, and Sclerospora philippinensis

Common Name Philippine downy mildew, Java downy mildew of corn, and sugarcane downy mildew

Type of Pest Fungal-like

Taxonomic Position Phylum: Oomycota, Class: Oomycetes, Order: Sclerosporales, Family: Sclerosporaceae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009; Agricultural Bioterrorism Protection Act of 2002 (Select Agent)

Pest Description Philippine downy mildew, caused by Peronosclerospora philippinensis, was initially reported from India in 1913. The pathogen at this time, however, was misidentified as Sclerospora maydis and later as S. indica (Bonde, 1982). The disease in India has never caused large losses (Payak, 1975). In contrast, in the Philippines, where the disease has been present at least since 1916 (Weston, 1920), it is economically very damaging. Philippine downy mildew is confined to parts of Asia and has not been reported within the United States.

Peronosclerospora philippinensis is an obligate parasite that will not grow on artificial media. The mycelia are branched, slender (8 µm in diameter), irregularly constricted, and inflated. Figure 1. Chlorotic symptoms of Erect conidiophores (15-26 x 150-400 µm) Philippine downy mildew. Photo grow out of stomata and are dichotomously courtesy of CIMMYT . branched two to four times. Branches are

209 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

robust. Sterigmata are ovoid to subulate, slightly curved, and 10 µm long. The conidia (17-21 x 27-39 µm) are elongate ovoid to round cylindrical, hyaline, and slightly rounded at the apex (Weston, 1920; Smith and Renfro, 1999). Haustoria are simple, vesiculiform to subdigitate, 8 x 2 µm.

Oospores are rarely produced and are not produced in corn tissue (Smith and Renfro, 1999). When produced, oospores are spherical, smooth-walled, approximately 22 µm in diameter; they germinate by a side germ tube (CABI, 2007).

Biology and Ecology Disease spreads locally via wind and rain from an infected crop. Perennial grass hosts may also serve as reservoir hosts to carry over the pathogen during unfavorable periods, or provide primary inoculum (Bonde, 1982). The known infective agents are the mycelium in these plants or the airborne conidia produced by the mycelium. Although conidia are produced from 18-23°C (64-73°F) and germinate from 10-35°C (50-95°F), the highest rates of infection occur at temperatures greater than 16°C (61°F) (Bonde et al., 1992). Dalmacio and Raymundo (1972) indicated that conidial production, germination, and infection required night time temperatures of 21-26°C (70-79°F). Movement of infected plant tissue could introduce Philippine downy mildew to new locations. Although Philippine downy mildew is reported to produce an over-wintering spore form (oospore), its role in the lifecycle has not been established.

In the host, the mycelium produces the conidiophores, which bear the conidia. The conidiophores emerge through host stomata in the chlorotic areas on both leaf surfaces (more so on the lower surface of corn, because of the higher density of stomata), and on sheaths, tassel rachis, glumes, and husks (Dalmacio and Exconde, 1969). The pathogen spreads intercellularly through the mesophyll cells. The fungus grows mainly downward through the leaf sheath to the stem where it moves into and persists in the shoot apex. When mycelium invades the meristematic tissues, chlorotic streaks soon appear on the leaves, followed by the fungus sporulating in these areas when conditions are favorable, producing secondary inoculum (Dalmacio and Exconde, 1969). The fungus can spread throughout the plant, but it is confined to the chlorotic, (not green) areas.

Spore production requires high humidity, with at least a thin film of water for 4-5 hours in the darkness on the infected leaf surface (Exconde, 1970; Dalmacio and Raymundo, 1972; Bonde, 1982). Germinating conidia produce germ tubes, which invade stomata. A mycelium develops in the mesophyll. Conidia form sparsely the morning after the first symptom appears, and abundantly after systemic symptoms appear (Dalmacio and Exconde, 1969). Sporulation may begin at midnight and continue until surface moisture dries (Weston, 1920). Sporulation on individual plants can continue for more than 2 months, releasing enormous number of conidia (Bonde, 1982). Once exposed, however, to drying conditions (such as sun, wind, or low humidity for 1-2 hours) the conidia shrivel and no longer germinate (Weston, 1920). Viable conidia germinate in less than 1 hour under favorable conditions. Penetration of the host occurs about 2 hours after inoculation (Dalmacio and Exconde, 1969).

210 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

Seedborne transmission may occur in corn, but there are no external symptoms on the seed. Transmission does not occur when seed moisture content is less than 14% (Exconde, 1976). In the laboratory, corn seed from infected plants produce infected seedlings, especially when hard dough kernels are planted soon after harvest. Because infected plants mature slower that uninfected plants, ears from the former are left in the field while the latter are harvested. The infected ears may then become a new inoculum source. No infections result from inoculations of healthy seed with conidia. The infective agent is the mycelium within the infected seed, reportedly located in the pericarp (Advincula and Exconde, 1976) or the embryo (Bains and Jhooty, 1982).

The number of systemically infected (non-yielding) plants is positively correlated with night time relative humidity, spore production, day time relative humidity and rainfall and negatively correlated with night and day temperatures and duration of sunlight (Bonde, 1982; Exconde, 1976).

Symptoms/Signs On corn, the first symptoms typically appear as chlorotic stripes at the first leaves as early as 9 days after planting. All leaves on a plant may show characteristic symptoms of long chlorotic (yellow) streaks (Fig. 1). Weston (1920) reported the collapse of badly infected cells and the destruction of chloroplasts, resulting in the characteristic yellow color of diseased leaves.

A downy (grayish) covering primarily on the underside of the leaves is characteristic beginning at the two-leaf stage and is present until the appearance of tassels and silks. This covering is the site of spore production (conidia on conidiophores) and the source for secondary spread of the disease to other susceptible plants.

As the plant ages, leaves may narrow, become abnormally erect, and appear somewhat Figure 2. Downy growth characteristic of downy dried-out. As the corn plant mildews. Photo of downy mildew of tobacco. Photo matures, tassels become courtesy of Tom Creswell, North Carolina State malformed and produce less University. pollen, ear formation is interrupted, and sterility of seeds can result (Expert Panel, 2006). If infection occurs early, plants are stunted and may die (Smith and Renfro, 1999).

211 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

There are no external symptoms on seeds. The fungus becomes established in the pericarp layer in the form of mycelium. The fungus is also present in the embryo and endosperm. There are no reports on the effect of P. philippinensis on seed quality (CABI, 2007).

Sugarcane has some of these symptoms.

Pest Importance Downy mildew of sorghum, maize, and sugarcane are among the world’s most destructive diseases (Bonde et al., 1992). Heaviest losses are incurred when the disease becomes systemic (spreads throughout the plant host) (Kenneth, 1981). P. philippinensis is considered the most virulent of the maize downy mildew pathogens (Payak, 1975). This obligate pathogen causes a serious disease to two major crops, corn and sugarcane. Yield losses of 40-60% and 25% have been observed in corn and sugarcane respectively (USDA, 1986). The national yield loss for corn in the Philippines in the 1974-1975 crop year was estimated at 8%, valued at US $23,000,000 (Exconde, 1976). No grain is produced because the seedlings quickly die or the plant slowly matures but produces little grain (Weston, 1920). For a susceptible sugar cane clone, 25-85% disease incidence was observed with losses in tons of cane per hectare ranging from 9 to 38%, losses in picul of sugar per ton from 2 to 35%, and losses in picul of sugar per hectare ranging from 10-58% (Husmillo, 1982).

Known Hosts Major hosts Saccharum officinarum (sugarcane) and Zea mays (corn)

Secondary hosts Avena spp. (oats) and Sorghum bicolor (sorghum)

Wild hosts Andropogon spp. (blue stem), Botriochloa spp. (bluestem), Euchlaena luxurians (Florida teosinte), Eulalia spp. (eulalia), Miscanthus spp., Saccharum spontaneum (wild sugarcane; kans grass), Sorghum halepense (Johnson grass), Sorghum plumosum (plum sorghum), Tripsacum spp. (gamagrasses), Zea diploperennis (diploperennial teosinte), Zea mexicana (teosinte), Zea perennis (perennial teosinte).

