2014

Table of Contents

Table of Contents

Authors, Reviewers, Draft Log ...... 4 Introduction to Reference ...... 6

Arthropods ...... 14 Primary Pests of Grape (Full Pest Datasheet) ...... 14 Autographa gamma ...... 14 Cryptoblabes gnidiella ...... 23 Diabrotica speciosa ...... 36 Epiphyas postvittana ...... 45 Eupoecilia ambiguella ...... 59 Heteronychus arator ...... 67 Lobesia botrana ...... 75 Scirtothrips dorsalis ...... 89 Spodoptera littoralis ...... 103 Spodoptera litura ...... 113 Secondary Pests of Grape (Truncated Pest Datasheet) ...... 124 Adoxophyes orana ...... 124 Aleurocanthus spiniferus ...... 128 Eudocima fullonia ...... 133 Eutetranychus orientalis ...... 138 Oxycarenus hyalinipennis ...... 143 Planococcus lilacinus ...... 149 Thaumatotibia leucotreta ...... 153 Pests of Grape Requested by the CAPS Community...... 161 Homalodisca vitripennis ...... 161

Mollusks ...... 171 Primary Pests of Grape (Full Pest Datasheet) ...... 171 None at this time ...... 171 Secondary Pests of Grape (Truncated Pest Datasheet) ...... 171 Cernuella virgata ...... 171 Lissachatina fulica ...... 174

Nematodes ...... 179 Primary Pests of Grape (Full Pest Datasheet) ...... 179 Xiphinema italiae ...... 179 Secondary Pests of Grape (Truncated Pest Datasheet) ...... 189 Meloidogyne mali ...... 189

Plant Pathogens ...... 193 Primary Pests of Grape (Full Pest Datasheet) ...... 193 ‘Candidatus Phytoplasma australiense’ ...... 193 ‘Candidatus Phytoplasma vitis’...... 206 Phellinus noxius ...... 221

2 Table of Contents

Pseudopezicula tracheiphila ...... 230

Appendix A: Diagnostic Resource Contacts ...... 237

Appendix B: Glossary of Terms ...... 239

3 Authors, Reviewers, Draft Log

Authors, Reviewers, Draft Log

Authors CAPS Commenters Melinda Sullivan Darryl Jewett – Pest Survey Specialist - NY Plant Pathologist National Weed Management Laboratory Gary Russell – State Operations Support 2301 Research Blvd., Suite 108 Officer – USDA-APHIS-PPQ – Phoenix, Fort Collins, CO 80526 Arizona

Nehalem Breiter Erin N Stiers- Pest Survey Specialist - KS, Biological Science Technician OK, CO National Weed Management Laboratory 2301 Research Blvd., Suite 108 Craig Webb- Domestic Identifier –USDA- Fort Collins, CO 80526 APHIS-PPQ – Manhattan, Kansas

Talitha Price Andrea Simao Biological Science Technician National Program Manager - LBAM USDA-APHIS-PPQ-CPHST USDA APHIS PPQ 1730 Varsity Dr., Suite 400 Riverdale, MD Raleigh, NC 27606 Eileen Smith CPHST Reviewers National Program Manager - EGVM Thomas Culliney USDA APHIS PPQ Entomologist Riverdale, MD Pest Epidemiology and Risk Analysis Laboratory Draft Log 1730 Varsity Drive July 2007 - Draft sent for CPHST review Raleigh, NC 27606

September, 2007 – Draft sent for CAPS Lisa Jackson comment. Biological Science Technician

CPHST Director’s Office December, 2007 – Draft sent to NAPIS for 1730 Varsity Drive posting Raleigh, NC 27606

February, 2008 – Revised draft posted on Kimberly Schwartzburg NAPIS Risk Analyst

Plant Epidemiology and Risk Analysis May, 2008 – Grape Commodity map Laboratory updated. 1730 Varsity Drive

Raleigh, NC 27606 September, 2009 – Updated risk maps added to document.

June, 2010 - Datasheets for Diabrotica speciosa and Eupoecilia ambiguella added, CAPS-Approved survey and diagnostic method information added, updated risk maps added, and tertiary pests removed. All tertiary pests were listed with name and photo only. A full datasheet will be added for each ‘tertiary’ pest as they become available and undergo the pest list prioritization process.

4 Authors, Reviewers, Draft Log

Draft Log Continued: July, 2010 – Added datasheet for Xiphinema italiae (a tertiary pest).

August, 2010 – Updated hyperlinks for links now on the CAPS Resource and Collaboration site

April, 2012 – Trap and lure information updated to reflect names used in the IPHIS Survey Supply Ordering System. Lobesia botrana lure effectiveness changed from three to four weeks. New maps added for pests. Xylella fastidiosa (citrus variegated chlorosis strain) datasheet was removed per suggestion of the CAPS Pest List Working Group. Minor changes made for a few pest datasheets (host/distribution information primarily). Information about Tort AI- Tortricids of Agricultural Importance, a new diagnostic tool developed by CPHST’s Identification Technology Program, was added where appropriate.

June 2012 – Removed Copitarsia spp. and Planococcus minor datasheets at the suggestion of the CAPS Pest List Working group for FY 2013 surveys. Planococcus minor has been deregulated. Both pests are not available for the 2013 survey season.

June 2013: Updated ‘Candidatus Phytoplasma australiense’ datasheet.

August 2013: Added datsheets for three new 2014 AHP CAPS targets: Candidatus Phytoplasma vitis, Cryptoblabes gnidiella, and Pseudopezicula tracheiphila.

August 2016: Removed outdated maps.

5 Introduction to the 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 C and D) and the Select Agent list (http://www.selectagents.gov/ or http://www.aphis.usda.gov/programs/ag_selectagent/ag_bioterr_toxinslist.html). The AHP prioritized pest list for FY 10/11’ and 12’ are also given in Appendices C and D. 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 United States, or if specifically requested

6 Introduction to the Reference

by the CAPS National Committee. States are not required to survey for all of the pests 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. 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, corn, cotton, small grains, stone fruit, oak forests, and pine forests.

Grape Commodity Survey Reference The Grape Commodity Survey Reference (CSR) is a companion document to the Grape Commodity Survey Guidelines (CSG). Both documents are intended to be tools to help survey professionals develop surveys for exotic grape pests. The Grape CSR is a collection of detailed data sheets on exotic pests of grape. Additionally, the authors have tried to identify native pests that these exotic pests may be easily confused with 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.

In contrast, the Grape Commodity Survey Guidelines 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 guidelines contain little information about biology. Instead, they focus on survey design, sampling strategies, and methods of identification. There is no 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 guidelines 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.

7 Introduction to the Reference

Purposes of the Grape 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 generally (but not always) 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

8 Introduction to the Reference available. PPQ offers diagnostic support for the CAPS program through NIS. A major 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 PPQ 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 A has a listing of NIS and Domestic Identifier contact information.

Organisms Included in the Grape Survey Reference Organisms included in the grape survey reference are organized first by:

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

2. Organisms are then divided by their pest status on grape [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

9 Introduction to the Reference

requested by the CAPS Community: Full pest datasheets will be developed for specific pests requested by the CAPS community.

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 2012 CAPS National Survey Guidelines (http://caps.ceris.purdue.edu/webfm_send/1063; http://caps.ceris.purdue.edu/node/223) 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.

10 Grape Background

Introduction to Grape Cultivated grapes, consumed as fresh fruit, raisins, wine, and juice, are an important crop in the United States. The many varieties and cultivars of grape belong to the Vitaceae family. The Vitis genus grows in eastern Asia, Europe, the Middle East and North America between 25° and 50° N latitude. Grapes can be classified by either food usage or by species (Reiger, 2006).

Grape production in the United States The United States ranks fourth worldwide in grape production, contributing 8% of the total pounds produced globally. The U.S. grape industry is valued at approximately $3.1 billion dollars. In 2006, the United States produced a total of 6.33 million tons of grapes, valued at $499 per ton. Grapes are the highest value non-citrus fruit/nut crop in terms of both utilized production and value. Grape-producing states include Arizona, Arkansas, California, Georgia, Michigan, Missouri, New York, North Carolina, Ohio, Oregon, Pennsylvania, Texas, Virginia, and Washington. The majority of U.S. grapes, 90%, are grown in California with 800,000 acres in production (NASS, 2007).

Species and Cultivars The genus Vitis is divided into two subgenera: Euvitis and Muscadinia. Euvitis, or ‘true grapes,’ are characterized by fruits growing in elongated clusters and attached to stems at maturity. Euvitis grapes have forked tendrils, diaphragms in pith at nodes, and long strips of loose bark detached from the vine. Muscadinia, or Muscadine grapes, have small clusters of thick-skinned fruit which detach as they mature. Tendrils of Muscadine grapes are simple,

11 Grape Background

diaphragms in pith at nodes are lacking, and vines have smooth bark with lenticels (Reiger, 2006).

Grape production worldwide primarily consists of only four species or hybrids. Vitis vinifera L., often named the ‘Old World grape,’ is prized for use in wine production and for table and raisin grapes. Worldwide production is dominated by V. vinifera, with 90% of grape production in this species with at least 500 V. vinifera cultivars grown. Vitis rotundifolia Michx is a muscadine grape that is disease tolerant and grows vigorously. This species is not cold hardy but is well adapted to the southeastern United States. Vitis labrusca L. includes the popular ‘Concord’ cultivar, among others, and is grown for grape juice, jams, and associated products. Lastly, French-American hybrids between vinifera grape and native species rootstocks are grown to provide increased Phylloxera resistance (Reiger, 2006).

Food Usage Classification of grapes by food usage includes table grapes, raisin grapes, sweet juice grapes, and wine grapes. Table grapes are those grown for and consumed as fresh fruit. V. vinifera ‘Thompson seedless’ grapes are widely grown as table grapes, as well as many other cultivars. ‘Thompson seedless’ is the most common raisin grape cultivar, making up 90% of raisin production in the United States. Sweet juice grapes grown for juice, jelly, jam, preserves and certain types of wine are typically ‘Concord’ grapes. Commercial wine grape production is dominated by cultivars of V. vinifera (Reiger, 2006).

Domestication It is thought that grapes, Vitis vinifera, are native to southwestern Asia near the Caspian Sea. Like other fruits native to that region, such as pears and apples, grapes most likely spread to new places as a result of trade. Grape seeds dating back to the Bronze Age have been found in excavated dwellings in south-central Europe. Egyptian hieroglyphics told of wine making. Grapes were probably brought to Greece, Rome and France by the Phoenicians. Early settlers to North America brought grape starts, but the grapes faired poorly on the east coast. Spanish missionaries introduced vinifera-type grapes to California in the 1700s, where they have flourished as a crop (Reiger, 2006).

Biology and Reproduction of Grapes All grapes are woody, climbing vines. Muscadine grapes have smooth bark with lenticels and small, round, unlobed leaves with dentate margins. Vinifera grapes are characterized by older vines with loose, flakey bark and large leaves. Vinifera leaves can vary greatly in both size and shape and may or may not be lobed. Small, green flowers appear on racemose panicles at the base of the current season’s growth. Flowers have five sepals, five petals and five stamens. Superior ovaries have two locules, each with two ovules. Vinifera and Concord grapes have perfect flowered, self- pollinating flowers, and some muscadine cultivars are only found with pistillate flowers. The majority of grapes are self-pollinating, with some exceptions in muscadine grapes. Fruits of grapes are true berries, round to oblong in shape, and have up to four seeds. Skin is variable in thickness, but most is thin. A fine, glaucous layer of wax may be

12 Grape Background present on the fruit surface. Anthocyanin compounds are found in the skin, which colors the fruit red, blue, purple, or black (Reiger, 2006). References NASS (National Agricultural Statistics Service). 2007. Noncitrus fruits and nuts: 2006 preliminary summary – Fr Nt 1-3 (07). Agricultural Statistics Board, USDA-NASS.

Reiger, M. 2006. Grapes – Vitis spp. (Introduction to Fruit Crops), University of Georgia. http://www.uga.edu/fruit/ Accessed on 2/26/2007.

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

Arthropods

Primary Pests of Grape (Full Pest Datasheet)

Autographa gamma

Scientific Name Autographa gamma L.

Synonyms: Phytometra gamma and Plusia gamma

Common Name(s) Silver-Y moth, beet worm

Type of Pest Moth A

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

Reason for Inclusion in Manual CAPS Target: AHP Prioritized List 2006 through 2009

Pest Description B Eggs: Semi-spherical, 0.57 mm (0.022 in.) in diameter. Eggs are yellowish-white Figure 1. Eggs (A) and larvae (B) of A. (Fig. 1A), later turning yellowish-orange to gamma (A.) and . Photos courtesy of brown. The number of ribs varies from 28 Jurgen Rodeland and P. Mazzei (www. to 29 (Paulian et al., 1975). The eggs are invasive.org), respectively. 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 (0.79 to 1.57 in.) (Emmett, 1980; Hill, 1983; Jones and Jones, 1984; USDA, 1986).

14 Autographa gamma Primary Pest of Grape 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 pupa is brown to black, greenish or even whitish-green on its ventral side, 16 to 21 mm long (0.63 to 0.83 in.), and 4.5 to 6.0 mm (0.18 to 0.24 in.) broad. Cremaster globular, with four pairs of hooklets (Paulian et al., 1975; Carter and Hargreaves, 1986).

Adult: The adults are gray-colored and the forewings are marbled in appearance; their color being A silvery-gray to reddish-gray to black with a velvety sheen. Wing expanse is 36 to 40 mm (1.42 to 1.57 in.). 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 able to breed. The silver-Y moth B 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 moths 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 batches, on the underside of leaves Figure 2. Cocoon (A) and adult (B) A. gamma. of low-growing plants. In temperate Photos courtesy of Alain Fraval and Jeremy Lee, regions, hatching may take 10 to respectively. 12 days (Hill, 1983). The incubation period lasts for three 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 they 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 Autographa gamma Primary Pest of Grape Arthropods Silver-Y moth Moth

15 to 16 days at 25°C (77°F), 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, 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 egg 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 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. Leaves may be skeletonized by larval feeding. Older leaves are preferred by larvae. The petioles or leaf stalks may be cut by the larvae. Frass may or may not be visible. Apart from damaging the foliage of their host plants, larvae can scrape the skin from grapes and feed on the contents of the fruits. A single larva can damage 20 or more mature grapes (Abdullagatov and Abdullagatov, 1986). Pupae are found in the folds of the lower leaves of the host plant. Webbing may be present. Adult moths feed on flowers 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, Asia and North Africa. The outbreak of 1928, which occurred in most of central Europe, caused widespread defoliation of peas in Poland. Damage from this insect and Pieris rapae (the small white butterfly) in areas of the Netherlands was valued at as much as 320,000 guilders 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. (mustards) and other vegetables, including legumes. Grape is considered a primary

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

host. 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. vulgaris (sugar beet), 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), Chrysanthemum indicum (chrysanthemum), Cicer arietinum (chickpea), Cichorium intybus (chicory), Cynara cardunculus (artichoke), Daucus carota (carrot), Glycine max (soybean), Gossypium spp. (cotton), Helianthus annuus (sunflower), Hyssopus officinalis (hyssop), Lactuca sativa (lettuce), Linum usitatissimum (flax), Medicago sativa (alfalfa), Nicotiana tabacum (tobacco), Pelargonium (geranium) hybrids, Petroselinum crispum (parsley), Solanum tuberosum (potato), Spinacia oleracea (spinach), Trifolium pratense (purple clover), 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 India and China; it also occurs in North Africa (USDA, 1958).

Asia: Azerbaijan, China, India, Iran, Iraq, Israel, Japan, Kazakhstan, Korea, Saudi Arabia, Syria, Turkey, and Uzbekistan. Europe: Albania, Andorra, Austria, Belarus, Belgium, Bosnia and Herzegovina, Bulgaria, Channel Islands, Croatia, Cyprus, Czech Republic, Denmark (including Faroe Islands), Estonia, Finland, , France (including Corsica), Germany, Greece (including Crete, Cyclades, Dodecanese), Hungary, Iceland, Ireland, Italy (Including Sardinia and Sicily), Latvia, Liechtenstein, Lithuania, Luxembourg, Macedonia, Malta, Moldova, Montenegro, the Netherlands, North Aegean Islands, Norway, Poland (including Azores and Madeira), Portugal, Romania, Russian Federation (including Kaliningrad Oblast), Serbia, Slovakia, Slovenia, Spain (including Balearic Islands, Canary Islands), Sweden, Switzerland, Ukraine, and United Kingdom (including Gibraltar). 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). A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates that many states in the United States have a low to moderate risk rating for A. gamma establishment based on host availability and climate within the continental United States. Establishment is precluded in most of the northern states. Areas of

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

Arkansas, Kentucky, Louisiana, Mississippi, North Carolina, Oklahoma, and Virginia have the highest risk of A. gamma establishment.

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination. The trap is Plastic Bucket Trap. The lure is effective for 28 days (4 weeks).

The Lure Product Name is “Autographa gamma Lure”.

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

Method Notes: This trap is also known as the unitrap. The trap has a green canopy, yellow funnel, and white bucket and is used with a dry kill strip. For instructions on using the trap, see Brambila et al. (2010).

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

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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 pheromone, (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

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

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of A. gamma is requires morphological identification by a trained entomologist/taxonomist.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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 (Fig. 4), A. pseudogamma (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. (1981) 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.

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

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

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.

20 Autographa gamma Primary Pest of Grape Arthropods Silver-Y moth Moth

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

References Abdullagatov, A.Z. and Abdullagatov, K.A. 1986. The gamma moth as a vineyard pest. Zashchita Rastenii, No. 11:40 (English Abstract).

Brambila, J., Jackson, L., and Meagher, R. 2010. Plastic bucket trap protocol. http://caps.ceris.purdue.edu/webfm_send/398.

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

Carter, D.J. and Hargreaves, B. (Illustrator). 1986. A field guide to caterpillars of butterflies 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.

21 Autographa gamma Primary Pest of Grape Arthropods Silver-Y moth Moth

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.

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. 1980-81. Redescription and classification of subfamily Plusiinae 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.

22 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

Cryptoblabes gnidiella

Scientific Name Cryptoblabes gnidiella Millière

Synonyms: Albinia casazzar Briosi Albinia gnidiella Millière Albinia wockiana Cryptoblabes aliena Swezey Cryptoblabes wockeana Ephestia gnidiella Millière

Common Name(s) Honeydew moth, Christmas berry webworm, citrus pyralid, earhead caterpillar, false blossom moth, and lemon borer moth

Type of Pest Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Figures 1 & 2. Cryptoblabes gnidiella adults Family: Pyralidae (http://pathpiva.wifeo.com/).

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2014

Pest Description Eggs: “Oval, irregularly reticulate; at first white, becoming yellow before hatching” (CABI, 2012).

Larvae: “Measures 12 mm [approx. ½ in] long when fully grown, narrowing towards each end. Head and thoracic plate reddish-brown to black; body yellowish, olivaceous, reddish- or brownish-grey with longitudinal stripes; peritreme of spiracles reddish-brown; setae yellowish-brown; pinacula blackish-brown surrounded by pale Figure 3. Cryptoblabes gnidiella larva ochreous coloration. Thoracic legs (http://pathpiva.wifeo.com/).

23 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth reddish-brown to black (Fig. 2)” (CABI, 2012).

Pupae: “Measures 5 – 6 mm long. At first greenish, becoming reddish-brown (Fig. 4). Fine hairs in two dorsal, two lateral, and two ventral rows. Cremaster with two slender prongs terminating in strongly angular hooks. Formed in a slight cocoon” (CABI, 2012).

Adults: “Wingspan 11 – 20 mm. Head and thorax greyish-brown. Antennae simple, finely ciliate. Forewing greyish- Figure 4. Cryptoblabes gnidiella pupa brown with a variable amount of whitish (http://pathpiva.wifeo.com/). suffusion, scattered reddish-brown scales give a purplish appearance (Fig. 1 & 2); postmedian and antemedian fascias pale, indistinct. Hindwing shining white; veins conspicuous, pale greyish-brown. Abdomen shining greyish-white” (CABI, 2012).

Descriptions of the life stages can also be found in Avidov and Harpaz (1969) and Carter (1984). Descriptions of the immature stages can be found in Heckford and Sterling (2004). Description of the adult can be found in Goater (1986).

Biology and Ecology Adults are active at night (Avidov and Gothilf, 1960) and mate on the same night of emergence; females begin oviposition the following day (Avidov and Harpaz, 1969). Most females will mate once, while males have been recorded mating multiple times (Wysoki et al., 1993). Adults are attracted to sweet material, including honeydew excreted by mealybugs and also fruits like grapes and pomegranates that have been injured by other insects (Avidov and Harpaz, 1969). Adults live for about a month during the cold season and 8 to 10 days during the warm season (Avidov and Gothilf, 1960).

Females lay their eggs on fruit, usually singly or in small batches (Avidov and Harpaz, 1969). In mangos, adult females will lay eggs on the tender leaves near the main rib, on the fruit, or on the tender branches (MALR, n.d.). In garlic, eggs are laid on the inner surface of the bulb sheath or in the garlic lobe depressions (Salama, 2008). Females live from one to four weeks and lay an average of 150 eggs during this time (Avidov and Harpaz, 1969).

Eggs hatch after about four days in Brazil (Botton et al., 2003). In Egypt, eggs hatch in 8 to 13 days (MALR, n.d.). Once larvae hatch, they will initially feed solely on the honeydew produced by mealybugs and insect remnants. As the larvae mature, they begin nibbling superficially on the skin of citrus fruit (Avidov and Harpaz, 1969). Bagnoli and Lucchi (2001) state that larval diet includes sweet matter, dry flower parts, berry juice, berry stalk, and even healthy grapes. Larval populations can be very high,

24 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

especially if they are close to a large colony of mealybugs. If no mealybugs are present, larvae can develop on grape berries that are almost ripe. Larvae develop slightly faster on grapes versus citrus fruit with mealybugs (Avidov and Harpaz, 1969). Larvae can be found in sheltered places among the fruit or between fruits and leaves. They are enveloped by webs and waste matter (Avidov and Gothilf, 1960). Larvae go through five larval instars.

The larvae pass through a prepupal stage in which they stop feeding and become sluggish. Larvae then secrete a thin silk cocoon before molting into a pupa (Salama, 2008). Pupation occurs close to the larval feeding site (Avidov and Harpaz, 1969), either on the host plant or on the ground. Overwintering can occur in either the larval or pupal stage (Carter, 1984).

Complete development can range from five weeks during the summer, to five months during the winter. In Israel, five to six generations occur per year on citrus. If the host cycle also includes grapes, then six to seven generations may occur per year (Avidov and Harpaz, 1969). Carter (1984) states that this species has three to four generations a year in southern Europe and up to five in North Africa. The number of generations per year is highly dependent on the climate and host plants utilized. The generations can move between different host plants depending on what is available at the time.

In Mediterranean climates, this species has three to four periods of adult flight activity: May to June, July, and August to October (Bagnoli and Lucchi, 2001). Avidov and Gothilf (1960) report the temperature threshold for this species is 13°C (55.4°F). One generation requires 455 degree days (Avidov and Gothilf, 1960). MALR (n.d.) states that the optimum temperature for this species is 25 to 30°C (77 to 86°F) with 65 to 67% relative humidity.

Symptoms/Signs Larvae of this species may be found as early as August in Israel; however, damage to fruit usually occurs only from late October onwards (Avidov and Harpaz, 1969). Symptoms include presence of eggs or larvae on the fruits, silk webbing produced by the larvae, and molds which can develop on the attacked fruits (MALR, n.d.). This species is attracted to honeydew of coccids, especially Pseudococcus citriculus (Zhang, 1994).

In grape berries, young larvae enter the fruit at the junction between the fruit and stalk (Avidov and Harpaz, 1969). Botton et al. (2003) state that the larvae feed on the stems when the grapes are green, causing wilting and grape fall. When attacked close to harvesting, juice leakage can lead to rot by secondary pathogens which can reduce wine quality (Botton et al., 2003). Larvae produce dense webbing spun around the host stalk (Carter, 1984).