Known Vectors (or associated insects) There is a synergistic relationship between downy mildew fungi and maize streak virus (MSV) on corn. Infection by MSV can mask symptoms of downy mildew infection. Reduction in height and biomass were significantly greater with pathogen combinations than with single pathogens (Damsteegt et al., 1993).

Known Distribution Asia: China, India, Indonesia, Japan, Nepal, Pakistan, Philippines, and Thailand. Africa: Democratic Republic of Congo, Mauritius, and South Africa.

212 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

Potential Distribution within the United States A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that portions of Illinois, Indiana, Kentucky, and Ohio have the greatest risk for P. philippinensis establishment based on host availability and climate within the continental United States. The remaining states have low to moderate levels of risk for establishment of P. philippinensis.

Survey CAPS-Approved Method: Use visual survey, sentinel plots, spore trapping, or a combination of methods. For visual survey collect symptomatic plants. Spore traps, similar to those used in soybean rust monitoring, can be used to detect spores. Unsprayed, susceptible plants (sentinel plots) that are scouted regularly can also be used for early detection.

Literature-Based Methods: Visual survey: Surveys should occur in areas of the country that are at the greatest risk for establishment. Survey for Philippine downy mildew is conducted via visual survey of plants for symptoms of the disease (chlorotic stripes or streaks) and signs of the pathogen (downy growth on underside of leaves consisting of conidia and conidiophores). Survey for diseased plants by examining corn seedlings that have larger and larger yellow streaks on succeeding leaves. Grayish white down will be in the chlorotic areas although sun or low humidity may have dried the mildew to matted fragments (USDA, 1986).

Trapping: Spore trapping using Burkhard spore traps and sentinel plots (unsprayed, susceptible plants that are scouted regularly) are suggested for early detection and have been employed for resistance screening for other exotic downy mildews (Cardwell et al., 1997; Expert Panel, 2006).

Key Diagnostics CAPS-Approved Method: Confirmation of P. philippinensis is by morphological identification. Pathogen may be identified morphologically by conidiophore structure and dimension and spore (conidia) shape and size. Isozyme comparisons have been used to identify Peronosclerospora spp., including P. philippinensis

Literature-Based Methods: Peronosclerospora spp. and other downy mildew genera (including Sclerospora and Sclerophthora) are primarily differentiated by pathogen morphology, including conidiophore structure and dimension and spore (conidia) shape and size. However, these characteristics can vary considerably under different culture conditions, at different developmental stages, and on different hosts. Identification of Peronosclerospora species often is difficult. They may be easily divided into three categories according to the shape of conidia: globose, ovoid to slightly elongate, and long or slipper-shaped, but within each group there are usually only minor morphological differences. Species are differentiated by only variations in the size and shape of their conidia and conidiophores, and sometimes differences in host ranges,

213 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

presence or lack of oospores, and differences in morphology (Bonde et al., 1992). Characters for downy mildew pathogens are listed in Appendix A.

Isozyme comparisons have been used to identify Peronosclerospora spp., including P. philippinensis (Bonde et al., 1984; Micales et al., 1988). P. philippinensis and P. sacchari can not be separated based on isozyme analysis. These species are believed to be conspecific (Yao et al., 1991b).

Some DNA-based approaches have been reported for other Peronosclerospora spp., which indicates that a molecular method for identification may be available in the future. Yao et al. (1991a) created a DNA clone that could be used as a species-specific hybridization probe for the detection and identification of P. sorghi. Yao et al. (1991b) then developed a DNA probe from a P. maydis genomic library to detect P. sacchari in maize leaves and seeds. Ladhalakshmi et al. (2009) used a DNA sequence characterized amplified region (SCAR) marker for identification of isolates of P. sorghi from maize using PCR. There has been some work on SSR markers for P. sorghi and related species that include this pathogen and will be helpful for identification and future PCR primer development (Perumal et al., 2008).

Easily Confused Pests Downy mildews of corn are caused by up to ten different species of Oomycete fungi in the genera Peronosclerospora, Scleropthora and Sclerospora. Peronosclerospora philippinensis may be confused with other Peronosclerospora spp. occurring on corn and other downy mildew genera. Other indigenous downy mildews (e.g., P. sorghii) and physiological conditions (fertility, weather, etc.) can cause similar symptoms.

References Advincula, B.A. and Exconde, O.R. 1976. Seed transmission of Sclerospora philippinensis Weston in maize. Philpp. Agric. 59: 244-245.

Bains, S.S. and Jhooty, J.S. 1982. Distribution, spread, and perpetuation of Peronosclerospora philippinensis in Punjab. Indian Phytopathology 35(4): 566-570.

Bonde, M.R. 1982. Epidemiology of downy mildew diseases of maize, sorghum, and pearl millet. Trop. Pest. Manage. 28(1): 49-60.

Bonde M.R., Peterson, G.L., Dowler, W.M., and May, B. 1984. Isozyme analysis to differentiate species of Peronosclerospora causing downy mildews of maize. Phytopathology 74(11): 1278-1283.

Bonde, M.R., Peterson, G.L., Kenneth, R.G., Vermeulen, H.D., Sumartini, and Bustaman, M. 1992. Effect of temperature on conidial germination and systemic infection of maize by Peronsclerospora species. Phytopathology 82(1): 104-109.

CABI. 2007. Crop Protection Compendium Wallingford, UK: CAB International.

http://www.cabi.org/compendia/cpc/H .

Cardwell, K.F., Kling, J.G., and Bock, C. 1997. Methods for screening maize against downy mildew Peronosclerospora sorghi. Plant Breeding 116: 221-226.

214 Peronosclerospora philippinensis Primary Pest of Corn Fungal-like Philippine downy mildew

Dalmacio, S.C. and Exconde, O.R. 1969. Penetration and infection of Sclerospora philippinensis Weston on corn. Philippine Agriculturist 53: 35-52.

Dalmacio, S.C. and Raymundo, A.D. 1972. Spore density of Sclerospora philippinensis in relation to field temperature, relative humidity, and downy mildew incidence. Philippine Phytopathology 8: 72-77.

Damsteegt, V.D., Bonde, M.R., and Hewings, A.D. 1993. Interactions between maize streak virus and downy mildew fungi in susceptible maize cultivars. Plant Disease 77: 390-392.

Exconde, O.R. 1970. Philippine corn downy mildew. Indian Phytopathology 23: 275-284.

Exconde, O.R. 1976. Philippine corn downy mildew: assessment of present knowledge and future research needs. Kasetsart Journal 10: 94-100.

Expert Panel. 2006. Recovery plan for Philippine downy mildew and brown stripe downy mildew of corm

caused by Peronosclerospora philippinensis and Sclerophthora rayssiae var. zeae, respectively. H http://www.ars.usda.gov/SP2UserFiles/Place/00000000/opmp/Corn%20Downy%20Mildew%2009-18- 06.pdf.

Husmillo, F.R.1982. Assessment of yield loss due to downy mildew of sugarcane caused by Peronosclerospora philippinensis (Weston) C.G. Shaw. Sugarcane Pathol. Newsl. 28: 17-24.

Kenneth, R.G. 1981. Downy mildew of Graminaceous crops. In The Downy Mildews. Spencer, D.M. (ed.). pgs. 367-394.

Ladhalakshmi, D., Vijayasamundeeswari, A., Paranidharan, V., Samiyappan, R., and Velazhahan, R. 2009. Molecular identification of isolates of Peronosclerospora sorghi from maize using PCR-based SCAR marker. World J. Microbiol. Biotechnol. 25: 2129-2135.

Micales, J.A., Bonde, M.R., and Peterson, G.L. 1988. Isozyme analysis and aminopeptidase activities within the genus Peronosclerospora. Phytopathology 78: 1396-1402.

Payak, M.M. 1975. Epidemiology of maize downy mildews with special reference to those occurring in Asia. Tropical Agriculture Research 8: 81-91.

Perumal, R., Nimmakayala, P. Erattaimuthu, S.R., No, E. –G., Reddy, U.K., Prom, L.K., Odovdy, G.N., Luster, D.G., and Magill, C.W. 2008. Simple sequence repeat markers useful for sorghum downy mildew (Peronsclerospora sorghi) and related species. BMC Genetics 9: 77.

Smith, D.R. and Renfro, B.L. 1999. Pg. 26, 28. In: D. G. White (ed.) Compendium of Corn Diseases, 3rd ed. APS Press, St. Paul, MN. 78 pp.

USDA. 1986. Pests not known to occur in the United States or of limited distribution. No. 77: Philippine downy mildew.

Weston, W.H. 1920. Philippine downy mildew of maize. J. Agric. Res., 19: 97-122.

Yao, C.L., Magill, C.W., and Frederickson, R.A. 1991a. Use of an A-T-rich DNA clone for identification and detection of Peronosclerospora sorghi. Applied and Environmental Microbiology 57(7): 2027-2032.

Yao, C.L., Magill, C.W., Frederickson, R.A., Bonde, M.R., Wang, Y., and Wu, P. 1991b. Detection and identification of Peronosclerospora sacchari in maize by DNA hybridization Phytopathology 81: 901-905.

215 Scleropthora rayssiae var. zeae Primary Pest of Corn Fungal-like Brown stripe downy mildew

Sclerophthora rayssiae var. zeae

Scientific Name Sclerophthora rayssiae var. zeae Payak & Renfro

Common Name(s) Brown stripe downy mildew, brown stripe, and maize downy mildew

Type of Pest Fungal-like

Taxonomic Position Phylum: Oomycota, Class: Oomycetes, Order: Sclerosporales, Family: Verrycalvaceae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009; Agricultural Bioterrorism Protection Act of 2002 (Select Agent)

Pest Description Brown stripe downy mildew is caused by Sclerophthora rayssiae var. zeae. S. rayssiae var. zeae is an obligate parasite that will not grow on artificial media. Sporangia develop at 22-25°C (72-77°F) atop determinate and short sporangiophores that grow from the substomatal spaces. Sporangia are hyaline and ovate to almost cylindrical, operculate 18.5-26 × 29-66.5 and germinate via zoospores at 20-22°C (68-77°F). Sporangia are produced in clusters of two to six and are hyaline, ovate to cylindrical, with an obvious peduncle. Sporangia walls are smooth and contain four to eight zoospores. Zoospores geminate at 15-30°C (59-86°F). Once encysted, the zoospores are hyaline, spherical and range from 7.5-11 µm (Payak and Renfro, 1967; Smith and Renfro, 1999).

The oogonia (33-44.5 µm in diameter) are subglobose, thin walled, and hyaline to light straw colored. Oospores (29.5-37 µm in diameter) are spherical to subspherical with smooth, glistening walls that are 4 µm thick and confluent with the oogonia. They have hyaline contents, including a prominent oil globule. The oospore stage in corn develops more readily and more abundantly where higher temperatures prevail (over 28°C, 82°F) (Payak, 1975). Oospores and oogonia are numerous and scattered in the leaf mesophyll or under stomata (Payak and Renfro, 1967; Smith and Renfro, 1999).

Biology and Ecology The disease is primarily soilborne, but can be seedborne. Young plants are most susceptible and susceptibility decreases as plants age. Plants may be predisposed to infection when zinc is limiting. Soil temperatures of 28-32°C favor disease development. It has also been shown that the disease severity varies with the rainfall. Where the annual rainfall varies from 4-60 cm the disease intensity was described as low, moderate in areas with 60-100 cm, and high in regions receiving 100-200 cm of rain.

216 Scleropthora rayssiae var. zeae Primary Pest of Corn Fungal-like Brown stripe downy mildew

Oospores survive in infected debris in the soil. The infective source responsible for initial outbreaks has been determined to be the oosporic stage (Payak, 1975). The pathogen may survive as oospores for as long as four years in infected plant debris in the soil (Singh, 1971a). When adequate moisture is present for at least 12 hours, the oospore germinates to produce a sporangiophore bearing a sporangium that liberates four to eight zoospores. In the presence of enough moisture or at high temperatures, the sporangium may produce a germ tube that can also infect corn leaves. Secondary spread is by sporangia. Pathogen dissemination is by rain water, and direct contact. Sporangia have been trapped 1.65 meters from an infected field, but the greatest numbers of sporangia were found to move less than 1 meter, suggesting that long distance transport via wind is unlikely (Singh and Renfro, 1971).

Moisture is essential for infection by S. rayssiae var. zeae. Sporangia production, germination, and infection require a film of water. Twelve hours of leaf wetness were required for infection via zoospores, with longer periods producing greater numbers of infected plants. Most sporangia are liberated at maturity during the day. Sporangial release occurs in the afternoon of sunny days when high moisture is present, rather than on cloudy or rainy days (Singh and Renfro, 1971). Generation time of secondary inoculum (sporangia) from primary inoculum (oospores) can be rapid. Under ideal conditions, sporangial production can occur as soon as 10 days post inoculation. Infected plants placed in a moist environment at 22 to 25°C can produce sporangia in as little as 3 hours, with a second generation of sporangia arising 9 hours later (Singh et al., 1970).

The fungus can be detected within the embryo of corn seeds and has been shown to be seed transmitted when seeds from infected plants are planted in sterile soil immediately after harvest (Singh et al., 1967a,b). Seed transmission was found to occur at less than 1% (Lal and Figure 1. Symptoms of brown stripe downy mildew. Image Prasad, 1989). Seeds courtesy C. De Leon. Reproduced from Compendium of Corn dried to 14% moisture Diseases, 3rd Ed., 1999, American Phytopathological Society, St. or less and stored for Paul, MN. 4 weeks or more do

217 Scleropthora rayssiae var. zeae Primary Pest of Corn Fungal-like Brown stripe downy mildew

not transmit this or other downy mildew diseases. The pathogen can also overseason in crabgrass as oospores or as mycelium from which sporangia are produced (Bains et al., 1978).

Symptoms/Signs S. rayssiae var. zeae causes leaf lesions only (Putnam, 2007). Initially, lesions develop on the leaves as narrow, chlorotic or yellowish stripes, similar to other downy mildews, but only 3-7 mm wide (Fig. 1). They have well-defined margins and are delimited by the veins. The stripes later become reddish to purple in some corn genotypes (Putnam, 2007). Lateral development of lesions causes severe striping and blotching. The disease may first be noticed on the lower leaves, which will show the greatest degree of striping; as a result they appear pale-brown and burnt, and severely affected leaves may be shed prematurely.

Seed development may be suppressed, seed may be smaller in size, and the plant may die prematurely if blotching occurs prior to flowering. Unlike other downy mildews, floral or vegetative parts are not malformed, and the leaves do not shred (Smith and Renfro, 1999; Putnam, 2007). The pathogen apparently does not systemically affect the plant.

Sclerophthora rayssiae var. zeae may sporulate on either side of the lesions and appear downy or woolly. Sporangia disappear as the lesions become necrotic. Oospores occur only in necrotic tissue, in the mesophyll, or beneath the stomata, but not in vascular tissue (Putnam, 2007).

Pest Importance Downy mildews of sorghum, maize, and sugarcane are among the world’s most destructive diseases (Bonde et al., 1992). Yield loss of up to 63% has been reported in India (Payak and Renfro, 1967). Singh (1971b) reported 20-70% infection in many corn growing areas in India with an incidence of 80-100% in certain high hilly tracts. Annual losses from this disease in India have been estimated in the million of dollars (Frederickson and Renfro, 1977). Maize genotypes vary in their reaction to S. rayssiae var. zeae (Payak and Renfro, 1967). Among 2113 Indian maize inbred lines and other germplasm scored in the field, 58 were highly resistant, 667 resistant, 772 moderately resistant, 478 susceptible, and 138 highly susceptible (Singh et al., 1970).

According to CABI (2007), losses due to the disease vary depending on when and how severely the tissue is affected. If three-quarters or more of the foliage is affected prior to flowering, then the loss may be total; ear formation is either totally suppressed or markedly attenuated. Grain yield reductions vary from 20-90%. Losses in the higher range only occur with highly susceptible cultivars in conditions conducive for disease development.

Known Hosts Digitaria bicornis (southern crabgrass), Digitaria sanguinalis (hairy crabgrass), Sorghum bicolor (sorghum), and Zea mays (corn).