In lemons, eggs are laid on young fruit. When hatching, larvae penetrate the underside of the egg into the fruit. Larvae do not usually go further than the flavedo (the outer part

25 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

of the rind). This can cause the fruit to gum, resulting in necrotic blotches on mature fruit (Moore, 2003).

In sweet orange, feeding by the larvae leads to fruit piercing and gallery formation on the fruit exocarp and endocarp. This can result in gum-exudation, yellowing of the fruit, and premature fruit drop (Silva and Mexia, 1999).

This species has been recorded destroying large quantities of loquat flowers (reviewed in Avidov and Harpaz, 1969). In avocado, severe injury can be caused by C. gnidiella’s superficial feeding on the skin of the avocado fruit. This can occur even in the absence of honeydew producing Homoptera (Ben Yehuda et al., 1991).

This species is usually associated with other pests of host plants like scale insects or mealybugs and their honeydew (Ben Yehuda et al., 1991). In Israel, this species coexists with Lobesia botrana in grapevines (Harari et al., 2007). In avocado, this species can be found with Pseudococcus longispinus (long-tailed mealybug).

Pest Importance This species is a pest of avocado, citrus, grapes, loquat, and pomegranates in Israel (Ascher et al., 1983). Losses associated with this species are not well quantified in the literature (CABI, 2012). This is likely due to the fact that this species is usually associated with a variety of other pests in host crops, including birds, bacteria and fungi, and insects (including aphids, mealybugs, and Lobesia botrana) (reviewed in Silva and Mexia, 1999). Many sources state that this species is not a serious pest; however, there are records of this species causing damage on certain hosts.

In the field, C. gnidiella mostly appears on grape bunches that have previously been damaged by grape berry moth. However, experiments have shown that this species can develop on healthy grapes as well (Avidov and Harpaz, 1969). Bagnoli and Lucchi (2001) state that this species is a frequent and harmful pest on grapes in Tuscany, Italy as this species does well in areas with Mediterranean conditions. They state that this species is only faintly destructive (Bagnoli and Lucchi, 2001). Bisotto-de-Oliveira et al. (2007) state that this species is an important pest of grapevine orchards in Rio Grande do Sul, Brazil. In addition, Sellanes et al. (2010) state that this species is an economically important pest of vineyards in Brazil and Uruguay.

Avidov and Harpaz (1969) state that larval infestation of citrus is not considered serious; while Gentry (1965) states that this species is regarded as a secondary feeder causing no serious primary damage. This species can cause occasional fruit drop as a result of infestation (Gentry, 1965). Damage on grapevine in Israel is considered secondary (Avidov and Harpaz, 1969). Larvae have been recorded on pomegranates which were previously infested by the pomegranate butterfly (Avidov and Harpaz, 1969).

Akanbi (1973) states that this species is found in association with trees in the Meliaceae family in Nigeria, while Chandel et al. (2010) state that this species is a serious pest of hybrid sorghum and bajra crops. In Egypt, this species is considered one of the most

26 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

important insect pests of stored garlic (El-Zemaity et al., 2009); it is also prevalent in damaged corn ears in Egypt (Gentry, 1965). Hashem et al. (1997) state that this species is a serious pest of fruit orchards and vegetable and field crops. EPPO (2004) states that this species can cause severe damage to oranges (cultivars Navel and Navelina) by causing premature fruit fall. Fruits remaining on the tree change color on the lowest part of the tree but remain green elsewhere on the tree (EPPO, 2004). Ben Yehuda et al. (1991) state that this species can also infest avocado as a primary pest. In South Africa, this species has been recorded causing up to 5% crop reduction in lemon on the Eastern Cape and up to 50% fruit damage on the Western Cape (Moore, 2003). Özturk and Ulusoy (2009) state that this species is a pest of pomegranate in Turkey. This species has been recorded causing damage to Macadamia ternifolia in Israel (Wysoki, 1986). Larvae have also been found on cotton bolls after a primary infestation by cotton pests (Avidov and Harpaz, 1969). Outside of Israel, this species has been recorded on previously injured fig and peach. Damage to other hosts, including wheat, corn, millet, and castor bean are usually light (Avidov and Harpaz, 1969).

This species is usually associated with other pests of host plants like scale insects or mealybugs and their honeydew (Ben Yehuda et al., 1991). In Israel, this species coexists with Lobesia botrana in grapevines. Harari et al. (2007) state that damage caused by C. gnidiella and L. botrana in Israel is twofold, direct damage “caused to clusters when the larvae feed within (L. botrana) and among (C. gnidiella) the berries” and indirect damage “characterized by fungal infestation of injured berries.”

Avidov and Harpaz (1969) state that control of this pest should focus on controlling the primary pest which allows for Cryptoblabes gnidiella infestation. Many countries use chemical control to lessen the effects caused by the primary pest and C. gnidiella (Harari et al., 2007). Other control methods include biological control (like Bacillus thuringiensis applications) and mating disruption (Wysoki et al., 1975; 1981; 1988a; b; Ben Yehuda et al., 1991; 1993; Sellanes et al., 2010).

Known Hosts In Israel, this species is only recorded on seven plant species: citrus, corn, cotton, grapevine, loquat, pomegranate, and sorghum (Avidov and Harpaz, 1969). In Brazil, this species has also been found on avocado, banana, and coffee (Botton et al., 2003). Carter (1984) also lists plum, peach, and apple as hosts.

Major hosts Citrus spp. (including orange, grapefruit, lemon*), Persea americana (avocado), Punica spp., Punica granatum (pomegranate), and Vitis spp. (grape) (reviewed in CABI, 2012).

Other hosts Acacia karroo (karroothorn), Acacia nilotica (gum arabic tree), Actinidia deliciosa (kiwi), Allium spp., Allium cepa (garden onion), Allium sativum (garlic), Annona muricata (soursop), Azolla spp. (mosquitofern), Azolla anabaena (water fern), Azolla pinnata (feathered mosquitofern), Ceratoniae siliqua (fig), Chaenomeles japonica (Maule’s

27 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth quince), Coffea spp. (coffee), Coffea arabica (Arabian coffee), Cyanea procera (Molokai cyanea), Cydonia oblonga (quince), Cyperus rotundus (nutgrass), Daphne gnidium, Daphne mezereum (paradise plant), Daucus carota (Queen Anne’s lace), Eleusine coracana (finger millet), Eriobotrya japonica (loquat), Feijoa sellowiana (feijoa), Ficus spp. (fig), Ficus carica (edible fig), Ficus macrophylla (Moreton Bay Fig), Gossypium spp. (cotton), Gossypium arboreum (tree cotton), Gossypium thurberi (Thurber’s cotton), Khaya senegalensis, Lantana spp. (lantana), Lythrum spp. (loosestrife), Lythrum salicaria (purple loosestrife), Macadamia spp. (macadamia), Malus spp. (apple), Malus pumila (paradise apple), Mangifera indica (mango), Mespilus spp. (medlar), Mespilus germanica (medlar), Morus alba (mora), Morus nigra (black mulberry), Musa spp. (banana), Musa acuminata (edible banana), Nephelium lappaceum (rambutan), Nerium oleander (oleander), Olinia spp., Oryza sativa (rice), Osmanthus spp. (devilwood), Paspalum dilatatum (dallisgrass), Pelargonium spp. (geranium), Pennisetum americanum (pearl millet), Pennisetum glaucum (=P. typhoideum) (bulrush millet), Phaseolus spp. (beans), Phaseolus vulgaris (kidney bean), Philodendron spp. (philodendron), Plectroniella spp., Prosopis spp. (mesquite), Prunus spp. (plums, peaches), Psidium spp. (guava), Pyrus spp. (pear), Pyrus communis (common pear), Ricinus spp., Ricinus communis (castor bean), Rosa spp. (rose), Saccharum spp. (sugarcane), Saccharum officinarum (sugarcane), Samanea saman (=Albizia saman) (raintree), Schinus spp. (pepper tree), Schinus terebinthifolius (Brazilian pepper tree), Sorghum spp., Sorghum bicolor (=S. vulgare) (sorghum), Swietenia spp. (mahogany), Tamarix spp. (tamarisk), Triticum aestivum (wheat), Vaccinium spp. (blueberry), and Zea mays (corn) (Gough, 1913; Akanbi, 1973; Gubbaiah, 1984; Zhang, 1994; Molina, 1998; reviewed in Silva and Mexia, 1999; McQuate et al., 2000; Zheng et al., 2006; Robinson et al., 2010; reviewed in CABI, 2012).

Ben Yehuda et al. (1991) give a host list for this species.

Interestingly, this species has also been recorded as a predator of whiteflies and other Homoptera, including Aleurocanthus woglumi and Stictococcus spp. (Clausen, 1940; Zhang, 1994). It is also capable of developing on honeydew alone (McQuate et al., 2000).

*Moore (2003) states that this species does not complete its lifecycle on lemon.

Known Vectors (or associated organisms) Cryptoblabes gnidiella is associated with different mealybug species including the citrus mealybug (Planococcus citri) and Green’s mealybug (Avidov and Harpaz, 1969). In grapes, this species usually attacks fruits following attacks by the European grapevine moth (Lobesia botrana) (Carter, 1984).

When grapes are attacked by larvae close to harvesting, juice leakage can lead to rot by secondary pathogens which can reduce wine quality (Botton et al., 2003). Grape grey mold can increase with high population densities of C. gnidiella on grape (reviewed in Silva and Mexia, 1999).

28 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

On dallisgrass (Paspalum dilatatum), this species is associated with the ergot fungus Claviceps paspali, which produces honeydew-like secretions on the host (Ben Yehuda et al., 1991).

Aside from its pest status, this species could serve as a potential vector of Botrytis cinerea for control of the weed Myrica faya (Duffy and Gardner, 1994).

Known Distribution Asia: India, Indonesia, Israel, Lebanon, Malaysia, Pakistan, Russia, Thailand, and Turkey; Africa: Congo, Egypt, Liberia, Malawi, Morocco, Nigeria, Sierra Leone, South Africa, and Zaire; Caribbean: Bermuda; Europe: Austria, Cyprus, France, Gibraltar, Greece, Italy (including Sardinia and Sicily), Malta, Portugal (including Azores and Madeira), and Spain (Canary Islands), and Ukraine; Oceania: Fiji, Hawaii, and New Zealand; South America: Brazil and Uruguay (reviewed in Wysoki, 1986; Zhang, 1994; reviewed in Silva and Mexia, 1999; Nuss et al., 2011; reviewed in CABI, 2012; Evenhuis, 2013).

This species is likely more widespread than reported in the literature as many interceptions at U.S. ports of entry have originated on material from countries not reported to have the pest, including: Albania, Bosnia and Herzegovina, Cote D’Ivoire, Croatia, Cuba, Dominican Republic, Eritrea, Georgia, Germany, Haiti, Iran, Iraq, Jordan, Kuwait, Mexico, Netherlands, Qatar, Romania, Saudi Arabia, South Korea, Syrian Arab Republic, Tunisia, United Kingdom*, Venezuela, former Yugoslavia (AQAS, 2013; queried March 8, 2013).

*According to Carter (1974), this species is not established in the United Kingdom as it cannot survive; it is likely a transient species.

Potential Distribution within the United States If introduced into the United States, this species is most likely to be found wherever associated coccids are found. Cryptoblabes gnidiella is associated with many commercial crops found in the United States, including: citrus, grapevine, cotton, and corn. These hosts are found throughout most of the United States.

Pathway Although this species is native to the Mediterranean region, it has been introduced into Malaysia, New Zealand, Hawaii, and parts of tropical and subtropical America. It is sometimes found in imported material in the British Isles (Beirne, 1952; Carter, 1984). Mau and Kessing (1992) state that this species most likely dispersed throughout Europe on infested fruit.

This species has been intercepted at U.S. ports of entry over 450 times. Almost all interceptions occurred on plant material, including: Punica granatum (319), Citrus sp. including C. sinensis (50), Psidium guajava (32), and Ananas sp. (8). Most

29 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

interceptions occurred in baggage (394), with the rest occurring on general cargo (38), permit cargo (11), and other (13). Intercepted material has originated from many different countries around the world, including: Italy (57), Jordan (55), Greece (49), Israel (48), Spain (33), Iran (30), former Yugoslavia (19), and Portugal (17) among others (AQAS, 2013; queried March 8, 2013).

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination. Both the wing trap and plastic bucket trap are approved traps for this species. The wing trap is available in a plastic or paper version; either type may be used for this target. The lure is effective for 28 days.

Any of the following Trap Product Names in the IPHIS Survey Supply Ordering System may be used for this target: 1) Wing Trap Kit, Paper 2) Wing Trap Kit, Plastic 3) Plastic Bucket Trap

The Lure Product Name is “Cryptoblabes gnidiella Lure.”

IMPORTANT: Do not include lures for other target species in the trap when trapping for this target.

Trap spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Survey Site Selection: This species has a wide host range that includes citrus, corn, cotton, grapevine, loquat, pomegranate, and sorghum (Avidov and Harpaz, 1969). Surveys should be focused on areas where host plants are present. These can include field crops, nurseries, and even some residential areas.

Time of year to survey: In Israel, males begin appearing in pheromone traps in low numbers in early spring. The number of adult males captured greatly increases from June to October (Harari et al., 2007). Ben Yehuda et al. (1991) trapped moths from March through December. The percentage of the total moths caught over this time period was: March to April (5% of total moths captured), June to September (75%), and October to December (20%). In Portugal, moths are trapped from March until late May (overwintering population) and trap catches increase again from early June until late August (first generation population). Moths are trapped until late November/early December (Silva and Mexia, 1999).

30 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

Literature-Based Methods: Trapping: Early work on pheromones for this species found that the synthetic compounds: Z11-16:Ald, El 1-16:Ald, Z13-18:Ald, and E13-18:Ald in 100:10:100:10 proportions was more attractive to adult males than virgin females (Bjostad et al., 1981).

Field work by Anshelevich et al. (1993) found that “at dosages of 2 or 0.2 mg, a binary blend containing (Z)-I 1-hexadecenal (Z11-16:Ald) and (Z)-13-octadecenal (Z13-18:Ald) (1:1) was as effective in attracting males as a quaternary blend containing Z11-16:Ald, (E)-11-hexadecenal (E11-16:Ald), Z13-18:Ald and (E)-13-octadecenal (E13-18:Ald), (10:1:10:1)”. The suggested replacement rate of the lure was two to three weeks. Anshelevich et al. (1993) found that the nonsticky IPS trap was as effective in capturing C. gnidiella adult males as the sticky Pherocon 1C trap.

Pheromones have been used in certain countries, including Israel, to develop mating disruption techniques against this species (Gordon et al., 2003).

Ben Yehuda et al. (1991) used delta traps placed trees at a height of 1.5 m when surveying for this species in avocado. Silva and Mexia (1999) used one funnel trap per grove when trapping for this species in Portugal. Traps contained pheromone dispensers (replaced every four weeks) and an insecticidal strip (replaced every week).

Not recommended: This species is attracted to light (Goater, 1986); however, this survey method is not species specific.

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of C. gnidiella is by morphological identification. Adults of C. gnidiella are easily confused with many North American species; genitalia must be examined for confirmation. Mature larvae and adults can be identified. A screening aid for CAPS target Pyraloidea (males), including C. gnidiella, can be found in Passoa (2009). A description of this species is found in Neunzig (1986).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Easily Confused Pests Larvae could be confused with larvae of the carob moth (Ectomyelois ceratoniae) which is present in parts of the United States (Nay and Perring, 2008). This species also attacks citrus fruit but can cause much more extensive fruit drop than C. gnidiella (Avidov and Gothilf, 1960; Avidov and Harpaz, 1969). A diagnostic comparison between the two pests can be found in Avidov and Gothilf (1960).

Pre-imaginal stages of Lobesia botrana and C. gnidiella can look similar. Both can occur simultaneously on grapevines (CABI, 2012). Tio et al. (1994) give characteristics to distinguish the two larval species.

31 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

A key to Pyraloidea larvae (including Cryptoblabes spp.) is provided in Solis (2006).

Commonly Encountered Non-targets In Italy, Bagnoli and Lucchi (2001) caught the following species when trapping for Cryptoblabes gnidiella using the 4-component pheromone lure. However, CAPS surveyors will use the 2-component lure, so it is unknown if these species will be attracted.

Duponchelia fovealis Zeller (Pyralidae, Pyraustinae)*, Grapholita funebrana (=Cydia funebrana) Treitschke (Tortricidae, Olethreutinae), Hoplodrina ambigua (Den. & Schiff.) (Noctuidae, Amphipyrinae) Mamestra dysodea (Den. & Schiff.) (Noctuidae, Hadeninae), Mythimna vitellina (Hübner) (Noctuidae, Hadeninae), Pammene inquilina Fletcher (Tortricidae).

*This species is present in the United States (Stocks and Hodges, 2013).

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Ascher, K. R. S., M. Eliyahu, E. Gurevitz, and S. Renneh. 1983. Rearing the honeydew moth, Cryptoblabes gnidiella, and the effect of diflubenzuron on its eggs. Phytoparasitica 11(3-4): 195-198.

Avidov, Z. and I. Harpaz. 1969. Plant Pests of Israel. Israel Universities Press, Jerusalem.

Avidov, Z. and S. Gothilf. 1960. Observations on the honeydew moth (Cryptoblabes gnidiella Milliere) in Israel. Ktavim, 10:109-124.

Bagnoli, B. and A. Lucchi. 2001. Bionomics of Cryptoblabes gnidiella (Millière) (Pyralidae Phycitinae) in Tuscan vineyards (abstract). Proceedings of the IOBC/WPRS Working Group “Integrated Control in Viticulture” at Ponte de Lima, Portugal. March 2-7, 2001.

Beirne, B. P. 1952. British Pyralid and Plume Moths. Frederick Warne & Co., Ltd. London and New York. 208 pp.

Ben Yehuda, S., M. Wysoki, and D. Rosen. 1991. Phenology of the honeydew moth, Cryptoblabes gnidiella (Milliere) (Lepidoptera: Pyralidae), on avocado in Israel. Israel Journal of Entomology Vol. XXV- XXVI: 149-160.

Ben Yehuda, S., M. Wysoki, and D. Rosen. 1993. Laboratory evaluation of microbial pesticides against the honeydew moth (abstract). Insect Science and its Application 14(5): 627-630.

Bisotto-de-Oliveira, R., L. R. Redaelli, J. Sant’Ana, and M. Botton. 2007. Parasitoids associated with Cryptoblabes gnidiella (Lepidoptera, Pyralidae) in grapevine, state of Rio Grande do Sul, Brazil (abstract). Arquivos do Instituto Biológico (São Paulo) 74 (2) São Paulo: Instituto Biológico 115-119.

32 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

Bjostad, L. B., E. Gurevitz, S. Gothilf, and W. L. Roelofs. 1981. Sex attractant for the honeydew moth, Cryptoblabes gnidiella. Phytoparasitica 9(2): 95-99.

Botton, M., S. de J. Soria, E. R. Hickel. 2003. Uvas Viníferas para processamento em Regiões de Clima Temperado (in Spanish). Embrapa Uva e Winho. Sistema de Produção 4: 32 pp.

CABI. 2012. Cryptoblabes gnidiella (citrus pyralid). Crop Protection Compendium. Accessed March 8, 2013 from: www.cabi.org/cpc.

Carter, D. J. 1984. Pest Lepidoptera of Europe with special reference to the British Isles. Dr W. Junk Publishers

Chandel, B. S., A. P. S. Bhadauriya, and A. K. S. Chauhan. 2010. Relative toxicity of certain insecticides against larvae of earhead caterpillar, Cryptoblabes gnidiella Milliere (Lepidoptera: Pyralidae) (abstract). J. Env. Bio-Sci. 24(1): 95-98.

Clausen, C. P. 1940. Entomophagous Insects. McGraw-Hill, New York. 688 pp.

Duffy, B. K. and D. E. Gardner. 1994. Locally established botrytis fruit of Myrica faya, a noxious weed in Hawaii (abstract). Plant Disease 78(9): 919-923.

El-Zemaity, M. S., S. I. Salama, A. A. Ahmed, and E. R. Mahmoud. 2009. Some factors affecting the susceptibility of four garlic cultivars to infestation with Cryptoblabes gnidiella Milliere. Arab Universities Journal of Agricultural Sciences 17(1): 185-192.

EPPO. 2004. Good Plant Protection Practice, Citrus. EPPO Bulletin 34: 43-56.

Evenhuis, N. L. 2013. Checklist of Fijian Lepidoptera. Accessed March 14, 2013 from: http://hbs.bishopmuseum.org/fiji/checklists/lepidoptera.html.

Gentry, J. W. 1965. Crop Insects of Northeast Africa-Southwest Asia. Agricultural Handbook No. 273. United States Department of Agriculture, Agricultural Research Service.

Goater, B. 1986. British Pyralid Moths, A Guide to their Identification. Harley Books. England. 175 pp.

Gordon, D., L. Anshelevich, T. Zahavi, S. Ovadia, E. Dunkelblum, and A. Harari. 2003. Integrating mating disruption techniques against the honeydew moth and the European grapevine moth in vineyards (abstract). Proceedings of the IOBC/WPRS working group 'Integrated Protection and Production in Viticulture', Volos, Greece, March 18-22, 2003.

Gough, L. H. 1913. A new cotton insect (abstract). Bull. Soc. ent. Egypte 6(1): 19-20.

Gubbaiah. 1984. Cryptoblabes gnidiella, a fern-feeding caterpillar, and its parasites (abstract). International Rice Research Newsletter 9(6): 20-21.

Harari, A. R., T. Zahavi, D. Gordon, L. Anshelevich, M. Harel, S. Ovadia, and E. Dunkelblum. 2007. Pest management programmes in vineyards using male mating disruption. Pest Management Science 63: 769-775.

Hashem, A. G., A. W. Tadros, and M. A. Abo-Sheasha. 1997. Monitoring the honeydew moth, Cryptoblabes gnidiella Mill in citrus, mango and grapevine orchards (Lepidoptera: Pyralidae) (abstract). Annals of Agricultural Science (Cairo) 42(1): 335-343.

Heckford, R. J. and P. H. Sterling. 2004. Notes on, and descriptions of, some larvae of Oecophoridae, Gelechiidae and Pyralidae (Lepidoptera). Entomologist's Gazette 55(3): 143-159.

33 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

MALR. No date. Mango. Ministry of Agriculture and Land Reclamation (MALR), Central Administration of Plant Quarantine (CAPQ), Cairo, Egypt. 91 pp.

Mau, R. F. L. and J. L M. Kessing. 1992. Crop Knowledge Master. Cyptoblabes [sic] gnidiella. Christmas berry webworm. Accessed March 8, 2013 from: http://www.extento.hawaii.edu/kbase/Crop/Type/cryptobl.htm.

McQuate, G., P. A. Follett, and J. M. Yoshimoto. 2000. Field infestation of rambutan fruits by internal- feeding pests in Hawaii. Journal of Economic Entomology 93(3): 846-851.

Molina, J. M. 1998. Lepidoptera associated with blueberry cultivation in Western Andalusia (abstract). Boletín de Sanidad Vegetal, Plagas 24(4): 763-772.

Moore, S.D. 2003. The lemon borer moth: a new citrus pest in South Africa (abstract). SA Fruit Journal 2(5): 37-41. Paarl: Deciduous Fruit Producers Trust.

Nay, J. E. and T. M. Perring. 2008. Blue Palo Verde, Parkinsonia florida (Benth. Ex Grey) S. Wats. (Fabaceae), a new host of the introduced carob moth, Ectomyelois ceratoniae (Zeller) (Lepidoptera: Pyralidae), in southern California (abstract). Pan-Pacific Entomologist 84(2):143-145.

Neunzig, H. H. 1986. The Moths of America North of Mexico. Fascicle 15.2. Pyraloidea, Pyralidae (Part). The Wedge Entomological Research Foundation.