218 Scleropthora rayssiae var. zeae Primary Pest of Corn Fungal-like Brown stripe downy mildew

The isolate of S. rayssiae var. zeae from D. bicornis was not able to infect maize (Singh et al., 1970). Additional studies are needed to clarify if the pathogen from D. bicornis could play a role in causing infection on corn.

Known Vectors (or associated insects) S. rayssiae var. zeae is not a known vector and does not have any associated organisms.

Known Distribution Asia: India, Myanmar, Nepal, Pakistan, and Thailand.

Potential Distribution within the United States Corn growing regions in the country with warm and wet early season growing conditions are suitable for disease development and would be at high risk for damage caused by this disease if the pathogen is introduced. A recent risk analysis by USDA-APHIS-PPQ- CPHST indicates that portions of Illinois, Indiana, Kentucky, and Ohio have the greatest risk for S. rayssiae var. zeae establishment based on host availability and climate within the continental United States. The remaining states have low to moderate levels of risk for establishment of S. rayssiae var. zeae.

Survey CAPS-Approved Method: Visual survey is the method to survey for S. rayssiae var. zeae. For visual survey collect symptomatic leaves.

Literature-Based Methods: Visual survey: Young plants should be visually inspected for those with narrow (3-7 mm wide) chlorotic stripes on leaves with well-defined margins and delimited by the veins. The stripes later turn reddish to purple in some genotypes. Check for downy growth on both sides of infected leaves on cool, damp mornings as the downy growth will usually disappear by late afternoon. Infected leaves remain intact.

Key Diagnostics CAPS-Approved Method: Confirmation of S. rayssiae var. zeae is by morphological identification. Pathogen may be identified morphologically by conidiophore structure and dimension and spore (conidia) shape and size.

Literature-Based Methods: Sclerophthora spp. and other downy mildew genera (including Sclerospora and Peronosclerospora) are primarily differentiated by pathogen morphology, including conidiophore structure and dimension and spore (conidia) shape and size, and if produced oogonia and oospores. However, these characteristics can vary considerably under different culture conditions, at different developmental stages, and on different hosts. Characters for downy mildew pathogens are listed in Appendix A.

219 Scleropthora rayssiae var. zeae Primary Pest of Corn Fungal-like Brown stripe downy mildew

There are no known serological or molecular diagnostic methods available. A standing operating procedure has been developed by the National Plant Diagnostic Network for diagnosing this pathogen.

Easily Confused Pests Downy mildews of corn are caused by up to ten different species of oomycete fungi in the genera Peronosclerospora, Sclerophthora and Sclerospora. S. rayssiae var. zeae may be confused with other Sclerophthora spp. occurring on corn and other downy mildew genera. Other indigenous downy mildews (e.g., P. sorghii) and physiological conditions (fertility, weather, etc.) can cause similar symptoms.

Brown stripe downy mildew could be confused with two potentially destructive downy mildew diseases of corn already established in the United States: crazy top (Sclerophthora macrospora) and sorghum downy mildew (Peronsclerospora sorghi) (USDA, 1984). In crazy top, growth in the tassel proliferates into a mass or tangle of leafy structures (phyllody), creating the ‘crazy top’ appearance. Also, numerous shoots or tillers sprout from the base of the original shoot. Phyllody and excessive tillering are not symptomatic of brown stripe downy mildew (USDA, 1984). Sorghum downy mildew causes infected plants to appear chlorotic and frequently stunted. White stripes may appear on the leaves. Also, the leaves are narrower and more erect than on healthy plants. Phyllodied tassels may be present. None of these symptoms are observed in brown stripe downy mildew (USDA, 1984).

Scleropthora rayssiae var. rayssiae causes a downy mildew disease in barley. This pathogen closely resembles S. rayssiae var. zeae, but they differ in host range (Payak et al., 1970; USDA, 1984).

References Bains, S.S., Jhooty, J.S., Sokhi, S.S., and Rewal, H.S. 1978. Role of Digitaria sanguinalis in outbreaks of brown stripe downy mildew of maize. Plant Disease Reporter 62(2): 143.

Bonde, M.R., Peterson, G.L., Kenneth, R.G., Vermeulen, H.D., Sumartini, and Bustaman, M. 1992. Effect of temperature on conidial germination and systemic infection of maize by Peronsclerospora species. Phytopathology 82(1): 104-109.

CABI. 2007. Crop Protection Compendium Wallingford, UK: CAB International.

http://www.cabi.org/compendia/cpc/H .

Frederiksen, R.A., and Renfro., B.L. 1977. Global status of maize downy mildew. Ann. Rev. Phytopathol. 15: 249-275.

Lal, S. and Prasad, T. 1989. Detection and management of seed-borne nature of downy mildew diseases of maize. Seeds Farms 15: 35-40.

Payak, M.M. 1975. Epidemiology of maize downy mildews with special reference to those occurring in Asia. Tropical Agriculture Research 8: 81-91.

Payak, M.M. and Renfro, B.L. 1967. A new downy mildew disease of maize. Phytopathology 57: 394- 397.

220 Scleropthora rayssiae var. zeae Primary Pest of Corn Fungal-like Brown stripe downy mildew

Payak, M.M., Renfro, B.L., and Lal, S. 1970. Downy mildew disease incited by Sclerophthora. Indian Phytopathol. 23(2): 183-193.

Putnam, M.L. 2007. Brown stripe downy mildew (Sclerophthora rayssiae var. zeae) of maize. Plant Health Progress doiL10.1094/PHP-2007-1108-01-DG. http://www.plantmanagementnetwork.org/pub/php/diagnosticguide/2007/stripe/.

Singh, J.P. 1971a. Infectivity and survival of oospores of Sclerophthora rayssiae var. zeae. Indian J. Exp. Biol. 9: 530-532.

Singh, J.P. 1971b. Pathological and histopathological studies of brown stripe downy mildew of maize. Indian J. Exp. Biol. 9(4): 493-495.

Singh, J.P., and Renfro, B.L. 1971. Studies on spore dispersal in Sclerophthora rayssiae var. zeae. Indian Phytopath. 24:457-461.

Singh, R.S, Chaube, H.S., Khanna, R.N., and Joshi, M.M. 1967a. Internally seedborne nature of two downy mildews of corn. Plant Disease Reporter 51: 1010-1012.

Singh, R.S., Joshi, M.M., and Chaube, H.S. 1967b. Further evidence of the seedborne nature of maize downy mildews and their possible control with chemicals. Plant Disease Reporter 52: 446-449.

Singh, J.P., Renfro, B.L., and Payak, M.M. 1970. Studies on the epidemiology and control of brown stripe downy mildew of maize (Sclerophthora rayssiae var. zeae). Indian Phytopath. 23: 194-208.

Smith, D.R. and Renfro, B.L. 1999. Pg. 26, 28. In: D. G. White (ed.) Compendium of Corn Diseases, 3rd ed. APS Press, St. Paul, MN. 78 pp.

USDA. 1984. Pests not known to occur in the United States or of limited distribution, No. 55: Brown stripe downy mildew. USDA-APHIS-PPQ.