Nuss, M., W. Speidel, and A. Segerer. 2011. Fauna Europaea: Cryptoblabes gnidiella. Fauna Europaea version 2.5. Accessed March 21, 2013 from: http://www.faunaeur.org.

Özturk, N. and M. R. Ulusoy. 2009. Pests and Natural Enemies Determined in Pomegranate Orchards in Turkey. Proc . Ist IS on Pomegranate. Acta Hort. 818: 277-284.

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 Maryland, March 2009.

Robinson, G. S., P. R. Ackery, I. J. Kitching, G. W. Beccaloni, and L. M. Hernández, 2010. HOSTS - A Database of the World's Lepidopteran Hostplants. Natural History Museum, London. Accessed March 21, 2013 from: http://www.nhm.ac.uk/hosts.

Salama, S. I. 2008. Biological studies on Cryptoblabes gnidiella (Lepidoptera: Pyralidae) infesting stored garlic. Annals Agric. Sci., Ain Shams Univ., Cairo, 53(2): 397-402.

Sellanes, C., C. Rossini, and A. González. 2010. Formate analogs as antagonists of the sex pheromone of the honeydew moth, Cryptoblabes gnidiella: electrophysiological, behavioral and field evidence. Journal of Chemical Ecology 36: 1234-1240.

Silva, E. B. and A. Mexia. 1999. The pest complex Cryptoblabes gnidiella (Millière) (Lepidoptera: Pyralidae) and Plannococcus citri (Risso) (Homoptera: Pseudococcidae) on sweet orange groves (citrus sinensis (L.) Osbeck) in Portugal: Interspecific association. Bol. San. Veg. Plagas, 25: 89-98.

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 lepidopterous larvae” by Weisman 1986. 58 pp.

Stocks, S. D. and A. Hodges. 2013. Featured Creatures, European pepper moth - Duponchelia fovealis. University of Florida. EENY-508. Accessed August 14, 2013 from: http://entnemdept.ufl.edu/creatures/veg/leps/european_pepper_moth.htm.

34 Cryptoblabes gnidiella Primary Pest of Grape Arthropods Honeydew moth Moth

Tio, R. del, R. Ocete, and M. E. Ocete. 1994. Practical indications for differentiating preimaginal stages of Lobesia botrana (Denis & Schiffermuller, 1775) (Lepidoptera: Tortricidae) and Cryptoblabes gnidiella (Milliere, 1867) (Lepidoptera: Pyralidae), two vine pests in Jerez, Spain. Sociedad Hispano-Luso- Americana de Lepidopterologia (SHILAP) 22(86):119-125.

Wysoki, M. 1986. New records of lepidopterous pests of macadamia in Israel. Phytoparasitica 14(2): 147.

Wysoki, M., B. S. Yehuda, and D. Rosen. 1993. Reproductive behavior of the honeydew moth, Cryptoblabes gnidiella (abstract). Invertebrate Reproduction and Development 24(3): 217-224.

Wysoki, M., E. Swirski, and Y. Izhar. 1981. Biological control of avocado pests in Israel (abstract). Protection Ecology 3(1): 25-28.

Wysoki, M., M. de Jong, and S. Rene. 1988a. Trichogramma platneri Nagarkatti (Hymenoptera: Trichogrammatidae), its biology and ability to search for eggs of two lepidopterous avocado pests, Boarmia (Ascotis) selenaria (Schiffermüller) (Geometridae) and Cryptoblabes gnidiella (Milliere) (Phycitidae) in Israel (abstract). Colloques de l'INRA (43): 295-301.

Wysoki, M., P. de Haan, and Y. Izhar. 1988b. Efficacy of Bacillus thuringiensis preparations containing dead and live spores against two avocado pests: the giant looper, Boarmia selenaria (Lep.: Geometridae) and the honeydew moth, Cryptoblabes gnidiella (Lep.: Phycitidae). Crop Protection 7:131-136.

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35 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle 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 through 2012

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 (0.029 x 0.014), 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 (0.047 in.) long, and mature third instars are about 8.5 mm (0.33 in.) 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.

36 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle

Pupae: Pupae are 5.8 to 7.1 mm (0.23 to 0.28 in.) long and white. Females with a pair of tubercles near the apex. Mature third instars build an 8 x 4 mm (0.31 x 0.16 in.) oval cell in the soil in which they pupate, and tenerals remain for about three 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 (0.22 to 0.29 in.) long; antennae 4 to 5 mm (0.16 to 0.20 in.) (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 Figure 1. Adult Diabrotica speciosa. Photo transverse spots, basal spots larger courtesy of Hernan Tolosa. and usually reddish toward the 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 (81°F) takes place after about eight 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 (corn) 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 root system, but mature larvae tend to congregate in the upper 10 cm (3.94 in.) of the root under the crown. The larval stage lasts 23 to 25 days (~12 days in laboratory conditions at 25°C (77°F)), including an inactive prepupal period of two-to-three days. At 25°C, the pupal stage lasts six days, and is followed by a period of three to five days during which the recently molted adults remain in the pupal cell, presumably for the cuticle to tan (USDA, 1957).

Young beetles have a yellowish or pale brown color, which turns green with bright yellow spots in three days if fresh food is provided. Under laboratory conditions, mating has been observed between four and six days after emergence, and some females were observed mating again at day 35. Each female laid an average of 1164 eggs

37 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle

during her lifetime, starting on day eight and extending for a maximum of 77 days. Peak oviposition was observed on days 16 through 56. In a laboratory environment, oviposition on 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 roots, 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 dahlias and chrysanthemums (USDA, 1957). Economic thresholds of two insects per plant for Phaseolus vulgaris (bean) 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 nutrient uptake. In corn, attack on young plants by larvae produces a

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

‘Gooseneck’ growth form of corn. Photo On corn, the most economically Figure 2. courtesy of The Ohio State University. 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 are especially associated with 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).

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). 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

39 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle

controlled as beetles can also be observed feeding on and moving into grape from surrounding weeds.

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), Arachis hypogaea (peanut), Artemisia spp. (absinthium, tarragon), Asparagus spp. (asparagus), Avena spp. (oats), Baccharis articulata, Beta vulgaris (beet), Brassica spp. (mustards), Bromus catharticus (prairie grass), Camellia sinensis var. sinensis (tea), Capsicum annuum (pepper), Capsicum frutescens (pepper), Carica papaya (papaya), Cattleya spp. (orchid), Cayaponia spp., Chenopodium spp., Chrysanthemum spp., Cichorium spp. (chicory, endive), Citrullus lanatus (watermelon), Citrus spp., Coriandrum sativum (coriander), Cucumis spp. (melons, cucumbers, gerkins), Cucurbita spp. (cucurbits), Cucurbitella asperata, Cynara spp. (artichoke), Cynodon dactylon (Bahama crass), Dahlia pinnata (pinnate dahlia), Datura 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), Ipomoea purpurea (common morning glory), Lactuca sativa (lettuce), Lagenaria siceraria (bottle gourd), Lavandula angustifolia subsp. angustifolia (English lavender), Lepidium didymum (twin cress),Lilium maculatum (sukash-yuri), Linum usitatissimum (flax), Lolium perenne (rye grass), Luffa spp. (loofah), , 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), Origanum vulgare (oregano), Oryza sativa (rice), Passiflora coerulea (passion flower), Petroselinum crispum (parsley), , Phaseolus spp. (beans), Physalis viscose (starhair groundcherry), Pimpinella anisum (anise), Pisum sativum (pea), Prunus spp. (stone fruit), Raphanus sativus (radish), Rosa spp. (rose), Sechium edule (chayote), Sicyos polyacanthus, Solanum spp., Solanum betaceum (tree tomato), Solanum lycopersicum (tomato), Solidago chilensis (goldenrod), Sorghum spp., Spinacia oleracea (spinach), Taraxicum officinale (dandelion), Trifolium spp. (clover),

40 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle

Tropaeolum majus (Nasturtium), Vaccinium virgatum (smallflower blueberry), Vaccinium corymbosum (highbrush blueberry), 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 virus), carmoviruses, and purple granadilla mosaic virus (Ribeiro et al., 1996; Germain, 2000). Lin et al. (1984) showed that D. speciosa transmitted cowpea 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. A recent risk map developed by USDA-APHIS-PPQ-CPHST (Fig. 4) indicates that many states in the United States have a low to moderate risk rating for D. speciosa establishment based on host availability and climate within the continental United States. Areas of Arkansas, Illinois, Indiana, Kansas, Kentucky, Louisiana, Mississippi, North Carolina, Oklahoma, and Tennessee have the highest risk of D. speciosa establishment.

Survey CAPS-Approved Method*: Visual inspection is the approved method to survey for D. speciosa.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Visual survey: Visual detection of adults is easy, as their feeding period spans from Figure 4. Adult banded cucumber beetle, dawn until dusk. Detection of larval Diabrotica balteata. Photo courtesy of John damage, on the other hand, is more L. Capinera, University of Florida 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

41 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle 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 of D. speciosa appear to be universally attracted to aromatic compounds from squash blossoms, though the specific compound(s) that attract the beetles varies from species 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 genus. 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.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/. Figure 5. Western corn rootworm, Diabrotica Literature-Based Methods: virgifera. Courtesy of USDA-ARS. Diabrotica speciosa somewhat resembles the other main pestiferous 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

42 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle

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 hosts of D. speciosa, could be attributed to several other root feeders, such as wireworms (Conoderus spp.; Elateridae), white grubs, (Phytalus spp., Cyclocephala spp., Diloboderus abderus; Melolonthidae), Pantomorus spp. and Listronotus bonariensis (Curculionidae), and several Figure 6. Southern corn rootworm, chrysomelids (Caeporis spp., Colaspis Diabrotica undecimpunctata. Courtesy of spp., Maecolaspis spp., Diphaulaca Clemson University - USDA Cooperative spp. and Cerotoma arcuata) (Gassen, Extension Slide Series, www.bugwood.org. 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.

43 Diabrotica speciosa Primary Pest of Grape Arthropods Cucurbit beetle Beetle

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.

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 leaf beetle, 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.

44 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Epiphyas postvittana

Scientific Name Epiphyas postvittana Walker

Synonyms: Austrotortrix postvittana, Dichelia foedana, D. retractana, D. reversana, D. vicariana, D. vicaureana, Pandemis consociana, Teras basialbana, T. scitulana, T. secretana, Tortrix dissipata, T. oenopa, T. phaeosticha, T. pyrrhula, and T. stipularis.

Common Names Light brown apple moth (LBAM), apple leafroller, Australian leafroller

Type of Pest Moth

Taxonomic Position Class: Insecta Order: Lepidoptera Family: Tortricidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List -2003 through 2008; PPQ Program Pest

Pest Description Eggs: Epiphyas postvittana egg masses (Fig. 1) are flat, broadly oval, translucent, and appear pale yellow to white in color (Brown et al., 2010). The chorion is reticulated, which separates eggs of this species from some, but not all, tortricids in North America Figure 1: Eggs of LBAM on a (Peterson, 1965). There are approximately 35 eggs leaf surface. Photo courtesy of T. in a mass, overlapping like “roof tiles or shingles”. M. Gilligan & M. E. Epstein, Females lay on average 100 to 300 eggs beginning LBAM ID (CSU, CDFA, and at two to three days of age. USDA/APHIS/ PPQ/CPHST).

Larvae: First instar larvae (Fig. 2) are approximately 1.5 to 1.6 mm (~0.06 in.) long with a dark head and light-colored body, and final instar larvae range from 10 to 20 mm (0.39 to 0.79 in.) in length. The body of a mature larva is yellowish green with paler subdorsal, subventral, and ventral lines. The first larval instar has a dark-brown head; all other instars have a light-fawn head and prothoracic plate. The succeeding instars have a darker body than fully grown instars. The head, prothoracic shield, legs, and anal plate are pale brown, the genal dash is present or absent, and the prothoracic shield is only slightly darker than the rest of the integument. All instars are dark dorsally, and the pinacula of later instars are paler than the surrounding integument (Gilligan and Epstein, 2009; Brown et al., 2010).

45 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Pupae: Pupae (Fig. 3) are green after pupation, but become brown within one day. Male pupae average 2.5 by 7.6 mm (0.098 to 0.30 in.); females average 2.9 by 9.8 mm (0.11 to 0.39 in.). The pupal stage is completed within the “nests” made up of rolled up leaves (Gilligan and Epstein, 2009; Brown et al., 2010)

Adult: Epiphyas postvittana is sexually dimorphic. Forewings of both sexes are light brown to pale yellow with brown to dark brown markings. Male light brown apple moth adults are usually smaller than females. Male forewing length ranges from 5.3 to 11.1 mm (0.21 to 0.44 in.), compared with 5.4 to 12.5 mm (0.21 to 0.49 in.) in females (Gilligan and Epstein, 2009). Males are more variable than females, although in most males the basal half of the forewing is lightly marked, the median fascia is well defined, and there is a dark mark on the costa distal to the median fascia. In California, males tend to be of three phenotypes (Fig. 4); the form with solid dark markings on the distal half of the forewing is the most uncommon. All males have a forewing costal fold. The female (Fig. 4) forewing color is more uniform, with a poorly defined median fascia and overall mottled or speckled appearance. Most females have a dark mark on the dorsum of each forewing and two dark spots on the posterior of the thorax. All females lack a forewing costal fold. The hindwing in both males and females is mottled with dark scales, although this pattern is usually more evident in females. Adults in other areas of the world can have incredibly variable forewing patterns (Gilligan and Epstein, 2009).

Male genitalia (Fig. 5) are distinctive and a dissection can be used to verify male specimen identity. Males possess a combination of the following characters: spatulate Figure 2: Early- (top), (spoon-shaped) uncus; reduced socii; short valva with a mid- (middle), and late- broad sacculus; membranous lobe on the apex of the (bottom) instar larvae of valva; and an aedeagus with 2 to 4 deciduous cornuti LBAM. Photos courtesy of (Gilligan and Epstein, 2009). T. M. Gilligan & M. E. Epstein, LBAM ID (CSU, Female genitalia (Fig. 5) are typical of many Archipini and CDFA, and females may be difficult to verify based on dissection USDA/APHIS/PPQ/ alone. LBAM females possess a combination of the CPHST). following characters: simple sterigma; long, straight ductus bursae which is 2/3 or more the length of the abdomen; and corpus bursae with a single, hook-shaped signum (Gilligan and Epstein, 2009).

46 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Biology and Ecology Epiphyas postvittana has two to four annual generations over much of its range; the exact number of generations varies by latitude. There is considerable overlap between generations, with development driven by temperature and larval host plant (Danthanarayana, 1975; Geier and Briese, 1980; Thomas, 1989). The highest rate of population increase was on Plantago lanceolata (ribwort plantain), followed by Rumex crispus (curly dock), apples (Malus domestica cv. Granny Smith) and Trifolium repens (white clover) (Danthanarayana et al., 1995). In northern New Zealand, four overlapping generations occur, with adults : LBAM Pupa. Photo flying during September to October, Figure 3 courtesy of T. M. Gilligan & M. E. December to January, February to March, Epstein, LBAM ID (CSU, CDFA, and and April to May. In southern Australia, three USDA/APHIS / PPQ/ CPHST). overlapping generations are completed, with adults flying during December to January, April to May, and September to October.

Populations in California appear to complete at least four overlapping generations, with adults present almost continuously from March to November. The upper and lower temperature thresholds for E. postvittana development have been determined to be 7.5°C (46°F) and 31°C (88°F) in laboratory studies, with an ideal development temperature of 20°C (68°F) (Danthanarayana, 1975; Brown et al., 2010).

Females lay eggs in a mass that contains from four to 96 eggs (mean 35) overlapping individual eggs (Wearing et al., 1991). Females deposit eggs at night (USDA, 1984). Eggs are laid on the upper surface of host plants with smooth leaf surfaces; females will refrain from depositing eggs on hairy leaves (Danthanarayana, 1975; Geier and Briese, 1981; Foster and Howard, 1998). Females often select the depression along the upper side midrib of leaves (Powell and Common, 1985). Egg development time varies with temperature and eggs will hatch in approximately eight to nine days at 20°C (68°F) (Gilligan and Epstein, 2009).

Larvae pass through five to six instars during their development. Larvae do not overwinter, although development during colder months is slower. The rate of development varies with temperature and host plant utilized; larval development takes approximately 25 days at a temperature of 20°C (68°F). Early instar larvae feed on the underside of leaves within a silk chamber. Later instar larvae may fold single leaves, create a nest of several leaves webbed together, or web leaves to fruit and feed on the surface.

47 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Pupation occurs within the larval nest. Complete pupal A B development takes approximately 10 days at a temperature of 20°C (Danthanarayana, 1975). Adult moths emerge after one to several weeks of pupation. Female moths emerge from protective pupal nests and mate soon after emergence (Geier and Briese, 1981). Danthanarayana (1975) C D suggests the preoviposition period is 2 to 7 days. Females copulate for slightly less than one hr. (Foster et al., 1995). Oviposition does not begin until females are two- to three-days old (Geier and Briese, 1981). The oviposition period lasts from one to 21 days Figure 4. A. Typically marked male. B. Male with dark (Danthanarayana, 1975). wings. C. Make with light wings. D. Typically marked Adult longevity is influenced female. Photos courtesy of T. M. Gilligan & M. E. Epstein, by host plant and LBAM ID (CSU, CDFA, and USDA/APHIS / PPQ/ temperature. In the CPHST). laboratory, female longevity can vary between 10 days (Geier and Briese, 1981) and 32.7 days (Danthanarayana, 1975); males can live up to approximately 33 days (Danthanarayana, 1975). Under field conditions in Australia, the life span of adult E. postvittana is 2 to 3 weeks (Magarey et al., 1994).

Moths are quiescent during the day and may be found on foliage of hosts (Geier and Briese, 1981). Long distance dispersal is typically achieved by adults (Geier and Briese, 1980; Suckling et al., 1994), although larval dispersal occurs over a short range. Flight occurs at dusk in calm conditions (Geier and Briese, 1981; USDA, 1984; Magarey et al., 1994). Adults are unlikely to disperse from areas with abundant, high-quality hosts (Geier and Briese, 1981). Males will disperse farther than females. In a mark- release-recapture study, 80% of recaptured males and 99% of recaptured females occurred within 100 m (328 ft.) of the release point (Suckling et al., 1994). Females do not appear to rely on plant volatiles to locate a host, but tactile cues are important (Foster and Howard, 1998). Humidity influences the dispersal ability of the pest (Danthanarayana et al., 1995).

48 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Although they are sheltered in silk, first instar larvae are more exposed to weather and insecticide treatments than are second and third instar larvae (Madge and Stirrat, 1991; Lo et al., 2000). After approximately three weeks, larvae leave the silken tunnels for a new leaf (USDA, 1984). Second and later instars have the ability to create their own protective feeding shelter by rolling a leaf or webbing multiple leaves together (Danthanarayana, 1975; Lo et al., 2000); behaviors characteristic of the Tortricidae.

Larvae move vigorously when disturbed, but are always connected to the leaf by a silken thread to avoid being removed from the leaf (Nuttal, 1983; USDA, 1984). When larvae happen to fall to the ground, they feed on ground-cover hosts or can survive without feeding for several months (Evans, 1937; Thomas, 1975; USDA, 1984).

E. postvittana is more abundant during the second generation than during other generations (MacLellan, 1973; Madge and Stirrat, 1991). Thus, the second generation causes the most economic damage (Evans, 1937; Thomas, 1975; Madge and Stirrat, 1991; Lo et al., 2000) as larvae move from foliage to fruit (MacLellan, 1973; Magarey et al., 1994). Figure 5. Top: Male genitalia. Symptoms/Signs Bottom: Female genitalia. Photos The insect will feed on foliage (Fig. 6), courtesy of T. M. Gilligan & M. E. flowers, and fruit. In spring, the pest feeds on Epstein, LBAM ID (CSU, CDFA, new buds while later generations feed on and USDA/APHIS / PPQ/ CPHST). ripened fruits (Buchanan et al., 1991). After the first molt, they construct typical leaf rolls (nests) by webbing together leaves, a bud and one or more leaves, leaves to a fruit, or by folding and webbing individual mature leaves. During the fruiting season, they also make nests among clusters of fruits, damaging the surface and sometimes tunneling into the fruits (Danthanarayana, 1975).

Fruit surface feeding is common within larval nest sites and is typically caused by later instars (Lo et al., 2000). Clusters of fruit are particularly susceptible. E. postvittana has been shown to introduce Botrytis cinerea, a fungal that causes gray mold, spores into

49 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

wounds via contaminated larvae, with up to 13% of berry damage (by weight) as a result (Bailey, 1997). On a fruit, the calyx offers protection from parasitoids and is probably the best feeding location for young larvae (Lo et al., 2000). Larvae entering the fruit through the calyx may cause internal damage. Wet conditions may allow the entry of rot organisms. Feeding on the foliage by larvae causes ragging and curling of the foliage.

Damage to apples is in the form of either pinpricks, which are flask-shaped holes about 3 mm (0.12 in.) deep into the fruit, or entries, which are holes extending deeper than 3 mm into the fruit that leaves some frass and webbing at the Figure 6: LBAM typical damage to surface. On apples, skin damage or host plant foliage. Photo courtesy of blemishes have an irregular cork-like T. M. Gilligan & M. E. Epstein, LBAM appearance. Larvae may excavate small ID (CSU, CDFA, and USDA/APHIS / round pits and produce scars similar to PPQ/ CPHST). the ‘stings’ of the larvae of Cydia pomonella, the codling moth. The first generation (in spring) causes the most damage to apples; while the second generation damages fruit harvested later in the season (Terauds, 1977). Peaches are damaged by feeding that occurs on the shoots and fruit.

Pest Importance The larva of E. postvittana is a serious pest of fruit and ornamentals in Australia and New Zealand. As a pest of pome fruits, particularly apples, it probably ranks second to Cydia pomonella, the codling moth. During a severe outbreak, damage by E. postvittana to fruit may be as much as 75%. In Tasmania, this species is the most injurious pest of apples. In years of abundance, populations of the light brown apple moth may cause as much as 25% loss of the apple crop. This pest damages fruit in storage; a few larvae may ruin a whole case of fruit. The markings on the fruit render it unfit for export (USDA, 1984).

E. postvittana is a highly polyphagous pest that attacks a wide number of fruits, ornamentals, and other plants. According to Geier and Briese (1981), “Economic damage results from feeding by caterpillars, which may destroy, stunt or deform young seedlings, spoil the appearance of ornamental plants, and/or injure deciduous fruit-tree crops, citrus, and grapes.” Losses in Australia were estimated to be AU$21million (~US22.15 million) per year, but there has been no similar estimation in other countries.

The larvae can be very damaging to grape, apple, and peach. In grape, 70,000 larvae/ha were documented to cause a loss of 4.7 tons of chardonnay fruit in 1992 with

50 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

an estimated cost of $2000/ha (Bailey et al., 1995). A single larva can destroy about 30 grams of mature grapes.

Mature larvae are the most difficult stage to control. E. postvittana is also difficult to control with sprays because of its leaf-rolling ability, and because there is evidence of resistance due to overuse of sprays (Geier and Briese, 1981).

Known Hosts Epiphyas postvittana is a polyphagous pest and can damage nursery stock, stone fruit (peaches and apricots), pome fruits (apples and pears), grapes, and citrus. This pest can feed on >500 plant species in 121 families and 363 genera giving it the potential to become extremely destructive (Brown et al., 2010; Suckling and Brockerhoff, 2010). Larvae prefer herbaceous plants over woody ones (Brown et al., 2010).