221 Appendix A: Downy mildew morphological identification

Secondary Pests of Corn (Truncated Pest Datasheet) None at this time

Appendix A: Characteristics of the downy mildew fungi found on corn and other hosts Taken from Smith and Renfro (1999)

PATHOGEN PERONOSCLEROSPORA SCLEROSPORA SCLEROPTHORA philippinensis maydis (Java sacchari sorghi graminicola macrospora rayssiae (Philippine downy (sugarcane (sorghum (Graminicola (crazy top) var. zeae downy mildew) downy downy downy mildew) (brown mildew) mildew) mildew) stripe downy mildew) Conidiophores Hyaline, Hyaline, Hyaline, Hyaline, Hyaline, length Hyaline, Hyaline, length 150- length 150- length 160- length 180- av. 268 µm, length av. short, 400 µm, 550 µm, 170 µm, 300 µm, bloated, non- 13.8 µm, hyphoid, bloated, bloated, bloated, bloated, often septate, simple, determinate widening dichotomously widening dichotomously irregularly hyphoid, abruptly, branched 2-4 gradually, branched 2-3 dichotomously determinate dichotomously times, dichotomously times, septate branched branched 2-4 ephemeral branched 2-3 near base, ephemeral times, times, ephemeral ephemeral ephemeral Asexual Conidia Conidia Conidia Conidia Sporangia Sporangia Sporangia spores hyaline, hyaline, hyaline, hyaline, oval hyaline, broadly hyaline, hyaline, elongate- spherical to elliptical, to almost elliptical, lemon- ovate to ovoid to subspherical, oblong or spherical, 15- operculate, shaped, almost round- 17-23 x 27-39 conical, apex 26.9 x 15-28.9 papillate, 12-21 x operculate, cylindrical, cylindrical, µm round, 15-239 µm 14-31 µm papillate, operculate, apex slightly x 25-41 µm 30-60 x 60- 18.5-26 x rounded, 17- 100 µm 29-66.5 µm 21 x 27-39 µm Asexual Germ tube Germ tube Germ tube Germ tube Zoospores Many Zoospores spores zoospores germinate by

222 Appendix A: Downy mildew morphological identification

PATHOGEN PERONOSCLEROSPORA SCLEROSPORA SCLEROPTHORA philippinensis maydis (Java sacchari sorghi graminicola macrospora rayssiae (Philippine downy (sugarcane (sorghum (Graminicola (crazy top) var. zeae downy mildew) downy downy downy mildew) (brown mildew) mildew) mildew) stripe downy mildew) Oospores Rare or Unknown Yellow to Usually brown Pale brown, Hyaline to Brown, nonexistent; yellow brown, to subhyaline spherical, usually pale yellow, spherical, spherical, globular to spherical, smooth-walled, mainly in 29.5-37 µm smooth slightly diameter 25- diameter 22.5-35 vascular in diameter walled, 22.6 angular; 40- 42.9 µm µm bundles, µm in 50 µm in diameter diameter diameter 45-75 µm Sexual spores Side germ - Germ tube Wide germ Germ tube or by Sporangia Sporangia germinate by tube tube sporangia Geographical Philippines, Australia, Australia, Fiji North America Worldwide, but North India, Distribution Indonesia, Indonesia Islands, (including the on corn found America Nepal, India, Nepal, Japan, Nepal, U.S.), Central only in Israel and (including Pakistan, China, New Guinea, America, the United States the U.S.), Thailand Thailand, India, Asia, Africa, Mexico, Pakistan Philippines, South South Taiwan, America, America, Thailand Australia, Europe, Europe Africa, Asia

223 Appendix B: Diagnostic resource contacts

Appendix B: Diagnostic Resource Contacts

National Identification Services:

Joseph Cavey National Identification Services, Branch Chief USDA, APHIS, PPQ 4700 River Road, Unit 133 Riverdale, MD 20737 Office: (301) 734-8547 Fax: (301) 734-5276 [email protected]

Joel P. Floyd National Identification Services, Domestic Diagnostics Coordinator USDA, APHIS, PPQ 4700 River Road, Unit 52 Riverdale, MD 20737 Office: (301) 734-4396 Fax: (301) 734-5276 [email protected]

Domestic Identifiers:

Western Region Craig A. Webb, Ph.D. Plant Pathologist - Domestic Identifier USDA, APHIS, PPQ Department of Plant Pathology Kansas State University 4024 Throckmorton Plant Sciences Manhattan, Kansas 66506-5502 Office: (785) 532-1349 Cell: (785) 633-9117 Fax: (785) 532-5692 [email protected]

Kira Zhaurova Entomologist - Domestic Identifier USDA, APHIS, PPQ Minnie Belle Heep 216D 2475 TAMU College Station, TX 77843 Cell: (979) 450-5492 [email protected]

Eastern Region Julieta Brambila Entomology - Domestic Identifier USDA, APHIS, PPQ PO Box 147100 Gainesville, FL 32614-7100 Office: (352) 372-3505 Fax: (352) 494-5841

224 Appendix B: Diagnostic resource contacts

[email protected]

Grace O'Keefe Plant Pathologist - Domestic Identifier USDA, APHIS, PPQ 105 Buckhout Lab Penn State University University Park, PA 16802 Office: (814) 865-9896 Cell: (814) 450-7186 Fax: (814) 863-8265 [email protected]

Western and Eastern Region Robert (Bobby) Brown Forest Entomology - Domestic Identifier USDA, APHIS, PPQ Purdue University Smith Hall 901 W. State Street West Lafayette, IN 47907 Office: (765) 496-9673 Fax: (765) 494-0420 [email protected]

225 Appendix C: Glossary of terms

Appendix C: Glossary of Terms

Aedeagus: In male insects, the penis or intromittent organ, situated below the scaphium and enclosed in a sheath.

Aestivation: Dormancy in summer during periods of continued high temperatures, or during a dry season.

Aggressiveness: Relative ability of a plant pathogen to colonize and cause damage to plants (see virulence).

Allopatric: Occurring in separate, non-overlapping geographic areas. Often used to describe populations of related organisms unable to crossbreed because of geographic separation.

Androconia: Also called scent scales. Modified wing scales on butterflies and moths that release pheromones. Only males have these scent scales.

Anastomosis: Fusion between branches of the same or different structures (e.g. hyphae) to make a network.

Annual: A plant that completes its life cycle and dies within one year (see perennial).

Areolated: Divided into small spaces or areolations; usually pertains to the cuticle of a nematode.

Bifurcate: Split or divided into.

Biserrate: Having saw-like notches with the notches themselves similarly notched.

Blight: Sudden, severe, and extensive spotting, discoloration, wilting, or destruction of leaves, flowers, stems, or entire plants.

Bolls: The spherical shaped fruits of cotton and flax.

Botryoid: Resembling a cluster of grapes in form.

Bt: Bacillus thuringiensis. A gram-positive, soil-dwelling bacterium, commonly used as a pesticide.

Cerarii: These are characteristic of mealybugs and consist of groups of large setae, usually conical, on the lateral margins of the body.

Chlorotic: Abnormal condition of plants in which the green tissue loses its color or turns yellow as a result of decreased chlorophyll production due to disease or lack of light.

226 Appendix C: Glossary of terms

Chorion: The outer shell or covering of the insect egg.

Ciliated: Having a margin or fringe of hairlike projections.

Coalesce: Grow together into one body or spot.

Cockchafer: Any of various large European beetles destructive to vegetation as both larvae and adult.

Colonize: To infect and ramify through plant tissue with the growth of a pathogen.

Concave: Curving inward.

Conical: Relating to or resembling a cone.

Conidium (pl. conidia): An asexual, nonmotile fungal spore that develops externally or is liberated from the cell that formed it.

Conidiophores: Simple or branched on which conidia are produced.

Conspecific: An organism belonging to the same species as another organism.

Convex: Curving or bulging outward.

Cornuti: Spines.

Corona: A crown; the whorl of structures between the corolla and stamens.

Costa: Any elevated ridge that is rounded at its crest; the thickened anterior margin of any wing, but usually the forewings of an insect.

Cotyledon: Embryonic leaf in seed-bearing plants.

Coxae (pl. of coxa): The basal segment of the leg of an insect, by means of which it is articulated to the body.

Cremaster: 1) The apex of the last segment of the abdomen; 2) the terminal spine or hooked process of the abdomen of subterranean pupa, which is used to facilitate emergence from the earth; 3) an anal hook by which some pupae are suspended.

Cyst: In fungi, a resting structure in a protective membrane or shell-like enclosure. In nematodes, the egg-laden carcass of a female nematode. In bacteria, a specialized type of bacterial cell enclosed in a thick wall, often dormant and resistant to environmental conditions.

227 Appendix C: Glossary of terms

Damping off: Death of a seedling before or shortly after emergence due to decomposition of the root and/or lower stem; it is common to distinguish between preemergence damping-off and postemergence damping-off.

Dead hearts: Drying of the central shoot produced when stem borer larvae kill the growing points of young shoots.

Defoliation: Loss of leaves from a plant, whether normal or premature.