Major Hosts: Acca sellowiana (feijoa fruit), Actinidia spp. (kiwi/Chinese gooseberry), Chrysanthemum spp. (chrysanthemum), Citrus spp. (citrus), Cotoneaster spp., Crataegus spp. (hawthorns), Diospyros spp. (malabar ebony), Eucalyptus spp. (eucalyptus), Humulus lupulus (hops), Jasminum spp. (jasmine), Ligustrum vulgare (privet), Litchi chinensis (lychee), Malus spp. (apple), Medicago sativa (alfalfa), Persea americana (avocado), Pinus spp. (pines), Pinus radiata (radiata pine), Populus spp. (poplars), Prunus armeniaca (apricot), Prunus persica (peach), Pyrus spp. (pears), Ribes spp. (currants), Rosa spp. (roses), Rubus spp. (blackberry, raspberry), Solanum spp. (potato/tomato), Trifolium spp. (clovers), Vaccinium spp. (blueberries), Vicia faba (broad bean), and Vitis vinifera (grapevine) (CABI, 2012).

Other Documented Hosts: Acacia spp. (wattles), Adiantum spp. (maidenhead fern), Alnus glutinosa (black alder), Amaranthus spp. (amaranth), Aquilegia spp. (columbine), Arbutus spp. (madrone), Arctotheca calendula (capeweed), Artemisia spp. (sagebrush), Astartea spp. (astartea), Aster spp. (aster), Baccharis spp. (baccharis), Billardiera spp. (billadriera), Boronia spp. (baronia), Brassica spp. (mustards), Breynia spp. (breynia), Bursaria spp. (bursaria), Buddleja spp. (butterfly bush), Calendula spp. (marigold), Callistemon spp. (bottlebrush), Camellia japonica (camellia), Campsis spp. (trumpet- vine), Cassia spp. (senna), Ceanothus spp. (red-root/lilac), Centhranthus spp. (fox- brush), Chenopodium album (lambsquarters/fat-hen), Choisya spp. (choisya), Clematis spp. (virgin’s-bower), Clerodendron spp. (glory-bower), Correa spp. (correa), Crocosmia spp. (montbretia), Cupressus spp. (cypress), Cydonia spp. (quince), Cytisus scoparius (Scotch broom), Dahlia spp. (dahlia), Datura spp. (thorn-apple), Daucus spp. (carrot), Dodonaea spp. (dodonea), Eriobotrya spp. (loquat), Eriostemon spp. (eristemon), Escallonia spp. (escallonia), Euonymus spp. (euonymus), Forsythia spp. (forsythia), Fortunella spp. (kumquat), Fragaria spp. (strawberry), Gelsemium spp. (jasmine), Genista spp. (broom), Gerbera spp. (daisy), Grevillea spp. (spider-flower), Hardenbergia spp. (hardenbergia), Hebe spp. (hebe/speedwell), Hedera spp. (ivy), Helichrysum spp. (everlasting), Hypericum perforatum (St. John’s wort), Juglans spp. (walnut), Lathyrus spp. (sweet pea), Lavandula spp. (lavender), Leucadendron spp. (leucodendron), Leptospermum spp. (manuka), Lonicera spp. (honeysuckle), Lupinus spp. (lupine), Macadamia spp. (macadamia), Mangifera spp. (mango), Melaleuca spp.

51 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

(bootlebrush), Mentha spp. (mint), Mesembryanthemum spp. (ice-plant), Michelia spp. (banana-shrub), Monotoca spp. (monotoca), Myoporum spp. (sandle-wood), Oxalis spp. (wood-sorrel), Parthenocissus spp. (ivy), Pelargonium spp. (geranium), Persoonia spp. (persoonia), Petroselinum spp. (parsley), Philadelphus spp. (mock-orange), Photinia spp. (photinia), Phyllanthus spp. (phyllanthus), Pittosporum spp. (pittosporum), Plantago lanceolata (plaintain/ribwort), Platysace spp. (platysace), Polygala spp. (milkwort), Polygonum spp. (knotweed), Pteris spp. (brake-fern), Pulcaria spp. (fleabane), Pyracantha spp. (fire-thorn), Quercus spp. (oak), Ranunculus spp. (buttercup), Raphanus spp. (radish), Reseda spp. (coneflower), Rumex spp. (dock), Salix spp. (willow), Salvia spp. (sage), Senecio spp. (ragwort), Sida spp. (side), Sisymbrium spp. (mustard), Smilax spp. (cat-brier), Tithonia spp. (sunflower), Trema spp. (trema), Triglochin spp. (arrow grass), Ulex europaeus (gorse), Urtica spp. (nettle), Viburnum spp. (arrow-wood), and Vinca spp. (periwinkle) (Danthanarayana, 1975; Wearing et al., 1991; Venette et al., 2003; Brown et al., 2010).

Known vectors (or associated organisms) An association between larvae of E. postvitanna and Botrytis cinerea (Fig. 7), gray mold, has been shown in grapes.

Figure 7. Discolored, shriveled berries caused by Botrytis bunch rot (left) and Botrytis cinerea sporulating on grape berries. Photos courtesy P. Sholberg, Agriculture & AgriFood Canada.

Known Distribution Epiphyas postvittana is indigenous to Australia. E. postvittana is widespread throughout Australia and New Zealand on many weedy hosts including gorse (Ulex europaeus) and broom (Cytisus scoparius). It is commonly present in gardens and unsprayed horticultural crops.

52 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Europe: Netherlands, Sweden, United Kingdom. North America: United States. Oceania: Australia and New Zealand (Meyrick, 1937; Bradley, 1973; Wolschrijn and Kuclein, 2006; Svensson, 2009).

Although it was reported from New Caledonia, its presence in that country could not be verified by Suckling and Brockerhoff (2010).

Potential Distribution within the United States E. postvittana has been reported to occur in Hawaii since 1896 (Zimmerman, 1978). On March 16, 2007, E. postvittana was confirmed in Alameda County, California. As of March 2012, further detections have occurred in Alameda, Contra Costa, Los Angeles, Fresno, Madera, Marin, Monterey, Napa, Sacramento, San Benito, San Diego, San Francisco, San Joaquin, San Luis Obispo, San Manteo, Santa Barbara, Santa Clara, Santa Cruz, Solano, Sonoma, Ventura, and Yolo Counties. A single moth of E. postvittana was detected in the summer of 2010 in Oregon. To date, despite extensive trapping, no additional moths have been trapped indicating that the moth is not established in Oregon.

A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates areas of California and the southern and southeastern United States have a moderate to high risk rating for E. postvittana establishment based on host availability and climate within the continental United States. Establishment is precluded in areas of the northern and northeastern United States.

Survey CAPS-Approved Method*: The CAPS approved method is a trap and lure combination. The preferred trap type is a Jackson trap. The lure is effective for 42 days (6 weeks).

In order to standardize data reporting and trap procurement for the LBAM Program, it is preferable that states use the Jackson trap. However, if states prefer to use the large plastic delta traps, the traps must be purchased with their own funding. Negative data may then be reported from the large plastic delta traps. Trap color is up to the state and does not affect trap efficacy.

Large plastic delta traps for Epiphyas postvittana should not be ordered through the IPHIS Survey Supply Ordering System.

IPHIS Survey Supply Ordering System Product Names: 1) Jackson Trap Body 2) Epiphyas postvittana Lure

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

53 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Lure Placement: Do not place lures for two or more target species in a trap unless otherwise recommended.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: (Taken from Venette et al., 2003 and CABI, 2009) Trapping: Pheromone traps have been widely used for detection and monitoring of populations of this species (Bellas et al., 1983). Two key components of the pheromone are (E)-11-tetradecenyl acetate and (E,E)-(9,11) tetradecadienyl acetate (Bellas et al., 1983). These compounds in a ratio of 20:1 are highly attractive to males. This lure is typically formulated on a rubber septum (1 to 3 mg). Due to the recent detections of E. postvittana in California, new formulations (e.g., plastic laminate) are under development and testing is planned at Otis. Delta traps have been used and placed from 5 to 6.5 ft. (1.5 to 2 m) above ground level.

Foster and Muggleston (1993) provide a detailed analysis of different designs of delta traps. In general, they found that traps with a greater length (i.e., the distance between the two openings of the trap) capture significantly more E. postvittana than shorter traps. This effect is not related to saturation of smaller sticky surfaces with insects or other debris. The addition of barriers to slow the exit of an insect from a trap also improves catch. In a separate analysis, Foster et al. (1991) found that placing the pheromone lure on the side of the trap helped to improve trap efficiency. The orientation of the trap relative to wind direction did not affect the number of E. postvittana that were attracted to the pheromone or were subsequently caught by the trap (Foster et al., 1991).

Visual survey: Visual inspections have been used to monitor population dynamics of E. postvittana eggs and larvae. In grape, 40 vines were inspected per sampling date (Buchanan, 1977). In apple and other tree fruits, 200 shoots and 200 fruit clusters (10 of each on 20 different trees) are often inspected (Bradley et al., 1998). Egg masses are most likely to be found on leaves (USDA ,1984). The egg masses may be jet black if parasitized by Trichogramma spp. (a trichogrammatid wasp) (Glenn and Hoffman, 1997). Larvae are most likely to be found near the calyx or in the endocarp; larvae may also create “irregular brown areas, round pits, or scars” on the surface of a fruit (USDA, 1984). Larvae may also be found inside furled leaves, and adults may occasionally be found on the lower leaf surface (USDA, 1984).

Not recommended: Adults are also attracted to fruit fermentation products as a 10% wine solution has been used as an attractant and killing agent for adults (Buchanan, 1977; Glenn and Hoffmann, 1997). The dilute wine (670 ml) in 1 liter jars was hung from grapevines on the edge of a block of grapes (Buchanan, 1977). Black light traps have been used to monitor adults of E. postvittana (Thwaite, 1976).

Key Diagnostics/Identification

54 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

CAPS-Approved Method*: Confirmation of E. postvittana is by morphological identification. Many native tortricids could be confused with E. postvittana. Identification requires dissection of male genitalia. Female specimens should be sent to a Lepidopteran specialist for identification. Sorting and Level 1 Screening may be performed without dissection by using Passoa et al. (n.d.).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: E. postvittana is similar to E. pulla and E. liadelpha, both not known to be present in the United States. Geier and Springett (1976) reported possible hybridization based on demographic characteristics. Larvae are similar to larvae of other leafrollers, which may be present (for example, in New Zealand, Planotortrix octo, P. excessana, Ctenopseustis obliquana, and C. herana may be present). Identity of the species must often be confirmed by examination of adult genitalia. Molecular diagnostics based on PCR amplification of ribosomal DNA have been developed and are especially useful for the identification of immature specimens (Armstrong et al., 1997).

TortAI: Tortricids of Agricultural Importance is designed for use by persons in the continental United States performing domestic surveys for exotic species. TortAI, which includes all of the tortricid species found in the digital identification tool LBAM ID, includes two image-rich interactive identification keys (adult and larvae), diagnostic fact sheets, a visual dictionary, support pages for the dissecting and preparing specimens, an image gallery, and a molecular search page. Because the world tortricid fauna is too large to treat as a whole, this digital identification tool is not designed to identify every tortricid encountered, but rather to reliably eliminate or confirm target taxa if or when they are encountered. This tool is available via the internet [http://idtools.org/id/leps/tortai/] and on CD.

Easily Confused Pests E. postvittana may be confused with many native tortricids. It can also be confused with Amorbia emigratella (Mexican leafroller), which has been reported from the United States, however, E. postvittana has ocelli which are lacking in A. emigratella. The undersides of E. postvittana hindwings are conspicuously immaculate as in A. emigratella, and the second abdominal tergite lacks the conspicuous median pit near the base which is present in A. emigratella (USDA, 1984).

References Armstrong, K.F., Cameron, C.M., Frampton, E.R., and Suckling, D.M. 1997 Aliens at the border and cadavers in the field: A molecular technique for species identification, pp. 316-321, Proceedings of the 50th New Zealand Plant Protection Conference. New Zealand Plant Protection Society, Rotorua, New Zealand.

Bailey, P., Catsipordas, A., Baker, G., and Lynn, B. 1995. Traps in monitoring light brown apple moth. The Australian Grapegrower and Winemaker: 130-132.

55 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Bailey, P. 1997. Light brown apple moth [Epiphyas postvittana] control options for the 1997/8 season. Australian & New Zealand Wine Industry Journal 12(3):267-268, 270.

Bellas, T.E., Bartell, R.J., Hill, A., 1983. Identification of two components of the sex pheromone of the moth Epiphyas postvittana (Lepidoptera, Tortricidae). Journal of Chemical Ecology 9(4): 503-512.

Bradley, J.D. 1973. Epiphyas postvittana (Walker), pp. 126-127, British Tortricid moths; Cochylidae and Tortricidae; Tortricinae. The Ray Society, London.

Bradley, S., Walker, J., Wearing, C., Shaw, P., and Hodson, A. 1998. The use of pheromone traps for leafroller action thresholds in pipfruit., pp. 173-178, Proceedings of the 51st New Zealand Plant Protection Conference. New Zealand Plant Protection Society, Rotorua, New Zealand.

Brown, J. W., Epstein, M.E., Gilligan, T.M., Passoa, S.C., and Powell, J.A. 2010. Biology, identification, and history of the light brown apple moth, Epiphyas postvittana (Walker) (Lepidoptera: Tortricidae: Archipini) in California: An example of the importance of local faunal surveys to document the establishment of exotic insects. American Entomologist 56: 26-35.

Buchanan, G. 1977. The seasonal abundance and control of light brown apple moth, Epiphyas postvittana (Walker) (Lepidoptera: Tortricidae), on grapevines in Victoria. Australian Journal of Agricultural Research 28: 125-132.

Buchanan, G.A., Stirrat, S.C., and Madge. D.G. 1991. Integrated control of light brown apple moth, Epiphyas postvittana (Walker), in vineyards. Wine Industry Journal 6: 220-222.

CABI. 2012. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Danthanarayana, W. 1975. The bionomics, distribution and host range of the light brown apple moth, Epiphyas postvittana (Walk.) (Tortricidae). Australian Journal of Zoology 23(3): 419-437.

Danthanarayana, W., Gu, H., Ashly, S. 1995. Population growth potential of Epiphyas postvittana, the lightbrown apple moth (Lepidoptera: Tortricidae) in relation to diet, temperature, and climate. Australian Journal of Zoology 43(4): 381-394.

Evans, J.W. 1937. The light-brown apple moth (Tortrix postvittana, Walk).The Tasmanian Journal of Agriculture 8: 1-18.

Foster, S., and Howard, A. 1998. Influence of stimuli from Camellia japonica on ovipositional behavior of generalist herbivore Epiphyas postvittana. Journal of Chemical Ecology 24: 1251-1275.

Foster, S., and Muggleston, S. 1993. Effect of design of a sex-pheromone baited delta trap on behavior and catch of male Epiphyas postvittana (Walker). Journal of Chemical Ecology 19: 2617-2633.

Foster, S.P., Howard, A., and Ayers, R.H. 1995. Age-related changes in reproductive characters of four species of tortricid moths. New Zealand Journal of Zoology 22: 271-280.

Foster, S., Muggleston, S., and Ball, R. 1991. Behavioral responses of male Epiphyas postvittana (Walker) to sex pheromone-baited delta trap in a wind tunnel. Journal of Chemical Ecology 17: 1449- 1468.

Geier, P., and Briese, D. 1980. The light-brown apple moth, Epiphyas postvittana (Walker): 4. Studies on population dynamics and injuriousness to apples in the Australian Capital Territory. Australian Journal of Ecology 5: 63-93.

56 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Geier, P.W., and Briese, D.T. 1981. The light-brown apple moth, Epiphyas postvittana (Walker); a native leafroller fostered by European settlement. In: Kitching RL, Jones RE, eds. The Ecology of Pests. Some Australian Case Histories. Melbourne, Australia: CSIRO.

Geier, P.W., and Springett, B.P. 1976. Population characteristics of Australian leafrollers (Epiphyas spp., Lepidoptera) infesting orchards. Australian Journal of Ecology 1(3):129-144.

Gilligan, T. M. and Epstein, M.E. 2009. LBAM ID, Tools for diagnosing light brown apple moth and related western U. S. leafrollers (Tortricidae: Archipini). Colorado State University, California Department of Food and Agriculture, and Center for Plant Health Science and Technology, USDA, APHIS, PPQ.

Glenn, D., and Hoffmann, A. 1997. Developing a commercially viable system for biological control of light brown apple moth (Lepidoptera: Tortricidae) in grapes using endemic Trichogramma (Hymenoptera: Trichogrammatidae). Journal of Economic Entomology 90: 370-382.

Lo, P., Suckling, D., Bradley, S., Walker, J., Shaw, P., and Burnip, G. 2000. Factors affecting feeding site preferences of lightbrown apple moth, Epiphyas postvittana (Lepidoptera: Tortricidae) on apple trees in New Zealand. New Zealand Journal of Crop and Horticultural Science 28: 235-243.

MacLellan, C. 1973. Natural enemies of the light brown apple moth, Epiphyas postvittana, in the Australian Capital Territory. Canadian Entomologist 105: 681-700.

Madge, D. G., and Stirrat. S.C. 1991. Development of a day-degree model to predict generation events for lightbrown apple moth Epiphyas postvittana (Walker) (Lepidoptera: Tortricidae) on grapevines in Australia. Plant Protection Quarterly 6: 39-42.

Magarey, P.A., Nicholas, P.R., and Wachtel, M.F. 1994. Control of the diseases of grapes in Australia and New Zealand. Wine Industry Journal 9: 197-225.

Meyrick, E. 1937. Tortrix postvittana Walk. (Microlepidoptera), a species new to Britain, Entomologist 70: 256.

Nuttal, M. 1983. Planotortrix excessana (Walker), Planotortrix notophaea (Turner), Epiphyas postvittana (Walker). Forest and Timber Insects of New Zealand no. 58.

Passoa, S., M. Epstein, T. Gilligan, J. Brambila, M. O'Donnell. n.d. Light Brown Apple Moth (LBAM) Epiphyas postvittana (Walker) Screening and Identification Aid.

Peterson, A. 1965. Some eggs of moths among the Olethreutidae and Tortricidae (Lepidoptera). Fla. Entomol. 48: 1-8.

Powell, J.A., and Common, I.F.B. 1985. Oviposition patterns and egg characteristics of Australian tortricine moths (Lepidoptera: Tortricidae). Aust. J. Zool. 179-216.

Suckling, D. M. and Brockerhoff, E.G. 2010. Invasion biology, ecology, and management of the light brown apple moth (Tortricidae). Annual Review of Entomology 55: 285-306.

Suckling, D., Brunner, J., Burnip, G., and Walker, J. 1994. Dispersal of Epiphyas postvittana (Walker) and Planotortrix octo Dugdale (Lepidoptera: Tortricidae) at a Canterbury, New Zealand orchard. New Zealand Journal of Crop and Horticultural Science 22: 225-234.

Svensson, I. 2009. Anmarkningsvarda fynd av smafjarilar (Microlepidoptera) I Sverige 2008 (Noteworthy finds of Microlepidoptera in Sweden 2008). Entomol. Tidskr. 130: 61-72.

57 Epiphyas postvittana Primary Pest of Grape Arthropods Light brown apple moth Moth

Terauds, A. 1977. Two methods of assessing damage to apples caused by light brown apple moth, Epiphyas postvittana (Walker). Journal of Australian Entomological Society 16: 367-369.

Thomas, W.P. 1975. Lightbrown apple moth, Life Cycle Chart. Auckland, New Zealand: Department of Scientific and Industrial Research.

Thomas, W.P. 1989. Epiphyas postvittana (Walker), lightbrown apple moth (Lepidoptera: Tortricidae). In: Cameron PJ, Hill RL, Bain J, Thomas WP, eds. A Review of Biological Control Invertebrate Pests and Weeds in New Zealand 1874 to 1987 Technical Communication. Wallingford, UK: CAB International, 187- 195.

Thwaite, W. 1976. Effect of reduced dosage of azinphos-methyl on control of codling moth, Cydia pomonella (L.) and light-brown apple moth, Epiphyas postvittana (Walk.),in an apple orchard. Zeitschrift für Angewandte Entomologie 80: 94-102.

USDA. 1984. Pests not known to occur in the United States or of limited distribution No. 50: Light-brown apple moth, pp. 1-12. APHIS-PPQ, Hyattsville, MD.

Venette, R.C., Davis, E.E., DaCosta, M., Heisler, H., and Larson, M. 2003, Mini Risk Assessment Light brown apple moth, Epiphyas postvittana (Walker) [Lepidoptera: Tortricidae]. 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/epostvittanapra.pd f.

Wearing, C.H., Thomas, W.P., Dugdale, J.W., and Danthanarayana, W. 1991. Tortricid pests of pome and stone fruits, Australian and New Zealand species. In: L.P.S. van der Geest and H.H. Evenhius (eds.) Tortricid pests: their biology, natural enemies, and control. World Crop Pests, Vol. 5. Elsevier, Amsterdam. Pp. 453-472.

Wolschrijn, J.B., and Kuchlein, J.H. 2006. Epiphyas postvittana, nieuw voor Nedlerland en het Europese continent (new to the Netherlands and the European continent) (Lepidoptera: Tortricidae). Tinea Ned. 1(5-6): 37-39.

Zimmerman, E.C. 1978. Insects of Hawaii: Microlepidoptera. University Press.

58 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Eupoecilia ambiguella

Scientific Name Eupoecilia ambiguella (Hübner)

Synonyms: Clysia ambiguella, Clysiana ambiguella, Cochylis ambiguella, Conchylis ambiguella, and Tinea ambiguella.

Common Names European grape berry moth, grape berry moth, grape bud moth, vine moth, grape moth

Type of Pest Moth

Taxonomic Position Class: Insecta, Order: Lepidoptera, Family: Tortricidae Figure 1. Top: E. ambiguella egg. Bottom: E. ambiguella pupa. Photos Reason for Inclusion in Manual courtesy of Rémi Coutin, OPIE Added per suggestion of the CPHST Otis laboratory

Pest Description Eggs: Eggs are 0.8 x 0.6 mm (0.03 x 0.023 in.); grayish brown when laid (Fig. 1), later becoming speckled with orange (Alford, 2007).

Larvae: Larvae (Fig. 3) are up to 12 mm (0.47 in.) long; body purplish brown or yellowish brown to olive green, with large, brown and moderately conspicuous pinacula; head and prothoracic plate dark brown or black; anal plate brownish to yellowish; abdominal prologs each with 25 to 30 crochets” (Alford, 2007).

Pupae: Pupa are 5 to 8 mm long (0.2 to 0.31 in.); reddish brown (Fig. 1); cremaster with a ring of 16 hook-tipped bristles and a pair of short horns dorsally (Alford, 2007).

Adults: Adults (Fig. 2) have a 12 to 15 mm (0.47 to 0.59 in.) wingspan; forewings whitish ochreous, marked with yellow ochreous and with a dark, brownish to black median fascia; hindwings gray (Alford, 2007).

59 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Biology and Ecology Lobesia botrana, European grapevine moth, and E. ambiguella share similar native ranges. In some years, both pests are present, while in others, one is more dominant (EPPO, 2002). E. ambiguella is found in more Northern, humid climates because of its preference for cooler weather (Arn et al., 1986). As such, it is more of a pest in northern countries (EPPO, 2002). E. ambiguella Figure 2. E. ambiguella adult. Photo courtesy of usually has two generations V. V. Neimorovets. per year, but a third generation can sometimes be observed (central Asia) (AgroAtlas, n.d.; Schmidt et al., 2003). Flight of the overwintering generation occurs soon after bud-break (EPPO, 2002).

When females are ready to mate, they stay in a “calling position” where pheromone is released (Schmieder-Wenzel and Schruft, 1990). The male uses this to orient himself towards the female over long range (Schmieder-Wenzel and Schruft, 1990). Once the male finds the female, visual cues are used to successfully complete the mating process (Schmieder-Wenzel and Schruft, 1990). Findings by Schmidt-Büsser et al. (2009) suggests that host plant chemostimuli along with sex pheromone help males find females on host plants

Overwintered females usually begin egg laying on the grape inflorescence and can lay up to 100 eggs (AgroAtlas, n.d.). Egg development at 15ºC (59ºF) takes 13 days; while development at 19 to 25ºC (66 to 77ºF) takes six-to-seven days (AgroAtlas, n.d.). After hatching, the first generation larvae penetrate single flower buds but will later tie several flower buds together with a silken web where they continue to feed; at this point tolerance is high and depends on grape variety (Ibrahim, 2004). Lab and field experiments have shown that larvae feed in the evening and early morning with mating occurring after midnight to early morning; eggs are laid during the afternoon and evening (Ibrahim, 2004). In the spring, eggs are laid singly on clusters while they are laid on berries during the summer (later generations) (Ibrahim, 2004). First generation larvae development lasts 15 to 25 days (AgroAtlas, n.d.). Schmidt et al. (2003) found through modeling that the population density and longevity of larvae is due to temperature and relative humidity. Development is most favorable at 70 to 90% relative humidity and temperatures of 18 to 25ºC (64 to 77ºF); the lower threshold of development is approximately 7ºC (45ºF) (AgroAtlas, n.d.). Second generation flight occurs 2 to 2.5 months after the first generation (HYPP Zoology, n.d.). Pupation occurs on the edge of leaves or trunks and is the overwintering stage (Ibrahim, 2004).

60 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Symptoms/Signs Larvae chew round holes in the berries eating the pulp and unripe seeds (AgroAtlas, n.d.). On average, larva can damage nine to 12 berries each (AgroAtlas, n.d.). As larvae grow, they begin to spin webs around surrounding flower buds or berries (depending on the generation) where they stay and continue to feed until pupation.

Larval damage is similar to damage caused by L. botrana. Flower buds, open flowers, and young fruitlets can be destroyed (Fig. 3), as well as developing or mature grapes later on in the season (Alford, 2007). Both the silken larval habitations and sticky exudations serve as contaminants in grape clusters (Alford, 2007).

Larvae damage can lead to infection by Botrytis cinerea, which causes a destructive gray mold rot on ripening berries and can lead to additional problems and crop losses (EPPO, 2002). Figure 3. E. ambiguella Pest Importance larva and damage. Photo This pest causes severe problems in northern courtesy of Rémi Coutin, European wine production areas (Schmidt et al., OPIE. 2003) and southern Germany (Kast, 2001). Both E. ambiguella and Lobesia botrana are considered the most important insect pests of European vineyards (Carde and Minks, 1995).

The presence of larvae, webs, and rotten fruit downgrades grape crops (HYPP Zoology, n.d.). Inflorescences will be destroyed by three or more webs made by larvae (AgroAtlas, n.d.). Although this pest can cause extensive damage, attacks earlier in the season are not usually as damaging as later season outbreaks (Alford, 2007).

Although damage is caused by destruction of fruit, a majority of damage is caused by the fungus Botrytis cinera, which develops after plant injury caused by the larvae of E. ambiguella (Meijerman and Ulenberg, 2000). This fungus can destroy entire grape clusters, making action thresholds low (Carde and Minks, 1995). Wine making becomes difficult when molds are present (HYPP zoology, n.d.).

Known Hosts The main host of this pest is the Vitis genus specifically Vitis vinifera (cultivated grapes), but larvae are considered polyphagous (AgroAtlas, n.d.).

Major hosts: Prunus domestica (plum), Ribes nigrum (blackcurrant), Vitis spp. (grapes), Vitis vinifera (grapes) (CABI, 2007).

61 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Minor hosts: Prunus salicina (Japanese plum) (CABI, 2007).

Reported as hosts: Acer spp. (maple), Acer campestre (hedge, field maple), Ampelopsis spp., Berberis aquifolium (hollyleaved barberry), Cissus quadrangularis (veldt-grape), Citrus limon (lemon), Clematis vitalba (old man's beard), Cornus spp. (dogwood), Cornus alba (redosier dogwood),Cornus mas (Cornelian cherry), Cornus sanguinea (bloodtwig dogwood), Cuscuta spp. (dodder), Daphne gnidium (flax-leaved daphne), Eleutherococcus spp. (aralia), Euonymus europaea (European spindle tree), Frangula spp. (buckthorn), Frangula alnus (alder buckthorn), Fraxinus spp.(ash), Galium spp. (yellow bedstraw), Galium mollugo (false baby’s breath), Hedera spp. (ivy), Hedera helix (English ivy), Ligustrum spp. (privet), Lonicera spp. (honeysuckle), Medicago sativa (alfalfa), Parthenocissus quinquefolia (Virginia creeper), Prunus spp. (stone fruit), Prunus spinosa (blackthorn), Pyracantha coccinea, Rhamnus spp. (buckthorn), Rhamnus cathartica (European buckthorn), Rhus glabra (sumac), Ribes spp. (currant), Rubus fruticosus (blackberry), Rubus ideaus (red raspberry), Sambucus racemosa (red elderberry), Silene uniflora subsp. prostrata(sculpit), Sorbus aucuparia (European mountain-ash), Symphoricarpos spp. (schisandra), Symphoricarpos albus (snowberry), Symphoricarpos rivularis, Syringa persica (Persian lilac), Syringa spp. (lilac), Syringa vulgaris (lilac), Trifolium pratense (red clover), Viburnum lantana (wayfaring tree), and Viburnum spp. (UK moths, n.d ; Whittle, 1986; Meijerman and Ulenberg, 2000; Ibrahim, 2004; Alford, 2007; Robinson et al., 2011).

Known vectors (or associated organisms) The entry holes made by E. ambiguella in grapes help facilitate the establishment of the fungus, Botryotinia fuckeliana (anamorph, Botrytis cinerea) which is the causal agent of gray mold (Bournier, 1976; EPPO, 2002). This pathogen is favored during rainy conditions and occurs from flowering time onward (EPPO, 2002).

Known Distribution This pest is widely distributed in central and southern Europe. It has a less southerly distribution than Lobesia botrana (European grapevine moth).

Asia: Armenia, Azerbaijan, China, Republic of Georgia, Japan, Kazakhstan, Korea, Kyrgyzstan, Pakistan, Taiwan, and Uzbekistan. Europe: Albania, Austria, Belgium, Bulgaria, Czech Republic, Denmark, Estonia, Finland, France (including Corsica), Germany, Greece*, Hungary, Italy (including Sardinia and Sicily), Latvia, Lithuania, Luxembourg, Macedonia, Moldova, the Netherlands, Norway, Poland, Portugal*, Romania, Russia, Serbia and Montenegro, Slovakia, Slovenia, Spain, Sweden, Switzerland, Ukraine, and United Kingdom.

This species has previously been reported from Brazil, but this record could not be verified (CABI, 1986).

* Aarvik (2011) states that E. ambiguella is absent from these countries.

62 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Potential Distribution within the United States This pest is found throughout Europe and parts of Asia. Due to its large range in similar climates, this pest may be able to establish throughout a large part of the United States. Because E. ambiguella is considered polyphagous (AgroAtlas, n.d.), establishment is not dependent on grape distribution.

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination.

Any of the following Trap Product Names in the IPHIS Survey Supply Ordering System may be used for this target: 1) Wing Trap Kit, Paper 2) Wing Trap Kit, Plastic

The Lure Product Name is “Eupoecilia ambiguella Lure.” The lure is effective for 42 days (6 weeks).

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

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

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: In Europe, pheromone traps are used to monitor the flight period although correlation between trapped insects and real populations is not strong (EPPO, 2002). Arn et al. (1986) found that the best attractant was a blend of Z-9- dodecenyl acetate, dodecyl acetate and octadecyl actetate with the last two serving as synergists to the main compound.

Using pheromones for mating disruption was found to be more expensive, but also more effective than using insecticides (Kast, 2001). Many regions recommend this technique for control to prevent undesirable effects on beneficial that insecticides would cause (EPPO, 2002). Several companies offer pheromone dispensers for mating disruption purposes. According to Carde and Minks (1995), ampullae dispensers are used in parts of Germany for pheromone releases. Some dispensers can be used for both L. botrana and E. ambiguella.

Population thresholds can be determined by monitoring symptoms (i.e., webbing tying flower buds together) of the first generation in order to correctly time treatments (EPPO, 2002).

63 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of E. ambiguella is by morphological identification. Identification requires dissection of the adult male genitalia; use Brown (2009) as an aid.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: A morphological identification is required for E. ambiguella. Key characteristics are provided below.

Male external characteristics: Labial palpi ochreous-cream, head and thorax cream, abdomen brownish. Antenna shortly ciliate. Forewing only slightly dilated posteriorly; ground color pale ochreous-white, extensively suffused and strigulated with yellow- ochreous, costa and dorsum strigulated or dotted with black, costa narrowly suffused with fuscous-black from base to beyond middle (outer margin of median fascia); markings dark gray; basal and sub-basal fasciae obsolescent, indicated only by costal striae; median fascia conspicuous, inwardly oblique, dilated on costa, diffusely strigulated or mixed with blackish, a variable ferruginous admixture above and below median fold, sometimes extending to dorsum; pre-apical spot obsolescent, indicated by weak costal striae; apex variably suffused with blackish; a small diffuse blackish spot in terminal margin near middle; cilia pale ochreous, a dark sub-basal line. Hindwing gray, paler basally; cilia light gray, with a dark sub-basal line (Razowski, 1970a; Bradley et al., 1973; Meijerman and Ulenberg, 2000).

Male genitalia: Socii expanding towards base such that bases touch each other; thin narrow terminals of socii situated laterally on broad bases. Vinculum divided ventrally, forming two sclerites. Caudal margin of valva oblique; sacculus broad at base, narrow beyond base, with short spinules on raised terminal portion; transtilla with strong median process. Aedeagus large, with one large thin cornutus and many small cornuti, arranged in three groups: 2 smaller groups and one in which the cornuti are arranged in a ring. In the latter group, the cornuti have plateshaped bases (Meijerman and Ulenberg, 2000).

Female external characteristics: Forewing markings similar to those of male; hindwing more uniformly gray (Meijerman and Ulenberg, 2000).

Female genitalia: Antrum very short, broad, connected to sterigma. Ductus bursae short, distal part membranous with many small denticles, anterior part sclerotized, reaching into the corpus bursae, covered with denticles. Anterior part of corpus bursae membranous (Meijerman and Ulenberg, 2000).

A new identification tool, Tort AI – Tortricids of Agricultural Importance, is available at http://idtools.org/id/leps/tortai/ from CPHST’s Identification Technology Program. This tool contains larval and adult keys, fact sheets, an image gallery, molecular search capacity, and more. Eupoecilia ambiguella is included in this tool.

64 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Easily Confused Pests This pest can be confused with Lobesia botrana (European grapevine moth) (Vasquez, 2009). L. botrana is present in California. Both have similar biologies, damage, and synchronous flight periods (Bournier, 1978). E. ambiguella can also be confused with Endopiza viteana (present in the United States).

References AgroAtlas. n.d. Pests: Eupoecilia ambiguella (Hubner) - European Grape Berry Moth, Grape Berry Moth, Grape Bud Moth, Vine Moth, Grape Moth. Interactive Agricultural Ecological Atlas of Russia and Neighboring Countries. Economic Plants and their Diseases, Pests and Weeds.

Alford, D.V. 2007. Pests of Fruit Crops: A Color Handbook. Manson Publishing Ltd, London. pp. 461.

Arn, H., Rauscher, S., Buser, H., and Guerin, P.M. 1986. Sex pheromone of Eupoecilia ambiguella female: analysis and male response to ternary blend. Journal of Chemical Ecology 12(6): 1417-1429.

Aarvik, L. 2011. Fauna Europaea: Eupoecilia ambiguella. In O. Karsholt and E. J. Nieukerken (eds.) Fauna Europaea: Lepidoptera, Moths, version 2.4. Accessed April 10, 2012 from: http://www.faunaeur.org.

Brown, J. 2009. Adult Lepidoptera Workshop. http://caps.ceris.purdue.edu/webfm_send/107.

Bournier, A. 1978. Grape insects. Annual Review of Entomology 22: 355-376.

CABI. 1986. Eupoecilia ambiguella (Huebner). CAB International Institute of Entomology Distribution Maps of Pests. Series A (Agricultural) Map no. 76.

Carde, R.T. and Minks, A.K. 1995. Control of moth pests by mating disruption: successes and constraints. Annual Review of Entomology 40: 559-585.

EPPO. 2002. EPPO standards: good plant protection practice, grapevine. Bulletin OEPP/EPPO Bulletin 32: 371-392

HYPPZ. n.d. Eupoecilia ambiguella Hubner. Encyclopédie des ravageurs européens: HYPPZ. http://www.inra.fr/hyppz/RAVAGEUR/6eupamb.htm. Accessed on: February 1, 2010.

Ibrahim, R.A.E.A. 2004. Biological control of grape berry moths Eupoecilia ambiguella Hb. and Lobesia botrana Schiff. (Lepidoptera: Tortricidae) by using egg parasitoids of the genus Trichogramma. Doctorate thesis, Justus Liebig University of Giessen, Germany. pp. 103.

Kast, W.K. 2001. Twelve years of practical experience using mating disruption against Eupoecilia ambiguella and Lobesia botrana in vineyards of the Wuerttemberg region, Germany. Pheromones for Insect Control in Orchards and Vineyards, IOBC wprs Bulletin 24(2): 71-73.

Meijerman, L., and Ulenberg, S.A. 2000. Arthropods of Economic Importance: Eurasian Tortricidae.

Robinson, G.S., Ackery, P.R., Kitching, I.J., Beccaloni, G.W., and Hernández, L.M. 2011. HOSTS - a Database of the World's Lepidopteran Hostplants: Eupoecilia ambiguella. London: Natural History Museum. Accessed April 10, 2012 from: http://www.nhm.ac.uk/research- curation/research/projects/hostplants/search/index.dsml.

Schmidt, K., Hoppmann, D., Holst, H. and Berkelmann-Löhnertz, B. 2003. Identifying weather-related covariates controlling grape berry moth dynamics. Bulletin OEPP/EPPO Bulletin 33: 517-524.

65 Eupoecilia ambiguella Primary Pest of Grape Arthropods European grape berry moth Moth

Schmidt-Büsser, D., von Arx, M. and Guerin, P.M. 2009. Host plant volatiles serve to increase the response of male European grape berry moths, Eupoecilia ambiguella, to their sex pheromone [Abstract]. Journal of Comparative Physiology A Neuroethology, Sensory, Neural, and Behavioral Physiology 195(9): 853-864.

Schmieder-Wenzel, C. and Schruft, G. 1990. Courtship behavior of the Eupoecilia ambiguella Hb. (Lep., Tortricidae) in regard to pheromonal and tactile stimuli. Journal of Applied Entomology 109: 341-346.

UK Moths. n.d. Catalogue of the Lepidoptera of Belgium: Eupoecilia ambiguella (Hübner, 1796). http://webh01.ua.ac.be/vve/Checklists/Lepidoptera/Tortricidae/Eambiguella.htm. Accessed on: February 2, 2010.

Vasquez, S.J. 2009. New invasive vineyard pest found in California. Raisin Ramblings, Agriculture and Natural Resources, University of California. http://ucanr.org/blogs/blogcore/postdetail.cfm?postnum=1943. Accessed on: February 1, 2011.

Whittle, K. 1986. Pests not known to occur in the United States or of limited distribution. No. 73: European grape berry moth. United States Department of Agriculture, Animal and Plant Health Inspection Service, Plant Protection and Quarantine. 11 pp.

66 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

Heteronychus arator

Scientific Name Heteronychus arator (Fabricius)

Synonyms: Heteronychus sanctaehelenae, Heteronychus transvaalensis, Scarabaeus arator

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

Type of Pest Beetle

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

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 (0.07 in.) 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 1 to 5 cm (0.39 to 1.97 in.). Females each lay Figure 1. Illustration of each stage of between 12 to 20 eggs total. In the field, eggs the life cycle of the black maize hatch after approximately 20 days. Larvae can beetle, showing a close up view of be seen clearly with the naked eye each stage and a background view (Matthiessen and Learmoth, 2005; CABI, showing that the eggs, larvae, and 2007). pupae are all underground stages with the adult beetles as the only Larvae: There are three larval instars. Larvae stage appearing above ground. are creamy-white except for the brown head Illustration courtesy of NSW capsule and hind segments, which appear Agriculture. dark where the contents of the gut show through the body wall. The head capsule is smooth textured, measuring 1.5 mm, 2.4 mm, and 4.0 mm (0.059, 0.094, and 0.16 in.) at each respective instar. The third-instar larva is approximately 25 mm (0.98 in.) long when fully developed. Black maize beetle larvae are soil-dwelling and resemble white 'curl grubs.’ They have three pairs of legs on the thorax, a prominent brown head with

67 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

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 Learmoth, 2005; CABI, 2007).

Figure 2. Eggs, larvae, and adult black maize beetle. Photo courtesy of Yates Ltd.

Pupae: The larvae, when fully grown, enter a short-lived pupal stage, which measures approximately 15 mm (0.59 in.) long and is typically coleopteran in form (cylindrical shape), initially pale yellow, but becoming reddish-brown nearer to the time of emergence (Matthiessen and Learmoth, 2005).

Adults: Beetles are 12 to 15 mm (0.47 to 0.59 in.) 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 beetle is the main pest stage (Matthiessen and Learmoth, 2005; CABI, 2007).

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

68 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

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 (5.9 in.). They are a shiny black and cylindrical cockchafer that is slow moving and is approximately 15 mm (0.59 in.) long. The adult is capable of flying, which serves to disperse the beetle to new sites (Matthiessen and Learmoth, 2005). 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 Learmoth, 2005).

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 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. Black maize 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 (Matthiessen and Learmoth, 2005).

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, 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 Learmoth, 2005).

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., 1981). 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 black maize 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

69 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

training. Partial damage to vines by black maize 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 Learmoth, 2005; CABI, 2007).

Known Hosts Major hosts Cynodon dactylon (Bermuda grass), Dactylis glomerata (cocksfoot grass), Digitaria sanguinalis (hairy crab grass), Echinochloa crus-galli (barnyard grass), Echinochloa frumentacea (Japanese barnyard millet), Eucalyptus spp. (Eucalyptus tree), Eragrostis tef (tef lovegrass), Festuca arundinacea (tall fescue), Hordeum vulgare (barley), Lolium spp. (ryegrass), Lolium x hybridum (hybrid ryegrass), Lolium perenne (perennial ryegrass), Lolium rigidum (annual ryegrass), Paspalum dilatatum (dallisgrass), Solanum tuberosum (potato), Vitis vinifera (grape), and Zea mays (corn) (King and Watson, 1982; CABI, 2007; Maddison and Crosby, 2009).

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), Nicotiana tabacum (tobacco), Olea spp. (olives), Oryza spp. (rice), 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), Saccharum officinarum (sugarcane), Secale strictum subsp. strictum (perennial rye), Solanum lycopersicum (tomato), Sorghum spp. (sorghum), and Triticum aestivum (wheat) (Litsinger et al., 1986; CABI, 2007; Gordh and Headrick, 2011.).

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 of the Congo, Ethiopia, Kenya, Lesotho, Madagascar, Malawi, Mozambique, Namibia, Saint Helena, South Africa, Tanzania (including Zanzibar), Zaire, Zambia, and Zimbabwe; Oceania: Australia, New Zealand, and Norfolk Island (CABI, 2007).

There are reports of this species in Brazil and Papua New Guinea, but these are considered unreliable records (EPPO, 2012).

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

70 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

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 southern United States, Arkansas, California, Florida, Georgia, Louisiana, Mississippi, North Carolina, Oklahoma, South Carolina, and Texas, have the greatest risk for H. arator establishment based on host availability and climate within the continental United States. Establishment is precluded from most of the northern United States.

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

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Visual survey: Areas that are rotated with or replace pasture lands are most at risk of damage from the black maize beetle. Most damage by the black maize 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. Black maize beetles eat the cutting and rootlings at or just below ground level, ring bark the vine, and cause wilting and collapse. Inspect immediately below the soil surface for signs of H. arator attack, in particular frayed chewing around the stem circumference.

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 (19.69 in.) long portions of hilled-up rows comprised the sample unit. A 70 x 30 cm (27.56 x 11.81 in.) piece of sheet steel is pressed into soil across the row with soil on one side of the metal sheet being Figure 3. Example of a excavated. The steel sheet is then removed to expose an pitfall trap. Photo courtesy undisturbed soil face of the potato hill. Presence of the of Mnolf. beetle or other pests and associated plant damage in the http://commons.wikimedia. top, center, or bottom of soil cross sections is recorded. org/wiki/File:Barber_pitfall_ Fifty samples were examined at each sampling time in a trap.jpg. uniform grid across a 0.2 ha (0.49 acre) crop area.

71 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

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 (3.94 x 5.9 in.) 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).

Trapping: Matthiessen and Learmonth (1998) used pitfall (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 (23.6 in.)-long 20 watt fluorescent black light, the center of which was 1.5 meters (4.92 ft.) above the ground. Four vertically-oriented 17.5 cm (6.89 in.) 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 (8.27 in.) 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 (8.27 in.) 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 (32.8 ft.) intervals at one location. Subsequent traps were made from a 10 cm (3.94 in.) 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 (16.4 ft.) intervals in each of two lines 10 meters (32.8 ft.) 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.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Black maize 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

72 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

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://padil.gov.au/pests-and- diseases/Pest/Main/135883.

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 (0.98 in.) 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 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 black maize beetle and are much shorter (only up to 3 to 4 mm (0.12 to 0.16 in.)). 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.

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

Gordh, G. and Headrick, D.A. 2011. Dictionary of Entomology. 2nd Edition. CAB International. 1526 pp.

King, P.D. and Watson, R.N. 1982. Prediction and monitoring of black beetle, Heteronychus arator (Coleoptera: Scarabaeidae), outbreaks in New Zealand. New Zealand Entomologist 7(3): 227-231.

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

Litsinger, J.A., Barrion, A.T., and Soekarna, D. 1986. Upland rice insect pests. Their ecology, importance, and control. In Progress in Upland Rice Research: Proceedings of the 1985 Jakarta Conference. The International Rice Research Institute. pp. 403-445.

73 Heteronychus arator Primary Pest of Grape Arthropods Black maize beetle Beetle

Maddison, P. A. and Crosby, T.K. 2009. Summary of plant-animal associations from “Maddison (1993) Pests and other fauna associated with plants, with botanical accounts of plants. Technical report. UNDP/FAO-SPEC Survey of Agricultural Pests and Diseases in the South Pacific, vol. 3. Auckland : Manaaki Whenua B Landcare Research.” Accessed April 11, 2012 from: http://nzac.landcareresearch.co.nz/.

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 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.

Matthiessen, J.N., and Ridsdill-Smith, T.J. 1991. Populations of African black beetle, Heteronychus arator Coleoptera: Scarabaeidae) in a Mediterranean-climate area of Australia. Bulletin of Entomological Research 81: 85-91.

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.

74 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Lobesia botrana

Scientific Name Lobesia botrana [Denis & Schiffermüller]

Synonyms: Cochylis vitisana, Cochylis botrana, Coccyx botrana, Eudemis botrana, Eudemis rosmarinana, Grapholita botrana, Lobesia rosmariana, Noctua romani, Paralobesia botrana, Penthina vitivorana, Polychrosis botrana, Tortrix botrana, Tortrix vitisana, Tinea premixtana, Tinea reliquana, Tortrix reliquana, and Tortrix romaniana.

Common Names European grapevine moth, grape fruit moth, grape leaf-roller, grape vine moth, grape moth, vine moth

Type of Pest Moth

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

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2003 through 2009, Program pest

Pest Description European grapevine moth (EGVM) is primarily a pest on the flowers and fruits of grape vines, but the moth has been known to infest stone fruit trees, privet and olives as well. Survey for this pest in stone fruit, privet and olives is important because, in general, these secondary hosts flower before grapes and L. botrana can be found on these earlier hosts before moving over to grapes, its preferred host.