Degree Days: Development of poikilothermic ("cold-blooded") organisms such as insects, fungi, and plants, is regulated by environmental temperatures. Development to particular stages in the life cycles of these organisms is largely controlled by how much heat they experience, where heat is considered as a function of temperature and time. Degree-days are an estimate of the amount of heat accumulated over a 24-hr period. They are calculated using lower and upper developmental thresholds unique to a particular organism and, typically, some approximation of the 24-hour temperature pattern derived from minimum and maximum daily temperatures (which are commonly available from local weather-recording stations). Only those temperatures falling between the lower and upper thresholds are included in the calculations. Degree-day values may be positive or equal zero (all temperatures above or below thresholds), but are never negative. Degree-days are calculated for each day and are then summed to provide cumulative (total) degree-days. Starting points for calculating cumulative degree-days are usually arbitrary, typically January 1 but often later (e.g. April 1) in areas with cold winter temperatures. Based on experimental data, cumulative degree- days are linked to specific development events of interest (e.g. adult insect emergence). Thus, a pest manager can anticipate or predict an event of interest based on local temperature data and an appropriate degree-day based developmental model (Hansen, 2007).

Delimited: Bounded; having the limits or boundaries established.

Detasseling: The act of removing the pollen-producing tassel from a corn (maize) plant and placing it on the ground.

Deutonymph: The third instar of a mite.

Diapause: A period of arrested development and reduced metabolic rate, during which growth, differentiation, and metamorphosis cease; a period of dormancy not immediately referable to adverse environmental conditions.

Dichotomous: Divided or dividing into two sharply distinguished parts or classifications.

Diffuse: To pour out and cause to spread freely.

Digitus: Having appendages of the feet (as found in members of the family Coccidae), which may be either broadly dilated or knobbed hairs; tenent hairs, empodial hairs.

228 Appendix C: Glossary of terms

Discal: On or relating to the disc of any surface or structure.

Distal: Far from the center.

Diurnal: Of or belonging to or active during the day.

Downy: Like down or as soft as down.

Downy mildew: A plant disease in which the fungus appears as a downy growth on the host surface; caused by a member of the Oomycetes.

Ecdysis: Molting; the process of shedding the exoskeleton.

Eclosion: The emergence of an insect from the pupa case, or of a larva from the egg.

Ectoparasite: Parasite that feeds from the exterior of its host (see endoparasite).

ELISA (Enzyme-Linked ImmunoSorbent Assay): A serological test in which the sensitivity of the reaction is increased by attaching an enzyme that produces a colored product to one of the reactants.

Elutriation: The operation of pulverizing substances and mixing them with water in order to separate the heavier constituents, which settle out in solution, from the lighter constituents.

Embryo: A minute rudimentary plant contained within a seed or an archegonium.

Endoparasite: Parasitic organism that lives and feeds from inside its host (see ectoparasite).

Endosperm: Nutritive tissue surrounding the embryo within seeds of flowering plants.

Exogenous: Derived or originating externally.

External: Outside.

Facultative diapause: May or may not need to diapause; not required for development.

Fecundity: The number of offspring per number of potential offspring (e.g., eggs).

Fenestration: For nematodes, a clear area around the anus of the cyst.

Filiform: Thread-like or hair-like.

Forewing: Either of the anterior pair of wings on an insect that has four wings.

229 Appendix C: Glossary of terms

Frass: Plant fragments made by a wood-boring insect usually mixed with excrement; solid larval insect excrement.

Fuscous: Of something having a dusky brownish gray color.

Gall: An abnormal swelling or localized outgrowth, often roughly spherical, produced by a plant as a result of attack by a fungus, bacterium, nematode, insect, or other organism.

Genetalia: A sex organ.

Germ tube: Hypha resulting from an outgrowth of the spore wall and cytoplasm after germination.

Glume: Small dry membranous bract found in inflorescences of Graminaceae and Cyperaceae.

Gubernaculum: Nematodes: Spicule guide; sclerotized accessory piece.

Haustorium (pl. haustoria): Specialized branch of a parasite formed inside host cells to absorb .

Hard dough: A description the grain at maximum dry weight. The grain is hard to puncture and has started to dry off.

Hemispherical: Shaped like the half of a globe or sphere.

Hemizonid: Nematodes: Lens-like structure situated between the cuticle and hypodermal layer on the ventral side of the body just anterior to the excretory pore; generally believed to be associated with the nervous system.

Hermaphrodite (adj. hermaphroditic): Having both male and female reproductive organs.

Hind wing: Either of the posterior wings of a 4-winged insect.

Honeydew: Sugary ooze or exudate, often from , and a characteristic symptom of some fungi.

Hyaline: Like glass, transparent and colorless.

Hypha (pl. hyphae): Single, tubular filament of a fungal thallus or mycelium; the basic structural unit of a fungus.

Immune: Cannot be infected by a given pathogen.

230 Appendix C: Glossary of terms

Imperfect state: The asexual form in the life cycle of a fungus, when asexual spores (such as conidia) or no spores are produced.

Inflorescence: A characteristic arrangement of flowers on a stem; a flower cluster.

Inoculum: Pathogen or its parts, capable of causing infection when transferred to a favorable location.

Instar: An insect or other between molts.

Intercellular: Between or among cells.

Internode: A segment of a stem between two nodes.

Iridescent: A display of lustrous rainbow-like colors.

Isozymes: The different forms of an enzyme that carry out the same enzymatic reaction but require different conditions (pH, temperature, etc.) for optimum activity.

Leach: Permeate or penetrate gradually.

Leaf sheath: The part of the blade that wraps around the stem.

Lodging: To fall over.

Looper: A caterpillar that moves by looping (placing the rear end of the body next to the thorax before extending the front part of its body).

Mesocotyl: That part of the young plant that lies between the seed (which remains buried) and the plumule (extends the shoot up to the soil surface), where secondary roots develop from just beneath the plumule.

Migratory: An insect or nematode that moves from place to place on a plant, from plant to plant when feeding, or across regions (see sedentary).

Multivoltine: Pertaining to organisms with many generations in a year or season.

Necrotic: Death of cells or tissue, usually accompanied by black or brown darkening.

Nematode: Nonsegmented roundworm (animal), parasitic on plants or animals, or free living in soil or water.

Neonate: A recently born larva.

Nocturnal: Belonging to or active during the night.

231 Appendix C: Glossary of terms

Node: The small swelling that is the part of a plant stem from which one or more leaves emerge.

Obligate: Restricted to a particular set of environmental conditions, without which an organism cannot survive (e.g., an obligate parasite can survive only by parasitizing another organism).

Oblique: Having a slanting or sloping direction, course, or position; inclined.

Ocelli: A simple eye of an insect or other arthropod.

Oogonium (pl. oogonia): Female gametangium of oomycetes, containing one or more gametes.

Oospore: Thick-walled, sexually-derived resting spore of oomycetes.

Orbicular: Circular in outline.

Overseason: To survive or persist from one planting season to the next.

Overwinter: To survive or persist through the winter period.

Oviparous: Reproduction in which young hatch from eggs.

Oviposition: To deposit or lay eggs or ova; the act of depositing eggs.

Parasite (adj. parasitic): Organism that lives in intimate association with another organism on which it depends for its nutrition; not necessarily a pathogen.

Parasitoid: A parasitoid is an organism that spends a significant portion of its life history attached to or within a single host organism which it ultimately kills (and often consumes) in the process.

PCR (acronym for polymerase chain reaction): A technique used to amplify the number of copies of a specific region of DNA in order to produce enough of the DNA for use in various applications such as identification and cloning.

Pedicel: Small slender stalk; stalk bearing an individual flower, inflorescence, or spore.

Perennial: Something that occurs year after year; plant that survives for several to many years (see annual, biennial).

Perfect state: Sexual state; capable of sexual reproduction.

Pericarp: The ripened and variously modified walls of a plant ovary.

232 Appendix C: Glossary of terms

Perineal pattern: Fingerprint-like pattern formed by cuticular striae surrounding the vulva and anus of the mature nematode females.

Phasmids: Either of a pair of caudal chemoreceptors occurring in nematodes of the class Secernentea.

Phenology: The relationship between the climate and biological events, such as flowering or leafing out in plants.

Pheromone: A substance given off by one individual that causes a specific reaction by other individuals of the same species, such as sex attractants, alarm substances etc.

Phloem: Good-conducting, food-storing tissue in the vascular system of roots, stems, and leaves.