Eggs: The egg of L. botrana is of the so-called ‘flat type’ with the long axis horizontal and the micropyle at one end. Eggs are elliptical, flattened, and slightly convex with a mean eccentricity of 0.65. The egg measures about 0.65 to 0.90 mm (0.03 to 0.04 in.) x 0.45 to 0.75 mm (0.02 to 0.03 in.). Freshly laid eggs are pale yellow, later becoming light gray and translucent with iridescent glints (opalescent). The chorion is macroscopically smooth but presents a slight polygonal reticulation in the border and around the micropyle. The time elapsed since the eggs were laid may be estimated by observing the eggs: there are five phases of embryonic development - visible embryo, visible eyes, visible mandibles, brown head, and black head. As typically occurs in the subfamily Olethreutinae, eggs are laid singly, and more rarely in small clusters of two or three (CABI, 2009; Gilligan et al., 2011).

Larvae: There are usually five larval (Fig. 1A) instars. Neonate larvae are about 0.95 to 1 mm (~0.04 in.) long, with head and prothoracic shield deep brown, nearly black, and

75 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth body light yellow to yellowish green. Mature larvae reach a length between 10 and 15 mm (0.39 to 0.59 in.), with the head and prothoracic shield lighter than neonate larvae and the body color varying from light yellowish green to pale brown, depending principally on A larval nourishment (CABI, 2009; Gilligan et al., 2011). The head is brown to light yellowish brown to honey colored, the antennae and thoracic legs are brown to black, and the prothoracic shield is variably shaded with dark brown to black on the posterior and lateral margins. All instars have a dark stemmatal area and genal dash (Gilligan et al., 2011).

Important structural features of L. botrana larvae include: mandibles B without inner teeth or a retinaculum; distance between P1 and AF2 on head equal to distance between P1 and P2; a horizontal line connecting the P2 setae on head passes through AF2; L pinaculum on T1 horizontal, not extending beneath spracle; SV groups on A1, 2, 7, 8, 9 with 3:3:3:2:2 setae; distance between V setae on A9 approximately 1.5 to 2.0 times the distance between V setae on A8; distance between D1 setae on anal shield equal to distance between D1 and SD1; anal comb with 5 to 6 teeth in California individuals, other authors report 6 to 8 teeth; and body spicules relatively dense (Gilligan et al., 2011). C

Pupae: Female pupae are larger Figure 1. Larva (A), pupa (B), and adult (5 to 9 mm; 0.20 to 0.35 in.) than male (C) L. botrana. Photos courtesy of males (4 to 7 mm; 0.16 to 0.28 Instituto Agrario S. Michele All’ Adigen, in.). Freshly formed pupae are HYPPZ Zoology, and Todd Gilligan, usually cream or light brown but Colorado State University, respectively.

76 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

also light green or blue, but a few hours later become brown or deep brown (Fig. 1B). Cast pupal skins, are somewhat unusual in retaining a greenish tint on the anterior abdominal segments. Pupal age may be estimated as a function of tegument transparency and coloring (CABI, 2009). The sexes may be distinguished by the position of genital sketches that are placed in the IX and VIII abdominal sternites in males and females, respectively. Moreover, the male genital orifice is placed between two small lateral prominences. When Figure 2. Male genetalia of L. botrana. adult emergence is imminent, pupae Photo courtesy of Todd Gilligan, perforate the cocoon, resting the exuvia Colorado State University. fixed outwardly in a characteristic position by cremaster spines.

Important structural features of L. botrana pupae include: head unmodified, without projections; clypeus with two pairs of setae; A4 and A5 with 22 to 24 spines between the D2 setae; dorsum of A10 with a patch of spine and no setae present on the anal rice. The cremaster is fan-shaped with a weakly emarginated caudal margin (Gilligan et al., 2011).

Adults: Forewing length ranges from 4.5 to 8.5 mm (0.18 to 0.33 in.) (Gilligan et al., 2011). Adult size is greatly affected by larval food quality (Torres-Vila, 1995). Forewing pattern exhibits little variation and no sexual dimorphism. Forewing pattern is as follows; ground color cream; interfascial areas overlaid with leaden gray; costal stringulae cream, well defined; fasciae brown to dark brown; subbasal fasica well defined, with black scaling medially; median fasica well defined, with triangular medial projection often suffused with black scaling; postmedian fasica broken, forming pretormal patch along dorsum with cluster of black scales; postmedian band forming large brown patch along termen; apex often with conspicuous black dot; termin outlined in cream; fringe brown. The males (Fig. 1C) lack a forewing costal fold. The male hind wing is whitish with a brown periphery; while the female hind wing is completely brown (Gilligan et al., 2011). Figure 3. Female genetalia of L. Male genetalia (Fig. 2) can be distinguished by a combination botrana. Photo of the following characteristics: socii short, lateral, apex with courtesy of Todd numerous setae; uncus reduced to small bilobed hump on Gilligan, Colorado State University.

77 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

tegument; gnathos weakly sclerotized; valvae long and narrow with dense row of strong spines on ventral margin; cucullus densely setose, separated from sacculus by distinct gap in row of ventral spines; sacculus weakly concave post-medially; phallus small; cornuti absent. Female genetalia (Fig. 3) are characterized by a long, slender ductus bursa that is undifferentiated from the corpus bursae, gradually expanded anteriorly, and an unusual, elongate, somewhat feather-shaped signum (Gilligan et al., 2011).

Biology and Ecology: The first flight of adults occurs in spring when daily average air temperature is above the minimal threshold temperature of 10°C (50°F) for 10 to 13 days. The second flight period begins in summer (USDA, 1985). In Israel, adults appear in the vineyard when grapevines flower. Adults are hard to discover during the day and may be noticed only when they take flight after being disturbed. They fly at dusk whenever the temperature is above 12°C (54°F), but rainfall and wind will reduce A B flight. Adults usually prefer hot, dry places protected from wind so they fly mainly between the first rows of grapevines close to windbreaks and on slopes facing the sun (Avidov and Harper, 1969).

Within a day or two of mating, Figure 4. Adult on grape fruit (A) and larvae feeding females begin to oviposit on inside a grape (B). Photos courtesy of Michael Breuer. the blossoms, leaves, and http://www.bio- tender twigs of the grapevine. pro.de/de/region/freiburg/magazin/01476/index.html. The female lays 300 or more eggs singly or in groups of two or three at a rate of more than 35 per day. During rearing experiments under laboratory conditions in Czechoslovakia, the optimum temperatures for oviposition were from 20 to 27°C (68 to 81°F) (Gabel, 1981). First generation eggs are laid on the flower buds or pedicels of the vine while second generation eggs are laid on individual grapes (USDA, 1985) (Fig. 4A). Eggs hatch in 5 to 10 days or 75 degree- days above a 10°C (50°F) threshold (Gilligan et al., 2011).

The European grapevine moth is a polyvoltine species (CABI, 2009). The number of generations in a given area is fixed by photoperiod together with temperature, acting on diapause induction and development rate, respectively. Short-day photophases (between 8 and 12 h) during the larval stage induce diapause in larvae that will be later expressed in pupae. The moth achieves two generations in northern cold areas and usually three generations in southern temperate areas, although this general latitudinal pattern is often modified by the altitude-derived gradient and/or microclimatic conditions in a given area. Thus the number of generations has a broader range, reported as one generation in Romania (Filip, 1986) to four generations (often partial) in Spain, Greece,

78 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Crete, Italy, and former Yugoslavia (Coscollá, 1997 and references therein). Five generations have been reported in Turkmenistan (Rodionov, 1945).

First generation larvae feed on bud clusters or flowers and spin webbing around them (glomerules) before pupating inside the web or under the rolled leaf. Second generation larvae enter an unripened grape (Fig. 4B) and feed before pupating inside the grape. Larvae of the third generation, the most damaging, feed on ripening grapes, migrating from one to another and spinning webs. The third generation larvae leave the fruit and shelter Figure 5. Glomerules of L. botrana. Photo under the bark, among dead courtesy of EFAPO-ES. leaves, or between clods of earth, where they pupate before overwintering. Few of these larvae pupate before harvest, and many are gathered with the grapes. Larval development is completed in approximately 20 to 28 days or 170 degree days for larvae feeding on flowers and 225 degree-days for larvae feeding on berries (Gilligan et al., 2011). Pupae complete development in approximately 12 to 14 days, or 130 degree-days, for non-diapausing individuals (Gilligan et al., 2011).

Moth activity (i.e., flight, feeding, calling, mating, and egg-laying) is principally displayed at dusk, although some activity can also occur at daybreak or at any time on cloudy days. Water availability is necessary for adults to reach their potential reproductive output (Torres-Vila et al., 1996). Females are usually monandrous, but several physiological factors may enhance multiple mating (Torres-Vila et al., 1997). On the other hand, males are largely polygynic Figure 6. Damage by L. botrana. Photo (Torres-Vila et al., 1995). courtesy of HYPPZ Zoology.

Symptoms/Signs Documentation on damage to stone fruits is limited, as most information is available on grape. This moth can cause damage to the flowers and the fruit of stone fruit hosts.

79 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

On grape inflorescences, neonate (first generation) larvae firstly penetrate single flower buds. Symptoms are not evident initially because larvae remain protected by the top bud. Later, when larval size increases, each larva agglomerates several flower buds with silk threads forming glomerules visible to the naked eye (Fig. 5), and the larvae continue feeding while protected inside. Larvae usually make one to three glomerules during their development. Despite hygienic behavior of larvae, frass may remain adhering to the glomerules. On grapes (summer generations), larvae feed externally and when berries are a little desiccated (Fig. 6), they penetrate them, bore into the pulp, and remain protected by the berry peel (Fig. 4B, 7). Larvae secure the pierced berries to surrounding ones by silk threads in order to avoid falling. Each larva directly damages several berries (one to six), but if the conditions are suitable for fungal or acid rot Figure 7. Larva inside grape fruit. development, a large number of berries Photo courtesy P. del Estal (CABI, placed around may be also affected. 2009). Damage is variety-dependent; generally it is more severe on grapevine varieties with dense grapes because this increases both larval installation and rot development. On both inflorescences and grapes, several larvae may co-exist in a single reproductive organ. Larval damage on growing points, shoots, or leaves is unusual.

First-generation larval feeding on the buds or flowers webs them and prevents further growth. If heavy flower damage occurs during the first moth generation, the affected flowers will fail to develop and yield will be low. Damage by summer larvae of the second and third generation results in many nibbled berries, which later shrivel. The berries may be eaten either partly (leading to rot) or completely (leaving only empty skins at the tip of the bunch). Sometimes berries drop, and only the stalks remain (USDA, 1985).

Pest Importance The European grapevine moth is a serious pest in the warm vine-growing countries where it is normally found. Larvae feed on flower buds, developing berries, and most destructively, on the ripening fruit of grape. The primary damage to grape berries attracts other insects and predisposes the fruit to fungal infection. Larval boring in grapes may promote a number of fungal rots (CABI, 2009). Loss of up to one-third of the vintage has been reported in areas of the Soviet Union, Syria, and Yugoslavia. Losses in Israel sometimes reach 40 to 50 percent among table grapes and up to 80 percent or more for wine grapes. Further loss is due to the time and labor spent in cleaning the grape bunches. When infestations are heavy, the work days spent in cleaning the fruit account for 30 to 40 percent of the time of those involved in harvesting (USDA, 1985).

80 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

On grapes (summer generations), indirect damage is usually more important than direct, at least in the event of less severe attacks. Thus global damage may appear of little importance if it is evaluated exclusively as weight loss (direct damage) because greater damage is due to rot-derived reduction in quality (indirect damage). Larval boring in grapes may promote a number of fungal rots including Aspergillus, Alternaria, Rhizopus, Cladosporium, Penicillium, and especially Botrytis cinerea (Fig. 8) (Fermaud and Le Menn, 1989; CABI, 2009).

Figure 8. Discolored, shriveled berries caused by Botrytis bunch rot (left) and Botrytis cinerea sporulating on grape berries (right). Photos courtesy P. Sholberg, Agriculture & AgriFood Canada.

Known Hosts This pest feeds primarily on the flowers and fruits of grapes. However, L. botrana demonstrates a curious behavior of feeding on many different plant families (approximately 27), but only a few species within each family are suitable. Grape cultivars with prolonged blossoming or late-ripening berries are usually more heavily infested than short-flowering or early ripening varieties (Avidov and Harper, 1969). L. botrana exhibits an oviposition preference for privet and certain grape cultivars, such as ‘Cabernet Sauvignon’ (Maher et al., 2000, 2001).

Both plums and peaches/nectarines are as suitable as grapes for the development of L. botrana. These fruit trees bloom earlier than grapes, and since the female moths are active during this time, oviposition onto Prunus flowers and fruits has been observed and studied. The development of larvae and the reproductive capacity of females that are raised on the flowers and fruits of plums and peaches/nectarines are equal to the moths reared on the flowers and fruits of grapevines (Stavridis and Savopoulou- Soultani, 1998). Sour cherries, apricots and other stone fruits do not seem to support the same level of successful development of L. botrana, and can be seen as only minor hosts of this moth.

81 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Major hosts Prunus domestica (plum), Prunus persica (peach/nectarine), Vitis vinifera (grape), and Vitis spp.

Minor hosts Actinidia chinensis (kiwi), Clematis vitalba (traveler’s joy), Coffea spp. (coffee), Daphne odora (winter daphne), Dianthus spp. (carnation), Diospyros kaki (persimmon), Diospyros virginiana (persimmon), Hypericum calycinum (Aaron’s beard), Hordeum vulgare (barley), Medicago sativa (alfalfa), Olea europaea subsp. europaea (olive), Prunus spp., Prunus avium (sweet cherry), Prunus domestica (plum), Prunus dulcis (almond), Prunus spinosa (blackthorn), Punica granatum (pomegranate), Pyrus communis (pear), Ribes spp. (currant), Ribes nigrum (blackcurrant), Ribes rubrum (red currant), Ribes uva-crispa (gooseberry), Rosa spp. (rose), Rubus fruticosus (European blackberry), and Solanum tuberosum (potato).

Wild hosts Arbutus unedo (arbutus), Berberis spp. (barberry), Berberis aquifolium (Oregon-grape), Berberis vulgaris (European barberry), Clematis vitalba (old man's beard), Cornis spp. (dogwood), Cornus alba (dogwood), Cornus mas (cornelian cherry), Cornus sanguinea (dogwood), Cucumis sativus (cucumber), Daphne gnidium (spurgeflax daphne), Galium mollugo (smooth bedstraw), Hedera helix (ivy), Lamium amplexicaule (henbit), Ligustrum vulgare (privet), Lonicera tatarica (Tatarian honeysuckle), Lonicera xylosteum (fly honeysuckle), Menispermum canadense (common moonseed), Parthenocissus quinquefolia (Virginia creeper), Rhus glabra (smooth sumac), Rosmarinus officinalis (rosemary), Rubus spp. (blackberry), Rubus caesius (dewberry), Rubus dumetorum (European dewberry), Silene vulgaris (bladder campion), Syringa vulgaris (lilac), Tanacetum vulgare (tansy), Trifolium pratense (red clover), Viburnum lantana (wayfaring tree), and Ziziphus jujuba (common jujube) (Hoenisch, n.d.; USDA, 1985; Whittle, 1985).

Known Vectors (or associated organisms) It has been shown that the nutritional alteration of berries caused by Botrytis cinerea may enhance female fecundity of L. botrana (Savopoulou-Soultani and Tzanakakis, 1988). Larval boring in grapes may promote a number of fungal rots including Aspergillus, Alternaria, Rhizopus, Cladosporium, Penicillium, and especially Botrytis cinerea.

Known Distribution Africa: Algeria, Egypt, Eritrea, Ethiopia, Kenya, Libya, and Morocco; Asia: Armenia, Azerbaijan, Republic of Georgia, Iran, Iraq, Israel, Japan, Jordan, Kazakhstan, Lebanon, Syria, Tajikistan, Turkey, Turkmenistan, and Uzbekistan; Europe: Austria, Belgium, Bulgaria, Cyprus, Czech Republic, France (including Corsica), Republic of Georgia, Germany, Greece (including Crete), Hungary, Italy (including Sardinia and Sicily), Lithuania, Luxembourg, Macedonia, Malta, Moldova, Montenegro, Poland, Portugal, Romania, Russia, Serbia, Slovakia, Slovenia, Spain, Sweden, Switzerland,

82 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Ukraine, and the United Kingdom (CABI, 2007). South America: Argentina and Chile (CIE, 1974; SENASCA, 2010; Aarvik, 2011; EPPO, 2012).

In 2008, the first report of L. botrana in Western Hemisphere occurred in Chile (Gonzalez, 2008). In March 2010, the Argentinean National Service for Agrifood Health and Quality reported L. botrana in Argentina at locations in the Maipu Department, Mendoza Province, close to the Chilean border (SENASCA, 2010).

North American records from the mid- to late- 1800s are misidentifications of Paralobesia viteana (Kearfott, 1904), a native North American grape-feeding tortricid that is extremely similar in morphology to L. botrana (Gilligan et al., 2011).

Potential Distribution within the United States Climatic conditions in the major grape growing areas of the United States favor the establishment of L. botrana (USDA, 1985; Venette et al., 2003). Venette et al. (2003) estimated that approximately 29% of the continental United States may be suitable for L. botrana establishment. This projection includes the major California wine-producing counties of Napa, Sonoma, Amador, Monterey, and San Luis Obispo.

A recent risk analysis by USDA-APHIS-PPQ-CPHST indicates, however, that areas of the southeastern United States have the greatest risk for L. botrana establishment based on host availability and climate within the continental United States. L. botrana is excluded from establishment in areas of the western and northeastern United States.

On September 15, 2009, Lobesia botrana was detected in commercial vineyard in Napa County, California. Since 2010, , the moth has been detected and a quarantine is in place for portions of 10 counties (Fresno, Mendocino, Merced, Napa, Nevada, San Joaquin, Santa Clara, Santa Cruz, Solano, and Sonoma )in California. In March 2012, L. botrana was declared eradicated from Fresno, Mendocino, Merced, and San Joaquin counties leaving only, Napa, Nevada, Santa Clara, Santa Cruz, Solano, and Sonoma under quarantine.

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination. The approved trap is a delta trap.

Any of the following Trap Product Names in the IPHIS Survey Supply Ordering System may be used for this target, 1) Paper Delta Trap, 2 sticky sides, Brown 2) Paper Delta Trap, 2 sticky sides, Green 3) Paper Delta Trap, 2 sticky sides, Orange 4) Paper Delta Trap, 3 sticky sides, Orange 5) Large Plastic Delta Trap Kits, Red

The Lure Product Name is “Lobesia botrana Lure.” The lure is effective for 28 days (4 weeks).

83 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

Method Notes: The paper delta trap (with 2 sticky sides) has been added as an approved method. Both the large plastic delta trap (red) and the orange paper delta trap (with 3 sticky sides) are acceptable for use and for data reporting. For 2011 and the foreseeable future, the PPQ Lobesia Program has chosen the 2-sided paper delta trap as the preferred trap for the program. When using the 2-sided trap, the lure should be placed in a lure hanger inside the trap.

The trap color may be decided by the State and does not affect trap efficacy. For the paper delta traps, all of the standard colors used for gypsy moth (brown, green, or orange) are acceptable. Red was the recommended color for the large plastic delta trap as it has been shown to reduce trap catches of non-target (beneficial) insects. Trap color has not been shown to increase or decrease catches of L. botrana.

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

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: From Venette et al. (2003) Trapping: A sex pheromone has been identified that is highly attractive to males. Males are most attracted to a five component blend of (E,Z)-(7,9)-dodecadienyl acetate, (E,Z)- (7,9)-dodecadien-1-ol, (Z)-9-dodecenyl acetate, (E)-9-dodecenyl acetate and 11- dodecenyl acetate in a ratio of 10: 0.5: 0.1: 0.1: 1. Males are slightly less attracted to a three component blend of (E,Z)-(7,9)-dodecadienyl acetate, (E,Z)-(7,9)-dodecadien-1-ol, (Z)-9-dodecenyl acetate (ratio of 10:0.5:0.1). Males were still attracted, but much less so, to the main pheromone component (E,Z)-(7,9)-dodecadienyl acetate. The main pheromone component has been used to disrupt mating as a method of pest control and to monitor the flight period of males. However, this compound is sensitive to sunlight and degrades, becoming non-attractive to L. botrana after 60 minutes of exposure to UV radiation.

Pheromone-baited traps (e.g., Pherocon 1C, Zoecon) have been used to monitor male flight activity (Anshelevich et al., 1994) and to make informed treatment decisions in grape production areas. Traps placed 4 ft. high (1.3 m) are generally more effective than traps placed at only 1 ft. (0.3 m). Delta traps catch relatively fewer moths than traps with a more open design, e.g., traptest traps described as “commercial type (Montedison, Milan, Italy), consisting of two triangular plastic roofs in Havana brown; with a sticky area of 9.89 dm2 [152 in2]”. When pheromone traps are used, care should be taken to keep foliage away from the entry to the trap (PPQ, 1993). Rubber septa used to dispense the pheromone should be replaced every 3 weeks (PPQ, 1993). Traps should be placed approximately 100 ft. (30.5 m) apart to avoid inter-trap interference.

84 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Visual survey: USDA (1985) suggests visually inspecting for eggs on flower buds or pedicels of vines and grapes. It is preferable to look for larval damage rather than for eggs, because detection of eggs is very tedious and time-consuming, especially under field conditions. Look for webbed bud clusters (glomerules) or flowers where the spring generation larvae feed. Inspect for pupae under rolled leaves in spring. Inspect grapes and look for eggs or damaged berries. Cut open grapes and search for summer generation larvae (Fig. 5) and pupae. Suspect adult specimens should be pinned and labeled for subsequent identification. Submit suspect larvae or pupae in alcohol. For field surveys, Badenhauser et al. (1999) recommended a sample unit of a grapevine. Sample units should be selected at random.

Not recommended: Light traps have been used, but their lack of specificity and the fact that this moth flies at dusk competing for light makes their use inadvisable when the appropriate pheromones are available. Feeding traps were largely used in the past before pheromone traps were developed, but may still be useful in particular situations. An earthen or glass pot is baited with a fermenting liquid (fruit juice, molasses, etc.), and the scents produced attract adults, which are then drowned. Practical problems include irregularity in trapping because fermentation strongly depends on seasonal temperature, trap maintenance (lure replenishment and foam elimination), and low selectivity.

A corrugated paper band technique has sometimes been employed to trap and quantify overwintering pupae. Bands are placed around grapevine trunks or primary branches, and diapausing larvae pupate inside. However, this method is only useful in the latter generations, and its reliability is uncertain.

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of L. botrana is by morphological identification. Larvae can be keyed out using Gilligan et al. (2008). Identification of adults requires dissection of the male genitalia; use Brown, (2009) and Passoa, (2009).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Morphological identification is required for L. botrana. Hindwing coloration and the male clasper lacks spine at base (Venette et al. 2003). The key in MacKay (1959), or the simplified version in Passoa (2008), can be used to separate L. botrana larvae from many other Olethreutinae in the United States.

A new identification tool, Tort AI – Tortricids of Agricultural Importance, is available at http://idtools.org/id/leps/tortai/ from CPHST’s Identification Technology Program. This tool contains larval and adult keys, fact sheets, an image gallery, molecular search capacity, and more. Lobesia botrana is included in this tool.

Easily Confused Pests

85 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Paralobesia viteana is a native North American pest of grapes with an almost identical larval morphology to L. botrana (Gilligan et al., 2011). Pupae of the two species are distinct. The broad cremaster lacking thick curved hooks at the lateral margin, the presence of spines on A9, the lack of setae on the anal rise and presence of a spine patch on A10 define Lobesia. Adults of the two species are similar in size and wing pattern but can be separated by genitalic structures. P. viteana has a sclerotized lobe projecting from the ventral base of the male cucullus that is absent in all other Nearctic olethreutines, and the female corpus bursae lacks a signum and has two small lobelike anterior accessory bursae (Gilligan et al., 2011). The two species presently have different distributions: P. viteana occurs in the eastern United States, ranging as far west as Colorado; while L. botrana is currently restricted to California.

L. botrana can also be confused with Endopiza viteana (present in the United States) and Eupoecilia ambiguella (not present in the United States). The American grape berry moth, Endopiza viteana [Polychrosis viteana], occurs in the eastern United States and presents similar bionomics to L. botrana (Venette et al., 2003).