Photoperiod: The physiological reaction of organisms to the length of day or night.

Phyllody: Change of floral organs to leaflike structures.

Phylogenetic: Of or relating to the evolutionary development of organisms.

Phytophagous: Plant-eating.

Pleural membrane: A thin serous membrane in mammals that envelopes each lung and folds back to make a lining for the chest cavity.

Plumbeous: Consisting of or resembling lead.

Pneumostome: Respiratory hole.

Polyphagous: Eating many kinds of food.

Primary inoculum: Inoculum, usually from an overwintering source, that initiates disease in the field, as opposed to inoculum that spreads disease during the season (see secondary inoculum).

Prolegs: 1) Any process or appendage that serves the purpose of a leg; 2) specifically, the pliant, non-segmental abdominal legs of caterpillars and some sawfly larvae. Not true segmented appendages.

Pronotum: The upper (dorsal) plate of the prothorax.

Protonymph: Second instar of a mite.

Pubescent: Hairy.

233 Appendix C: Glossary of terms

Pygidium: Pertaining to the pygidium (i.e., the last dorsal segment of the abdomen).

Quiescent: Quiet and at rest, but not necessarily dormant and having the potential for resumed activity; can apply to non-meristematic cells.

Rachis: Floral thorn or point; axis of a compound leaf or compound inflorescence.

Random Amplified Polymorphic DNA (RAPD): A technique using single, short (usually 10-mer) synthetic oligonucleotide primers for PCR. The primer, whose sequence has been chosen at random, initiates replication at its complementary sites on the DNA, producing fragments up to about 2 kb long, which can be separated by electrophoresis and stained with ethidium bromide. A primer can exhibit polymorphism between individuals, and polymorphic fragments can be used as markers.

Real Time PCR: A laboratory technique based on polymerase chain reaction, which is used to amplify and simultaneously quantify a targeted DNA molecule. It enables both detection and quantification (as absolute number of copies or relative amount when normalized to DNA input or additional normalizing genes) of a specific sequence in a DNA sample. The procedure follows the general principle of polymerase chain reaction; its key feature is that the amplified DNA is quantified as it accumulates in the reaction in real time after each amplification cycle.

Reniform: Kidney-shaped.

Resistant (n. resistance): Possessing properties that prevent or impede disease development (see susceptible).

Restriction fragment length polymorphism (RFLP): A variation in DNA sequence that is easily recognized because it occurs at a site where a restriction enzyme cuts a specific sequence, producing DNA fragments of varying lengths. RFLP's often serve as genetic markers.

Rhizoid: Resembling roots.

Saccus: A sac.

Saturated: Impregnated; infused or filled completely; usually referring to water.

Sclerotia: A vegetative resting body of a fungus, composed of a compact mass of hyphae with or without host tissue, usually with a darkened rind.

Sclerotized: Hardened.

Secondary inoculum: Inoculum produced by infections that took place during the same growing season (see primary inoculum).

234 Appendix C: Glossary of terms

Secondary infection or secondary spread: Infection resulting from the spread of infectious material produced after a primary infection or from secondary infections without an intervening inactive period.

Sedentary: Not active; settled or remaining in one place.

Semi-looper: A caterpillar in which 1-2 pairs of the abdominal legs are absent and movement is restricted to progression only in small loops.

Sentinel plot: Unsprayed, susceptible plants that are scouted regularly as part of an early pest detection program.

Sequence Characterized Amplified Region (SCAR): SCARs are DNA fragments amplified by the Polymerase Chain Reaction (PCR) using specific 15-30 bp primers, designed from nucleotide sequences established in cloned RAPD (Random Amplified Polymorphic DNA) fragments linked to a trait of interest. By using longer PCR primers, SCARs do not face the problem of low reproducibility generally encountered with RAPDs. Obtaining a codominant marker may be an additional advantage of converting RAPDs into SCARs.

Serology: A method using the specificity of the antigen-antibody reaction for the detection and identification of antigenic substances and the organisms that carry them.

Setae: Bristles; commonly known as hairs.

Sexual dimorphism: Sexes are different in form or color in the same species; may be seasonal or geographic; male and female look different by color, form, etc.

Sign: Indication of disease from direct observation of a pathogen or its parts (see symptom).

Sinuate: Curved or curving in and out.

Skeletonize: To remove leaf tissue between the veins, leaving the network of veins intact.

Soilborne: Carried on or beneath the soil surface.

Sooty mold: Ascomycete fungi that grow from the sugary honeydew secreted by plants and insects (aphids, scales, whiteflies) that suck sap from their host plants.

Spatulate: Rounded and broad at the top, attenuate at base. Shaped like a spoon, with a narrow end at the base.

Spinose: Having spines.

235 Appendix C: Glossary of terms

Spiracles: Breathing pores; in the plural, the lateral openings on the segments of the insect body through which air enters the trachea.

Split-cane trap: Bundles of 6-8 split lengths of sugarcane, wrapped in black plastic with the ends left open.

Spore: A specialized reproductive body in fungi (and some other organisms), containing one or more cells, capable of developing into an adult.

Sporangiophore: Sporangium-bearing body of a fungus.

Sporangium (pl. sporangia): Saclike fungal structure in which the entire contents are converted into an indefinite number of asexual spores.

Sporulation: Producing spores.

Stadium (pl. stadia): The intervals between the molts of an insect.

Sterile: Unable to reproduce.

Sternites: The ventral shield or plate of each segment of the body of an insect or other arthropod.

Stippling: Series of small dots or speckles in which chlorophyll is absent.

Striated: Numerous parallel, fine, and impressed lines.

Stunting: Reduction in height of a vertical axis resulting from a progressive reduction in the length of successive internodes or a decrease in their number.

Stylet: A stiff, slender, hollow feeding organ of plant-parasitic nematodes or sap- sucking insects, such as aphids or leafhoppers.

Suffused: Cause to spread or flush or flood through, over, or across.

Superficial: Affecting, or being on or near the surface.

Susceptible: Prone to develop disease when infected by a pathogen (see resistance).

Symmetrical: Having similarity in size, shape, and relative position of corresponding parts.

Symptom: Indication of disease by reaction of the host, e.g. canker, leaf spot, wilt (see sign).

236 Appendix C: Glossary of terms

Syncytium (pl. syncytia): A multinucleate structure in root tissue formed by dissolution of common cell walls induced by secretions of certain sedentary plant-parasitic nematodes (e.g. cyst nematodes).

Synergism: Greater than additive effect of interacting factors.

Synthetic: Man-made; not of natural origin.

Tarsus: The leg segment immediately beyond the tibia, consisting of one or more segments or subdivisions.

Temperate: Free from extremes; mild.

Tergite: A dorsal sclerite or part of a segment, especially when such part consists of a single sclerite.

Tillers (subterranean): A lateral shoot, culm, or stalk arising from a crown bud; common in grasses.

Transgenic: Of, relating to, or being an organism whose genome has been altered by the transfer of a gene or genes from another species or breed.

Transient: One who stays for only a short time.

Translocate: Move from one place to another.

Translucent: Allowing light to pass through diffusely.

Transmit (n. transmission): To spread or transfer, as in spreading an infectious pathogen from plant to plant or from one plant generation to another.

Transverse: Extending or lying across; in a crosswise direction.

Trematodes: A fluke; parasitic flatworms having external suckers for attaching to a host.

Truck farming: A horticultural practice of growing one or more vegetable crops on a large scale for shipment to distant markets. At first this type of farming depended entirely on local or regional markets. As the use of railroads and large-capacity trucks expanded and refrigerated carriers were introduced, truck farms spread to the cheaper lands of the West and South, shipping seasonal crops to relatively distant markets where their cultivation is limited by climate. The major truck-farming areas are in California, Texas, Florida, along the Atlantic Coastal Plain, and in the Great Lakes area. Centers for specific crops vary with the season. Among the most important truck crops are tomatoes, lettuce, melons, beets, broccoli, celery, radishes, onions, cabbage, and strawberries.

237 Appendix C: Glossary of terms

Tubercle: Nodule; small rounded wart-like protuberance.

Tufted: Growing in small dense clumps or tufts.

Univoltine: Having only one generation per year.