In the Palaearctic vine-growing areas, other lepidopteran species have an ecological niche similar to that of L. botrana, including Eupoecilia ambiguella, Argyrotaenia pulchellana [Argyrotaenia ljungiana], Clepsis spectrana, Cryptoblabes gnidiella, Euzophera bigella, and Ephestia parasitella. Even the primarily phytophagous Sparganothis pilleriana may sometimes damage grapes. However, only the first of these, E. ambiguella, may cause comparable damage to L. botrana, at least in northern European vineyards. Adults of these species may be easily differentiated macroscopically using a photographic key. E. ambiguella forewings are cream with a median fascia bluish dark brown. In field conditions, larvae may be distinguished because (i) the head of E. ambiguella is darker than that of L. botrana; (ii) L. botrana larvae do not carry any protective silk cover; and (iii) the behavior of L. botrana when disturbed is quicker and even violent. Moreover, L. botrana pupation occurs inside a grayish white cocoon that usually does not incorporate vegetal residues and frass, as occurs in E. ambiguella (CABI, 2009).

References Aarvik, L. 2011. Fauna Europaea: Lobesia botrana. In O. Karsholt and E. J. Nieukerken (eds.) Fauna Europaea: Lepidoptera, Moths, version 2.4. Accessed April 11, 2012 from: http://www.faunaeur.org.

Anshelevich, L., Kehat, M., Dunkelblum, E., and Greenberg, S. 1994. Sex pheromone traps for monitoring the European vine moth, Lobesia botrana: Effect of dispenser type, pheromone dose, field aging of dispenser, and type of trap on male captures. Phytoparasitica 22: 281-290.

Avidov, Z., and Harper, I. 1969. Lobesia (=Polychrosis) botrana (Schiff,). In: Plant pests of Israel. Jerusalem: Israel Univ. Press: 380-384.

Badenhausser, I., Lecharpentier, P., Delbac, L., and Pracros. P. 1999. Contributions of Monte-Carlo test procedures for the study of the spatial distribution of the European vine moth, Lobesia botrana (Lepidoptera: Tortricidae) in European vineyards. European Journal of Entomology 96: 375-380.

Brown, J. 2009. Adult Lepidoptera Workshop.

86 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

CABI. 2009. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

CIE. 1974. Lobesia botrana (Schiff.). Distribution Maps of Pests, Series A (Agricultural). Map No. 70 (revised). Commonwealth Institute of Entomology.

Coscollá, R. 1997. La polilla del racimo de la vid (Lobesia botrana Den. y Schiff.). Valencia, Spain: Generalitat Valenciana, Consejerfa de Agricultura, Pesca y Alimentación.

EPPO. 2012. EPPO Plant Quarantine Information Retrieval System (PQR), version 5.0.5540. European and Mediterranean Plant Protection Organization.

Fermaud, M., and Le Menn, R. 1989. Association of Botrytis cinerea with grape berry moth larvae. Phytopathology 79(6):651-656.

Filip, I. 1986. Breeding zones of the grape moth (Lobesia botrana Den & Schiff) in Romania. Probleme de Protectia Plantelor 14(1):25-30.

Gabel, B. 1981. On the influence of temperature on the development and reproduction of the vine moth, Lobesia botrana Den. & Schiff. (Lepid., Tortricidae). Anz. Schadlingskunde Pflanz. Umweltschutz 54 (6): 83-87 (In German; English Summary). Taken from Rev. Appl. Entomol. Ser. A 70 (3): 186.

Gilligan T.M., Wright, T.J., and Gibson, L. 2008. Olethreutine Moths of the Midwestern United States. An Identification Guide. Bulletin of the Ohio Biological Survey, new series, Volume 16(2) 334 pp.

Gilligan, T.M., Epstein, M.E., Passoa, S.C., Powell, J.A., Sage, O.C., and Brown, J.W. 2011. Discovery of Lobesia botrana ([Denis & Schiffermuller]) in California: an invasive species new to North America (Lepidoptera: Tortricidae). Proceedings of the Entomological Society of Washington 113(1): 14- 30.

Gonzalez, R. H. 2008. Biologıa, desarrollo, caracterizacion, de danos y manejo fitosanitario de la polilla Europea de la vid, Lobesia botrana (D and S) (Lep., Tortricidae). Universidad de Chile, Direccion de Extension, Santiago, Chile. 25 pp.

Hoenisch, R.W. No date. The European Grapevine Moth, Lobesia botrana Denis & Schiffermüller, 1775 - Lepidoptera, Tortricidae First U.S. Report. Western Plant Diagnostic Network Pest Alert. 4 pp.

Kearfott, W. D. 1904. North American Tortricidae. Transactions of the American Entomological Society 30: 287–299.

MacKay, M. R. 1959. Larvae of the North American Olethreutidae (Lepidoptera). The Canadian Entomologist Supplement 10. 338 pp.

Maher, N. M., Toulouse, M.E., Jolivet, J., and Thiéry, D. 2000. Oviposition preference of the European grapevine moth, Lobesia botrana (Lepidoptera: Tortricidae) for host and non-host plants present in Bourdeaux area. Bulletin OILB-SROP 23: 131-134.

Maher, N. M., Jolivet, J., Thiéry, D., and Lozzia, C. 2001. Preference on the European grapevine moth, Lobesia botrana (Lepidoptera, Tortricidae) between different types of vine: influence of chemical information on the surface of berries. Bulletin OILB-SROP 24: 103-107.

Passoa, S. 2008. Part III: Immature stages, pp. 295–314. In: T. M. Gilligan, D. J. Wright, and L. D. Gibson, eds. Olethreutine Moths of the Midwestern United States, an Identification Guide. Ohio Biological Survey. Columbus, Ohio.

87 Lobesia botrana Primary Pest of Grape Arthropods European grapevine moth

Passoa, S. 2009. Screening Key for CAPS Target Tortricidae in the Eastern and Midwestern United States (males). Lab Manual for the Lepidoptera Identification Workshop. University of Maryland.

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

Robinson, G.S., Ackery, P.R., Kitching, I.J., Beccaloni, G.W., and Hernández, L.M. 2011. HOSTS - a Database of the World's Lepidopteran Hostplants: Lobesia botrana. London: Natural History Museum. Accessed April 13, 2012, from http://www.nhm.ac.uk/research- curation/research/projects/hostplants/search//index.dsml.

Rodionov, Z.S. 1945. Pest of vines in the Turkmen SSR. (English abstract in Rev. Appl. Entomol., 1948, 36:3).

Savopoulou-Soultani, M. and Tzanakakis, M.E. 1988. Development of Lobesia botrana (Lepidoptera: Tortricidae) on grapes and apples infected with the fungus Botrytis cinerea. Environmental Entomology 17(1):1-6.

SENASA (Servicio Nacional de Sanidad y Calidad Agroalimentaria). 2010. El Senasa declaro´ la emergencia fitosanitaria por la deteccio´n de la plaga Lobesia botrana. http://www.senasa.gov. ar/contenido.php?to=n&in=1453&ino=0&io=12316.

Stavridis, D.G. and Savopoulou-Soultani, M. 1998. Larval performance on and oviposition preference for known and potential hosts by Lobesia botrana (Lepidoptera: Tortricidae). European Journal of Entomology 95: 55-63.

Torres-Vila, L.M. 1995. Factores reguladores del potencial biótico y de la poliandria en la polilla del racimo de la vid Lobesia botrana Den. y Schiff., (Lepidoptera: Tortricidae). Madrid, Spain: Tesis Doctoral de la ETS de Ingenieros Agrónomos de la Universidad Politécnica de Madrid.

Torres-Vila, L.M., Stockel, J., and Roehrich, R. 1995. The reproductive potential and associated biotic variables in the male of the European vine moth Lobesia botrana. Entomologia Experimentalis et Applicata 77(1):105-119.

Torres-Vila, L.M. Stockel, and J. Rodriguez-Molina, M.C. 1996. Efecto de la indisponibilidad de agua sobre el potencial biótico de la polilla del racimo Lobesia botrana Den. y. Schiff. (Lepidoptera: Tortricidae). Boletfn de Sanidad Vegetal Plagas 22:443-449.

Torres-Vila, L.M., Stockel, J., and Rodríguez-Molina, M.C. 1997. Physiological factors regulating polyandry in Lobesia botrana (Lepidoptera: Tortricidae). Physiological Entomology 22(4):387-393.

USDA. 1985. Pests not know to occur in the United States or of limited distribution. No. 60: European Grape Vine Moth.

Venette, R. C., Davis, E.E., Dacosta, M., Heisler, H., and Larson, M. 2003. Mini Risk Assessment – Grape berry moth, Lobesia botrana (Denis & Schiffernuller) [Lepidoptera: Tortricidae]. http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/lbotranapra.pdf.

88 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Scirtothrips dorsalis

Scientific Name Scirtothrips dorsalis Hood

Synonyms: Anaphothrips andreae, Heliothrips minutissimus, Neophysopus fragariae, Scirtothrips andreae, S. fragariae, S. minutissimus, and S. padmae Common Name(s) Castor thrips, chilli thrips, yellow tea thrips, grapevine berry thrips, strawberry thrips

Type of Pest Thrips

Taxonomic Position Class: Insecta, Order: Thysanoptera, Family: Thripidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List (2006- 2008)

Figure 1. Scirtothrips dorsalis Pest Description adults. Courtesy of T. Skarlinsky Scirtothrips dorsalis is a widespread pest, (USDA APHIS PPQ) described as a new species by Hood in 1919. S. dorsalis is a very small (0.5 to 1.2 mm), pale yellow-colored thrips (Fig. 1) that can be found feeding on leaves, flowers, and calyxes of fruit on a wide variety of host crops. It is difficult to recognize this thrips with the naked eye, and definitive identification is best accomplished at approximately 40 to 80X magnification.

Eggs: Typically oval, whitish to yellowish, narrow anteriorly, with an incubation period of six to eight days (CABI, 2007). Eggs are about 0.075 mm (0.00295 in.) long and 0.070 mm (0.00275 in.) wide, and are inserted inside plant tissue.

Larvae: Two larval stages (first and second instar) last for six to seven days. The larvae are off-white in color. First instar: transparent; body short, legs longer; antennae short, swollen; mouth cone bent and short; and antennae seven-segmented and cylindrical. Sclerotization not distinct, head and thorax reticulate (CABI, 2007).

89 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Second instar: antennae longer, cylindrical, seven-segmented; mouth cone longer; maxillary palpi three-segmented; body setae longer than the first instar; head and thorax reticulate with sclerotization of head (CABI, 2007).

Pre-pupae: Yellowish; antennae swollen, short, with distinct segmentation; two pairs of external wing buds on each meso- and metathorax (CABI, 2007). The pre-pupal period is short (~24 hours).

Pupae: Dark yellow with eyes and ocelli bearing red pigmentation; wing buds are elongate; antennae short and reflected over the head; female pupae with larger pointed abdomen; males have a smaller, blunt abdomen (CABI, 2007). The pupal period lasts two to three days. Pupation takes place in the axils of leaves, in leaf curls, and under the calyxes of flower and fruits.

Adults: Almost white on emergence, turning yellowish subsequently (Fig. 1) with incomplete dark stripes on the dorsal surface where the adjacent abdominal segments meet. A technical description follows: abdominal tergites with median dark patch, tergites and sternites with dark antecostal ridge; ocellar setae pair III situated between posterior ocelli; two pairs of median post-ocular setae present; pronotum with four pairs of posteromarginal setae, major setae 25 to 30 µm long; metanotum medially with elongate recticles or striations, arcuate in anterior third, median setae not at anterior margin; forewings with fore marginal setae, second vein with two setae, cilia straight; tergal microtrichial fields with three discal setae, VIII and IX with microtrichia medially; sternites with numerous microtrichia, more than two complete rows medially; male without drepanae on tergite IX (Palmer and Mound, 1983).

Biology and Ecology: Reproduction is both sexual and parthenogenetic. In India, where the life cycle has been studied particularly, adults typically mate two to three days after their pupal molt and females start ovipositing on Ricinus spp. three to five days after emergence. The total number of eggs laid ranges from 40 to 68 (Venette and Davis, 2004). The life cycle is completed in 15 to 20 days, and the sex ratio is 6:1 females to males. On chillies, a single female lays two to four eggs per day for a period of about 32 days. In the Guntur area of India, S. dorsalis appears in two distinct periods: in the nurseries in August- September, when it is not serious, and from the third week of November to March. S. dorsalis typically undergoes four to eight generations per year (Venette and Davis, 2004). A degree day model was developed by Meisner et al. (2005) to determine the potential number of generations per year that the United States climate could support, as well as pinpoint the areas that could climatically support establishment. The model was based on a degree day (DD) requirement of 281 days per generation. In addition the model excluded regions where the minimum temperature reached -4°C (24.8°F) or below for five consecutive days. The results showed that parts of Florida, Texas, Arizona, California, and Nevada could sustain up to 18 generations per year and that several additional states (Louisiana, Alabama, Louisiana, Georgia, and South Carolina) are susceptible to establishment (Fig. 2).

90 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Figure 2. Generational potential (based on a generational requirement of 281 DD and a base and upper development temperature of 9.7°C (49.46°F) and 33.0°C (91.4°C), respectively) outside of the predicted cold temperature exclusion boundary (areas where the minimum daily temperature reaches -4ºC (24.8°F) or below on five or more days per year) for S. dorsalis in the United Statesand Mexico (from Meisner et al., 2005).

Pest Importance S. dorsalis is a significant pest of chilli pepper, citrus, castor bean, cotton, onion, and other crops in tropical and subtropical regions of Asia, Africa, eastern Europe, Oceania, and Japan. Recently, S. dorsalis was confirmed for the first time from several Caribbean Islands, Puerto Rico, Florida, and Texas. S. dorsalis is widespread in Florida, but it is not known if it is present or established in any other part of the continental United States.

On grapes, heavy feeding damage to flower clusters and developing fruit has resulted in reduced fruit set and reduced marketability of damaged fruits (Ananthakrishnan, 1971). In Asia, S. dorsalis is a pest of economic importance in pest of citrus and feeding by the thrips causes significant leaf and flower deformation, fruit damage, and yield reduction. In Taiwan, a range of damage on mango, from fruit scarring to total plant defoliation has been reported (Lee and Wen, 1982).S. dorsalis is also an economically important pest of chilli pepper, castor bean, cotton, and onion; in these crops, thrips feeding can wilt, distort, or stunt young leaves/shoots and cause premature leaf, bud, or flower drop. In some varieties of chilli peppers, ~75% of leaves may be deformed due to the activity of piercing-sucking insects. Yield losses attributable to S. dorsalis in chilli have ranged from 20% (Ahamad et al., 1987) to nearly 50% (Sanap and Nawale, 1987; Varadharajan and Veeravel, 1996). In cotton, sucking pests, including S. dorsalis, reduced yield of seed by 77%; fiber yields and quality were also diminished (Gupta and Gupta, 1999). Estimated yield loss has been recorded at 25 to 67% when

91 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

population density is high. On tea, feeding occurs on new growth, including leaves, shoots and buds, and occasionally on older leaves, resulting in browning and defoliation (Ananthakrishnan, 1971). Scirtothrips dorsalis is also considered a major pest of roses in India, where its feeding distorts or destroys leaves, buds, and flowers and reduces marketability (Gahukar, 1999). On mango, feeding damage occurs on new growth, on the underside of leaf surfaces at the midrib, and on fruits (Zaman and Maiti, 1994). Damaged tissues appear dark in color, and leaves curl and drop prematurely (Kumar et al., 1994).

This insect is also a key vector of Tomato spotted wilt virus (TSWV), which causes bud necrosis disease (BND), an important disease of peanut in India (Amin et al., 1981; Mound and Palmer, 1981; Ananthakrishnan, 1993; Lewis, 1997). TSWV is widely distributed in the eastern United States on a variety of ornamental, field crops (peanut, tobacco, tomato), and weeds. S. dorsalis is also known to vector peanut chlorotic fan- spot virus, peanut yellow spot virus and tobacco streak virus (Rao et al., 2003). It is also reported to transmit the bacterial leaf spot bacterium (Ananthadrishnan, 1993; Mound, 1996; and Mound and Palmer, 1981).

Symptoms/Signs As with other plant-feeding thrips, damage is caused by sucking out sap from individual epidermal cells, leading to necrosis of the tissue. Damage is most severe at the growing tips, on young leaves and shoots, or on flowers and young fruits. Heavy feeding damage turns tender leaves, buds, and fruits bronze to black in color. Damaged leaves curl upward (Fig. 3) and appear distorted. Infested plants become stunted or dwarfed, and leaves with petioles detach from the stem, causing defoliation in some plants. Symptoms include: silvering of the leaf surface; frass-marked staining or scarring of the fruit, particularly around the apex or at the rim of the calyx; distortion and staining Figure 3. Leaf curling of both leaves (curling and thickening of the lamina) symptom. Photo courtesy and fruit; and ultimately, the early senescence of the of CABI, 2007. leaves. S. dorsalis rarely feeds on mature leaves.

In tropical regions, the abundance of chilli thrips is low in the rainy season, but becomes high during the dry season. As is typical of species in the genus Scirtothrips, eggs are laid in the youngest tissues of plants, and feeding by adults and larvae can result in extensive cell damage to these developing tissues, leading to leaf and fruit distortion, and flower fall.

On chillies, S. dorsalis causes 'leaf curl disease.’ Heavy infestation of the tender shoots, buds and flowers causes the leaves to curl badly (Fig. 3) and drop; and fresh buds to become brittle and subsequently drop down.

92 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

On grape, the fruits and leaves are attacked, which causes damage and/or A scarring on the leaves (Fig. 4A,) on the surface of the berries (Fig. 4B), on the rachis (Fig. 4C). Berries are attacked, starting at even the button stage, and significant scarring develops if control measures are not taken during berry formation. In severe cases of infestation, the berry bursts and exposes the seeds. The infestation can last until fruit maturation and facilitate the secondary infestation of the fruit by certain flies and fungi. The affected fruit become unfit for consumption, canning, or preservation (Perumal, 1972). On B berries that grow after harvest or out of season, damage can be severe (Fig. 5).

Known Hosts S. dorsalis is highly polyphagous and has been recorded from more than 112 A plant species spread across 40 different families. Its main wild host plants are thought to be various Fabaceae such as Acacia spp., Brownea spp., Mimosa spp., and Saraca spp. Economically important hosts are identified below. Venette & Davis (2004) provide a thorough review of host plants known C from the scientific literature. Recent field surveys in the Caribbean have shown that S. dorsalis is found on cucumbers, cantaloupe, watermelon, pumpkin and squash, hot peppers, tomato, eggplant, okra, and beans (Ciomperlik & Seal, 2004).

Major hosts: Allium cepa (onion), Anacardium occidentale (cashew nut), Arachis hypogaea (peanut), Camellia sinensis Figure 4. Damage caused by S. dorsalis (tea), Capsicum frutescens (chilli on grape leaves and fruit. Photos courtesy pepper), Citrus spp., Fragaria spp. of Dr. Magally Quiros, University of Zulia, (strawberry), Gossypium spp. (cotton), Maracaibo, Venezuela.

Hevea spp. (rubber), Hydrangea spp.,

93 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Mangifera indica (mango), Nelumbo spp. (rose), Solanum lycopersicum (tomato), Tamarindus indica (tamarind), and Vitis spp. (grape). S. dorsalis is only cited as a significant pest of Citrus in Japan and Taiwan.

Minor hosts: Abelmoschus esculentus (okra), Acer spp. (maple), Asparagus officinalis (asparagus), Chrysanthemum spp., Cucumis spp. (cucumber and melon), Cucurbita moschata (pumpkin), Dahlia spp., Euonymus japonicus, Ficus carica (common fig), Glycine max (soybean), Laurus nobilis (bay laurel), Momordica charantia (balsam pear), Musa spp. (banana), Phaseolus vulgaris (bean), Pieris japonica, Prunus spp. (cherry), Pyrus spp. (pear), Quercus glauca (Japanese blue oak), Rhododendron spp., Solanum melongena (egg plant), Syzygium malaccense (malay apple), Virburnum spp., and Vigna radiata (mung bean).

Known Vectors (or associated organisms) S. dorsalis is a vector of Tomato spotted wilt virus (TSWV). S. dorsalis is also known to vector peanut bud necrosis virus (PBNV), peanut chlorotic fan-spot virus (groundnut chlorotic fan-spot virus), peanut yellow spot virus, Tobacco streak virus, (Hodges et al., 2005; Koike et al., 2008)

Known Distribution Asia: Bangladesh, Brunei, China (including Hong Kong), India, Figure 5. Severely damaged grape. Photo Indonesia, Israel, Japan (including courtesy of Dr. Magally Quiros, University Ryukyu Islands),Korea, Malaysia, of Zulia, Maracaibo, Venezuela. Myanmar, Pakistan, Philippines, Sri Lanka, Taiwan, Thailand, and Vietnam. Africa: Côte d’Ivoire. North America: United States (Florida, Georgia, Hawaii, and Texas). Oceania: Australia, New Caledonia, Papua New Guinea, and Solomon Islands. Caribbean: Recently reported as an invasive species in Puerto Rico, Barbados, Jamaica, Saint Lucia, Saint Vincent and the Grenadines, and Trinidad and Tobago. South America: Reported in Suriname and reported causing damage to grapevine in Venezuela (CABI, 2010; Riley et al., 2011; PIPDL, 2012).

This species has been intercepted in the United Kingdom (UK ) from material originating from Ghana (Jones, 2005). The population in the UK is considered transient and under eradication (EPPO, 2012).

94 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Potential Distribution in the United States S. dorsalis was first identified from Hawaii in 1987. In October 2005, S. dorsalis was positively identified by USDA-ARS-SEL from specimens submitted by Palm Beach County, Florida. As of 11/30/05, S. dorsalis has been collected in Florida 77 times in 16 counties. The pest was primarily found on potted roses sold in retail outlets from the Florida Keys to Tallahassee. A few positive detections occurred on pepper and Illicium spp. Only 2 of 77 positives were in a natural environment, so it is unknown at this time if S. dorsalis is established in the natural environment. Subsequent to the October detection, on 11/16/05, additional specimens were identified from Texas. Surveys in Texas have resulted in positive detections in 3 counties on Capsicum spp. on 11/10/05 and 11/14/05. Recently in early 2006, S. dorsalis was confirmed from samples collected in Jardin La Ceiba, Puerto Rico. Venette and Davis (2004) indicate that the potential geographic distribution of S. dorsalis in North America would extend from southern Florida to north of the Canadian boundary, as well as to Puerto Rico and the entire Caribbean region with approximately 28% of the continental United States having a suitable climate for S. dorsalis. Meisner et al. (2005) indicated that permanent establishment would likely be limited to southern and West Coast states. A recent risk analysis by USDA-APHIS-PPQ-CPHST shows that the greatest risk for establishment of S. dorsalis based on host presence and climate occurs in the eastern United States, California, Colorado, Idaho, Kansas, Nebraska, Oklahoma, Oregon, New Mexico, North Dakota, South Dakota, Texas, Utah, and Washington.

Survey CAPS-Approved Method*: Visually inspect hosts for damage symptoms. Collect terminal foliage and rinse with 70% ethanol. Collect ethanol and thrips for identification

Method Note: Sticky traps are not to be used for Chilli thrips surveys. Removing specimens for identification from the traps is nearly impossible.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Visual survey: Researchers typically depend on visual inspections of plant material to monitor populations of S. dorsalis. S. dorsalis feeds on new growth of nearly all vegetative plant parts (buds, leaves, flowers, fruits, and stems) (Ananthakrishnan, 1971; Chang et al., 1995). As with many thrips species, feeding deforms young leaves and stains or scars fruits (Chang et al. 1995). At the initial stage of infestation, the underside surfaces of the leaves become shiny. S. dorsalis is found on the leaves, flowers, and fruits of the hosts listed in this document. Thus, malformed fruits and foliage should be examined during inspections of potential host material. However, because of their small size and propensity to occur inside flowers or buds, S. dorsalis easily could be missed during visual inspections. Visual inspections are particularly likely to be ineffective if thrips densities are low. Pupae may be found in the axils of leaves, in leaf curls, and under the calices of flower and fruits, as well as in the soil.