Vascular bundle: A unit strand of the vascular system in stems and leaves of higher plants consisting essentially of xylem and phloem.

Vascular wilt: A xylem disease that disrupts normal uptake of water and minerals, resulting in wilting and yellowing of foliage.

Vector: Literally a bearer; specifically a host of a disease transmissible to another species of organism.

Ventral: Pertaining to the under surface of the abdomen.

Vermiform: Worm-shaped.

Vesica: Lepidoptera: the penis, or terminal part of the aedeagus. The vesica is membranous and eversible, typically held within the tubular part of the aedeagus, but everted and inflated during copulation.

Vesiculiform: Of or resembling a vesicle.

Viable: The state of being alive; able to germinate, as seeds, fungus spores, sclerotia, etc.; capable of growth.

Virulence: Degree or measure of pathogenicity; relative capacity to cause disease.

Virulent: Highly pathogenic; having the capacity to cause severe disease.

White heads: Empty panicles or with a few filled grains.

Whorl: Spiral pattern.

Wilt: Drooping of leaves and stems from lack of water (inadequate water supply or excessive transpiration); vascular disease that interrupts normal water uptake.

Xylem: Water and mineral conducting, food-storing, supporting tissue of a plant.

Zoospores: Fungal spore with flagella, capable of locomotion in water.

Definitions taken from:

Agrios, G.N. Plant Pathology. Fourth Edition. Academic Press, San Diego. 1997.

238 Appendix C: Glossary of terms

American Phytopathological Society. 2005. Illustrated Glossary of Plant Pathology. http://www.apsnet.org/education/IllustratedGlossary/default.htmH .

Borror, C.J., DeLong, D.M., and Triplehorn, C.A. 1976. An Introduction to the Study of Insects. Fourth edition. Holt, Rinehart and Winston, New York.

De La Torre-Bueno, J.R. 1985. A Glossary of Entomology. New York Entomological Soc., New York.

Hansen, R. 2007. Personal Communication.

Newfeldt, V. ed. 1988. Webster’s New World Dictionary Third College Edition. Cleveland and New York.

239

Appendix D: FY09 CAPS Prioritized Pest List and Commodity Matrix

240

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Appendix E: FY10 CAPS Prioritized Pest List and Commodity Matrix

243

244

245

246

Appendix F: 2008 Exotic Pest Detection Survey Order Form

247

248

Appendix G: Plastic Bucket Trap Protocol

Plastic Bucket Trap Protocol

The plastic bucket trap is a long-lasting insect trap used in conjunction with a lure to monitor or detect various species of moths. The plastic bucket trap is the preferred trap for some moth species as it is able to catch large numbers of moths without damaging some of their identifying characters. The trap has four parts: 1) lid, 2) lure basket with cap, 3) funnel, and 4) bucket. The trap is available in various color combinations. For PPQ programs, the trap consists of a green lid, yellow funnel, and white bucket. Fig. 1 is a photograph of a trap cut in half.

1 Lid

Lure basket

Funnel

Bucket

Fig. 1. Plastic bucket trap cut in half to show its interior.

249

Follow the steps below to prepare the bucket traps for use in the field.

1. Pheromone Unwrap a pheromone lure and place it inside the lure basket. Handle lures with gloves (see Fig. 4). Close the basket with a cap and insert the basket through the circular opening on the center of the lid (Fig. 2). If the cap no longer snaps snuggly into the trap lid opening, secure it with a piece of tape.

2

Cap Lure basket

Fig. 2. Lure basket with cap inserted through center of lid.

The synthetic pheromone is embedded in a small rubberized square (as seen in the photos below) or septum (similar to a pencil eraser). If the lure is flat and small (Figs. 3 and 4) you may attach the lure to a small paper clip and fold the clip so that the lure does not fall out of the basket. If a lure basket is not available, attach the lure to a cork with a pin and place the cork in the lid’s opening. Always carry extra corks.

3 4

Figs. 3 and 4. Lure made of a small rubberized square with embedded synthetic pheromone chemicals.

When not in use, the lures should be stored, unwrapped, in a freezer not used for food or drinks. MSDS documents for the pheromones to be used should be available and should be read.

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2. Handle Attach a wire handle to the lid through its two loops, as shown in the photos below (Figs. 5 and 6). A wire handle is usually included with each purchased trap. If a handle is not included, is lost, or is damaged and needs to be replaced, make one with a 12-inch long wire or with string, but the latter does not last as long as the wire.

5 6

Figs. 5 and 6. Wire handle attached to trap’s lid.

3a. Sponge Place a dry cellulose sponge in the bottom of the trap, as shown in Fig. 7. The sponge will absorb rainwater (except for extremely heavy amounts) that may enter the trap, keeping the moths somewhat dry.

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Fig. 7. Cellulose sponge inside the trap.

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3b. Wire screen Alternatively, the bottom part of the trap, the bucket, requires two modifications. Drill two to four drain holes in the bottom (see Fig. 8). If water remains in the trap, the killing agent (the pesticide) can spoil; in addition, the trapped insects may decay, making identification impossible.

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Fig. 8. Bucket with four drilled holes.

Then, add a wire screen slightly larger than the bucket bottom’s inside diameter (Figs. 9 and 10). The screen keeps the pesticide strip(s) and the moths from getting too wet from rainwater accumulated in the trap. Prepare a cardboard template for long term use. Cut the wire mesh with metal-cutting scissors.

9 10

Figs. 9 and 10. Metal wire screen inside the bucket.

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4. Insecticidal strips Place two insecticidal strips (Figs. 11 and 12), which kill the moths that enter the bucket. The active ingredient in the strips is Dichlorvos, also known as DDVP and Vapona. The strips should be handled with gloves. Read and have available the MSDS documents for this product. Store unopened strips in a freezer not used for food or drink. Rain, wind, high heat or an abundance of captured moths may reduce its potency from 3 to 4 weeks to a week or less. If using only one kill strip, change it every 2 weeks.

12

11

Figs. 11 and 12. Pesticide strips.

5. Label the trap Attach a rain-proof printed label (see Fig. 13) or handwrite a note with a water- proof black marker on the bucket trap. It should indicate that the trap belongs to a state or a PPQ program. Include a phone number in case someone has concerns or questions about the trap.

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Fig. 13. Label on the trap’s lid.

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6. Placement of traps The traps function best when placed in the open, away from foliage, as illustrated on Fig. 14. When hung under foliage, the 3-dimensional shape of the pheromone plume (chemical in the air) is disrupted and the effectiveness of the trap is much reduced. Hang the traps from such places as greenhouse roofs or in the open using metal rods (see Fig. 14) or other materials.

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Figs. 14. Trap set away from foliage, in open field.

In the field, transfer the caught moths to labeled zip-loc bags and store them in a cooler (Figs. 15 and 16). Place them overnight in a freezer to kill any surviving specimens. 16 15

Figs. 15 and 16. Moths placed in a ziploc bag and stored in a cooler.

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Prior to shipping, screen the samples. Remove any moth vastly different from the target and all other arthropods (beetles, flies, spiders). Write on PPQ Form 391 the approximate number of moths being submitted. Place an absorbent paper, such as a piece of a paper towel, inside each plastic bag to reduce moisture and to pad the specimens for their protection. The specimens should be well padded inside a box to prevent the specimens from being crushed or otherwise damaged. If longer-term storing is necessary, freezing works best, but refrigeration is acceptable as well.

The general recommendation for maintenance of the plastic bucket traps is to wash them occasionally with soap and water to keep them clean, and to store them indoors, or at least protected from sun, rain and dust. Keep the wire handle and the wire screen in good repair. The traps can be used multiple times and for multiple species since the chemicals degrade quickly in outdoor conditions. These traps usually last more than 5 years.

This protocol is designed to aid in the detection of exotic moths of concern by giving instructions on how to use generic plastic bucket traps. All photos were taken by J. Brambila and R. Meagher. These instructions are primarily based on work by R. Meagher. This aid was prepared by Julieta Brambila (USDA/APHIS/PPQ Eastern Region), Lisa Jackson (USDA/APHIS/PPQ/CPHST), and Dr. Robert L. Meagher (USDA/ARS/CMAVE), on April 2010.

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