95 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Suwanbutr et al. (1992) rinsed thrips from plant material using 70% ethanol and counted individuals collected on a fine muslin sieve. In Florida, five to 20 leaves from symptomatic plants are collected at random and placed in a ziplock bag to prevent the adults from escaping. The bag is labeled with collection locality information, host plant, date collected, and the name of collector. The samples are sent for next-day delivery to an expert for further processing to establish or confirm their identity. Foliage is washed in 70% ethanol to remove the adults and immatures. The alcohol and thrips samples are screened through a mesh diameter of 0.5 mm (0.02 in.) or less and observed under a stereomicroscope. Species level identifications should be made by a qualified taxonomist. The Florida method is recommended to enable morphological and/or molecular diagnostics to be used.

Trapping: No pheromones have yet been identified for this species.

Not recommended: Adults may also be attracted to yellowish-green, green, or yellow boards (Tsuchiya et al., 1995). In Japan, yellow sticky traps (10 x 20 cm (3.94 x 7.87 in.) were used to monitor S. dorsalis in vineyards, and these traps provided a coarse, but representative, estimate of population density on foliage (Shibao et al., 1990). A round, yellow sticky trap (15 cm (5.90 in.) diameter x 30 cm (11.81 in.) height) has also been used (Shibao et al,. 1993); traps were placed 1.2 m (3.94 ft.) above the ground at a density of four traps per 9 m2 (29.53 ft2). Counts on yellow sticky traps are correlated with damage to grape (Shibao, 1996). Chu et al. (2006) evaluated the effectiveness of a Blue-D, CC, and sticky traps for monitoring S. dorsalis in Taiwan and St. Vincent. The authors recommended that a combination of visual observation, yellow sticky traps, and CC traps may be an effective S. dorsalis population detection and monitoring system. For CAPS surveys, yellow sticky traps have traditionally been employed; however, thrips specimens may be damaged in the traps, thereby hindering morphological identification. The same is true for white, blue, or yellow CC traps. When possible, whole plant specimens or thrips rinsed from specimens should be sent to diagnostic labs.

Other methods include: (1) shaking inflorescences over black paper to count nymph and adult thrips (Gowda et al, 1979); (2) dislodging thrips from young shoots on a single plant over a piece of black cardboard and counting the recovered insects (Bagle, 1993; Sureshkumar and Ananthakrishnan, 1987); (3) using sticky suction traps to monitor the flight of S. dorsalis (Takagi, 1978); and (4) directly counting the number of thrips on plant shoots or terminal leaves (Khanpara and Patel, 2002). However, these methods are usually used in regions where the pest is established. Due to the difficulty of identification of S. dorsalis, these methods are not recommended for an early detection survey.

A Berlese funnel has been used to determine the presence of thrips in bulky plant material. However, its efficiency has not been evaluated for Scirtothrips dorsalis. (See http://www.extento.hawaii.edu/Kbase/reports/berlese_funnel.htm for further information).

96 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

For identifiers at the ports, a shaker box technique has been developed to detect S. dorsalis and other actionable pests in peppers from St. Lucia and St. Vincent and was found to be superior to detecting thrips when compared to a 2% visual inspection.

For additional information on chilli thrips, including photos of symptomatic plants and chilli trips, sampling, management, and more see: http://mrec.ifas.ufl.edu/lso/thripslinks.htm.

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation requires a morphological identification. Identification of adults and late instar larvae is possible with some expertise/ experience in Scirtothrips/Thysanoptera as per National Identification Service (NIS), states may go to reliable entomologists/ taxonomists for identification. The Chilli Thrips Task Force has named G.B. Edwards, Gillian Watson, Jack Reed, and Cindy McKenzie as approved taxonomists for chilli thrips identification. Other thrips families and genera, Frankliniella schultzei, S. aurantii, and S. oligochaetus may be confused with S. dorsalis.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Identification during immature and adult stages can be done most reliably by polymerase chain reaction and restriction fragment length polymorphism (PCR-RFLP), using a method described by Toda and Komazaki (2002) and Brunner et al. (2002).

Adult members of the genus Scirtothrips are readily distinguished from all other Thripidae by the following characters: surface of pronotum covered with many closely spaced transverse striae; abdominal tergites laterally with numerous parallel rows of tiny microtrichia; sternites with marginal setae arising at posterior margin; and metanotum with median pair of setae arising near anterior margin. Larvae are pale and indistinguishable from the larvae of other thrips species (CABI, 2007)

To diagnose S. dorsalis: Abdominal sternite with dark antecostal ridge. Ridge is not always visible for teneral adults. Lateral microtrichial fields of abdominal tergites with three discal setae. Posteromarginal comb on abdominal tergite VIII complete. Forewing shaded, lighter distally with straight cilia. Second vein: incomplete with two or three intermittent setae in distal half. Forked sense cone. Antennal segments I-II pale, III-VIIII dark. Head with three pairs of ocellar setae. Ocellar setae III between posterior ocelli.

97 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

Figure 6. Chaetanaphothrips orchidii (left) and Selenothrips rubrocinctus (right). Photos courtesy of Lance Osbourne (http://mrec.ifas.ufl.edu/lso/thripslinks.htm) and the University of Florida, respectively.

Note: the multiple transverse striae are characteristic of the genus. Posteromarginal seta II is broader and about 1.5 times longer than posteromarginal setae I and III. For images and further information on the diagnostic features of S. dorsalis see http://mrec.ifas.ufl.edu/lso/DOCUMENTS/Scirtothrips%2520dorsalis%2520Hood%2520I D%2520Aid.pdf.

An identification system, fully illustrated with photomicrographs of structural details, together with a molecular method for distinguishing this species from related species is provided in the thrips identification tool, “Pest Thrips of the World”, available for purchase at http://www.cbit.uq.edu.au/software/pestthrips/purchase.htm.

Easily Confused Pests Thrips are extremely small and very difficult to distinguish from one another, especially when immature. Adults and late-instar larvae may be identified upon close examination of morphological characters by a taxonomist (Toda and Komazaki, 2002).

Scirtothrips dorsalis has reportedly been confused with S. aurantii (South African citrus thrips), S. oligochaetus, and Frankliniella schultzei on various hosts (Amin and Palmer, 1985; Gilbert, 1986; Toda and Komazaki, 2002). Another similar species is Drepanothrips reuteri, a native European pest of grapevine, which has six-segmented antennae (the 3 terminal segments being fused) instead of the eight-segmented found in S. dorsalis (CABI, 2007).

Chaetanaphothrips orchidii (orchid thrips) is sometimes confused with S. dorsalis in Florida (Fig. 6). This species of thrips looks very much like chilli thrips. However, there is a “break” in the dark coloring of the wings. This gives the orchid thrips the appearance of having t dark spots on the front portion of the abdomen. The red banded

98 Scirtothrips dorsalis Primary Pest of Grape Arthropods Chilli thrips

thrips, Selenothrips rubrocinctus (Fig. 6), causes similar damage to roses but the adult red banded thrips is black in color whereas the chilli thrips is much smaller and very light yellow, brown, or straw colored.

References Ahamad, K., Mohamed, M.G., and Murthy, N.S.R. 1987. Yield losses due to various pests in hot pepper. Capsicum-Newsletter: 83-84.

Amin, P.W., and Palmer, J.M. 1985. Identification of groundnut Thysanoptera. Tropical Pest Management 31: 286-291, 340, 344.

Amin, P.W., Reddy, D.V.R., and Ghanekar, A.M. 1981. Transmission of tomato spotted wilt virus, the causal agent of bud necrosis of peanut, by Scirotothrips dorsalis and Frankliniella schultzei. Plant Disease 65: 663-665.

Ananthakrishnan, T.N. 1971. Thrips in agriculture, horticulture and forestry - diagnosis, bionomics and control. Journal of Scientific and Industrial Research (CSIR), New Delhi, 30:130-146.

Ananthakrishnan, T.N. 1993. Bionomics of thrips. Annual Review of Entomology, 38:71-92.

Bagle, B.G. 1993. Seasonal incidence and control of thrips Scirtothrips dorsalis Hood in pomegranate. Indian Journal of Entomology 55: 148-153.

Brunner, P.C., Fleming, C., and Frey, J.E. 2002. A molecular identification key for economically important thrips species (Thysanoptera: Thripidae) using direct sequencing and a PCR-RFLP-based approach. Agricultural and Forest Entomology 4: 127-136.

CABI. 2007. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/. CABI. 2010. Distribution Maps of Plant Pests, Scirtothrips dorsalis. Map No. 475, 1st revision.

Chang, N. T., Parker, B.L., Skinner, M. and Lewis, T. 1995. Major pest thrips in Taiwan, pp. 105-108, Thrips biology and management: Proceedings of the 1993 International Conference on Thysanoptera. Plenum Press, New York.

Chu, C.C., Ciomperlik, M.A., Chang N.-T., Richards, M., Henneberry, T.J. 2006. Developing and evaluating traps for monitoring Scirtothrips dorsalis (Hood). Florida Entomologist 89(1): 48-55.

Ciomperlik, M. and Seal, D. 2004. Surveys of St. Lucia and St. Vincent for Scirtothrips dorsalis Hood, January 14-23, 2004. Edinburg, TX, USDA-APHIS-PPQ-CPHST-PDDML: 19.

Gahukar, R.T. 1999. New record of thrips attacking roses in central India. Journal of Insect Science (India) 12: 89.

Gilbert, M.J. 1986. First African record of Scirtothrips dorsalis Hood (Thysanoptera: Thripidae) a potential pest of citrus and other crops in southern Africa. Journal of the Entomological Society of Southern Africa 49: 159-161.

Gowda, G., Ramaiah, E., and Reddy, C.V.K. 1979. Scirtothrips dorsalis (Hood) (Thysanoptera: Terebrantia: Thripidae) a new pest on cashew (Anacardium occidentale L). Current Research 8: 116-117.

Gupta, M.P., and Gupta, D.P. 1999. Effect of incidence of insect pests and their control measures on fibre quality of cotton hybrids. Journal of Insect Science (India) 12: 58-62.

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Hodges, A., Osborne, L., Beck, H. and Ludwig, S. 2009. Chilli Thrips, Scirtothrips dorsalis E-Learning Module. http://cbc.at.ufl.edu/

Jones, D.R. 2005. Plant viruses transmitted by thrips. European Journal of Plant Pathology 113:119-157.

Khanpara, D.V., and Patel, G.M. 2002. Need based plant protection and avoidable losses in hybrid castor. Indian Journal of Entomology 64: 175-184.

Koike, S.T., Davis, R.M., Subbarao, K.V., and Falk, B.W. 2008. How to Manage Pests: UC Pest Management Guidelines. Peppers - Tomato Spotted Wilt Virus (Publication 3460). University of California, Agriculture and Natural Resources, Statewide IPM Program. http://www.ipm.ucdavis.edu/PMG/r604100911.html.

Kumar, S., Patel, C.B., Bhatt, R.I., and Rai, A.B. 1994. Population dynamics and insecticidal management of the mango thrips, Scirtothrips dorsalis Hood (Thysanoptera: Thripidae) in South Gujarat. Pest Management and Economic Zoology 2: 59-62.

Lee, H.S., and Wen, H.C. 1982. Seasonal occurrence of and injury caused by thrips and their control on mangoes. Plant Protection Bulletin, Taiwan. 24: 179-187.

Lewis, T. [ed.] 1997. Thrips as Crop Pests. CAB International, Wallingford, UK.

Meissner, H., Lemay, A., Borchert, D., Nietschke, B., Neeley, A., Magarey, R., Ciomperlik, M., Brodel, C., and Dobbs, T. 2005. Evaluation of Possible Pathways of Introduction for Scirtothrips dorsalis Hood (Thysanoptera: Thripidae) from the Caribbean into the Continental United States. Center for Plant Health Science and Technology, USDA-APHIS, Raleigh, North Carolina, 125 pp.

Mound, L.A. 1996. The Thysanoptera vector species of tospoviruses. In: Kuo, C.G. (Ed.), Tospoviruses and Thrips of Floral and Vegetable Crops. Acta Horiticulturae. An International Symposium, Taichung, China. Technical Communications of the International Society of Horticultural Science, vol. 431, pp. 298- 309.

Mound, L.A., and Palmer, J.M. 1981. Identification, distribution, and host plants of the pest species of Scirtothrips (Thysanoptera:Thripidae). Bull. Entomol. Res. 71: 467-479.

Natarajan, K., Sundaramurthy, V.T., and Chidambaram, P. 1991. Usefulness of fish oil rosin soap in the management of whitefly and other sap feeding insects on cotton. Entomon 16: 229-232.

Okada, T., and Kudo, I. 1982. Relative abundance and phenology of Thysanoptera in a tea field. Japanese Journal of Applied Entomology and Zoology 26: 96-102.

Palmer, J.M. and Mound, L.A. 1983. The Scirtothrips species of Australia and New Zealand (Thysanoptera: Thripidae). Journal of Natural History, 17(4):507-518

Panickar, B.K., and Patel, J.R. 2001. Population dynamics of different species of thrips on chilli, cotton and pigeonpea. Indian Journal of Entomology 63: 170-175.

Perumal, R.S. 1972. Grapevine berry thrips Scirtothrips dorsalis H. and its control. Farm Fact 6(5): 23-24.

PIPLD. 2012. Host list for Scirtothrips dorsalis in New Caledonia. Accessed April 13, 2012 from: http://pld.spc.int/pld/.

Rao, R.D.V.D.J.P.. Reddy, A.S., Reddy, S.V., Thirumala-Devi, K., Rao, S.C., Kumar, V.M., Subramaniam, K., Reddy, T.Y., Nogram, S.M., and Reddy, D.V.R. 2003. The host range of tobacco streak virus in India and transmission by thrips. Annals of Applied Biology 142(3): 365-368.

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Riley, D.G., Joseph, S.V., Srinivasan, R., and Diffie, S. 2011. Thrips vectors of Tospoviruses. J. Integ. Pest Mngmt 1(2): 10 pp.

Saboo, K.C., and Puri, S.N. 1978. Effect of insecticides in incidence of sucking pests and yield of groundnut, Arachis hypogea Linn. Indian Journal of Entomology 40: 311-315.

Sanap, M. M., and Nawale, R.N. 1987. Chemical control of chilli thrips, Scirtothrips dorsalis Hood (Thysanoptera: Thripidae). Vegetable Science 14: 195-199.

Saxena, P., Vijayaraghavan, M.R., Sarbhoy, R.K., and Raizada, U. 1996. Pollination and gene flow in chillies with Scirtothrips dorsalis as pollen vectors. Phytomorphology 46: 317-327.

Schall, R. 1995. NPAG Data: Scirtothrips dorsalis Chilli (Assam) thrips.

Seal, D.R., Ciomperlik, M.A., Richards, M.L., and Klassen, W. 2006. Distribution of chilli trips, Scirtothrips dorsalis (Thysanoptera: Thripidae), in pepper fields and pepper plants on St. Vincent. Florida Entomology 89(3): 312-320.

Shibao, M. 1996. Damage analysis of chilli thrips, Scirtothrips dorsalis Hood (Thysanoptera: Thripidae) on grape. Japanese Journal of Applied Entomology and Zoology 40: 293-297.

Shibao, M., Tanaka, F., and Nakasuji, F. 1990. Seasonal changes and infestation sites of the chilli thrips, Scirtothrips dorsalis (Thysanoptera: Thripidae) on grapes. Japanese Journal of Applied Entomology and Zoology 34: 145-152.

Shibao, M., Tanaka, F., Fujisaki, K., and Nakasuji, F. 1993. Effects of lateral shoot cutting on population density of the chilli thrips, Scirtothrips dorsalis Hood (Thysanoptera: Thripidae) on grape. Applied Entomology and Zoology (Japan) 28: 35-41.

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102 Spodoptera littoralis Primary Pest of Grape 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, 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, 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 2003 through 2012

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 whitish-yellow in color, changing to black just prior to hatching, due to the big head of the larva showing through the transparent shell Figure 1. Eggs and neonates. Eggs are (Pinhey, 1975). laid in batches covered with orange- brown hair scales. Photo courtesy of Larvae: Upon hatching, larvae are 2 to 3 http://www.defra.gov.uk/planth/pestnote/ mm (0.079 to 0.12 in.) long with white spod.htm. bodies and black heads and are very difficult to detect visually. Larvae grow to 40 to 45 mm (1.57 to 1.77 in.) and are

103 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth

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 segment. The larva of S. littoralis is figured by Bishari (1934) and Brown and Dewhurst (1975). Larvae are nocturnal and during the day can be found at the base of the plants or under pots.

Figure 2. Larva of S. littoralis. Photos courtesy of CABI, 2007.

A B

Figure 3. Pupa and adult of S. littoralis on soil (A). Adult moth of S. littoralis (museum set specimen) (B). Photos courtesy of CABI, 2007 and Entopix.

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 to 20 x 5 mm (0.55 to 0.79 x 0.20 in.), tapering towards the posterior segments of the abdomen. The last segment ends in two strong straight hooks (Pinhey, 1975).

Adults: Moth with gray-brown body (Fig. 3), 15 to 20 mm (0.59 to 0.79 in.) long; wingspan 30 to 38 mm (1.18 to 1.50 in.); forewings gray to reddish brown with paler lines along the veins (in males, bluish areas occur on the wing base and tip); the ocellus

104 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth is marked by two or three oblique whitish stripes. Hindwings are grayish white, iridescent with gray margins, and usually lack darker veins (EPPO, 1997).

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 to 1,000 eggs) on the lower leaf surface of younger leaves or upper parts of the plant. Eggs begin to hatch after 28.6 degree days (DD) at a base temperature of 14.8°C (58.64°F). The optimal temperature for egg hatch is 28 to 30°C (82.4 to 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 on the loose surface of the soil downwards until it reaches more solid ground 3 to 5 cm (1.18 to 1.97 in.) deep. It then creates a clay ‘cell’ or cocoon in which it usually pupates within five to six hours. Emergence of adult moths occurs at night, and they have a life span of five to ten days. Adults fly at night, mostly between the hours of 8 pm and midnight.

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 cowpeas 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).

105 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth

Symptoms/Signs On most crops, damage arises from extensive feeding by larvae, leading to complete stripping of the plants. On grape, larvae gnaw holes in the leaves until sometimes only the veins remain. The damage caused by larvae to grapevines is not merely temporary; vines may suffer so severely from exposure to intense sunlight during the summer that their development in the following year will be retarded. Larvae also gnaw at grape bunch stalks, which as a result, dry up, and the larvae feed on the grape berries (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). Economically important hosts are identified below.

Major Hosts Abelmoschus esculentus (okra), Allium spp. (onion), Amaranthus spp., Apios spp. (groundnut), Arachis hypogaea (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 annuus (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 spp. (rose), Solanum lycopersicum (tomato), Saccharum officinarum (sugarcane), 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 (cashew), 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 domestica (apple), Mentha spp. (mint), Phoenix dactylifera (date palm), Pinus spp. (pine), and Zinnia spp. (zinnia).

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

106 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth

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, Ascension Island, Benin, Botswana, Burkina Faso, Burundi, Cameroon, Cape Verde, Central African Republic, Chad, Comoros, Congo, Democratic Republic of the Congo, Cote d’Ivoire, Egypt, Equatorial Guinea (including Bioko), Eritrea, Ethiopia, Gabon, Gambia, Ghana, Guinea, Kenya, Liberia, Libya, Madagascar, Malawi, Mali, Mauritania, Mauritius (Including Rodrigues), Morocco, Mozambique, Namibia, Niger, Nigeria, Reunion, Rwanda, Saint Helena, Sao Tome and Principe, Senegal, Seychelles, Sierra Leone, Somalia, South Africa, Sudan, Swaziland, Tanzania, Togo, Tunisia, Uganda, Zaire, Zambia, and Zimbabwe. Asia: Afghanistan, Bangladesh, Brunei, India, and Turkey. Europe: Albania, Greece (including Crete and Dodecanese), Italy (including Sardinia and Sicily), Malta, Portugal (including Azores and Madeira), and Spain (including Balearic Islands and Canary Islands). Middle East: Bahrain, Cyprus, Iran, Iraq, Israel, Jordan, Lebanon, Oman, Saudi Arabia, Syria, United Arab Emirates, and Yemen. Oceania: American Samoa and Fiji (CIE, 1964; Evenhuis, 2010; Fibiger and Skule, 2011; EPPO, 2012).

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 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 analysis by USDA-APHIS-PPQ-CPHST shows that portions of Alabama, Arkansas, Florida, Georgia, Louisiana, Mississippi, South Carolina, and Texas are at the greatest risk from S. littoralis. Portions of most states within the continental United States have low to moderate risk of S. littoralis establishment based on climate and host range. Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination. The trap is Plastic Bucket Trap. The lure is effective for 84 days (12 weeks).

The Lure Product Name is “Spodoptera littoralis Lure”.

107 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

Method Notes: This trap is also known as the unitrap. The trap has a green canopy, yellow funnel, and white bucket and is used with a dry kill strip. For instructions on using the trap, see Brambila et al. (2010).

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

Lure Notes: Place S. litura and S. littoralis lures in different traps and separate at least 20 meters (65 feet).

Though the lures for Spodoptera littoralis and S. litura are composed of the same two compounds (Z,E,9,11-14:AC and Z,E,9,12-14:AC), the compounds are loaded into the lure dispensers in different amounts depending on the target species. Therefore, it is necessary to use the specific lure for each of the two targets.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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 two weeks, but effectiveness declined after three to four weeks of use (Ahmad, 1988). To monitor male flight activity in vegetable production areas, delta traps were placed 1.7 m (5.58 ft.) above the ground at a rate of two traps/ha (approximately one trap/acre) (Ahmad, 1988). Pheromone lures impregnated with 2 mg of the pheromone blend (blend not specified) were replaced after four weeks of use (Ahmad, 1988). Traps are deployed at a similar height (1.5 m; 4.92 ft.) 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

108 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth 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 to 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. See the Field Diagnostics and Wing Diagnostics aids by Brambila (2008a, b) for additional Figure 4. Larva of S. exigua. Photo information courtesy of Oklahoma State (http://caps.ceris.purdue.edu/webfm_send/553 University. and http://caps.ceris.purdue.edu/webfm_send/554).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Observation of adult genitalia is often the only certain method to separate species.

Easily Confused Pests Figure 5. Adult of S. ornithogalli. S. littoralis is often confused with S. litura, and Photo courtesy of Mississippi the variability and similarity of the two species Entomological Museum. makes correct identification difficult; http://mothphotographersgroup.mss examination of adult genitalia is often the only tate.edu/Files/JV/JV50.7.shtml. certain method to separate the two species. For

109 Spodoptera littoralis Primary Pest of Grape Arthropods Egyptian cotton leafworm Moth

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 bursae are the same length in female S. littoralis, whereas they are different 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. 4), 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. 5), 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 (USDA, 1982).

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.

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.

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.

Brambila, J. 2008a. Egyptian Cottonworm, Field Diagnostics - Spodoptera littoralis. http://caps.ceris.purdue.edu/webfm_send/553.

Brambila, J. 2008b. Egyptian Cottonworm, Wing Diagnostics - Spodoptera littoralis. http://caps.ceris.purdue.edu/webfm_send/554.

Brambila, J., Jackson, L., and Meagher, R. 2010. Plastic bucket trap protocol. http://caps.ceris.purdue.edu/webfm_send/398.

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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.

CIE. 1964. Spodoptera littoralis (Boisd.). Distribution Maps of Pests, Series A (Agricultural). Map No. 232. Commonwealth Institute of Entomology.

El-Mezayyen, G.A., El-Dahan, A.A., Moawad, G.M., and Tadros, M.S., 1997. A modified light trap as a tool for insect 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.

EPPO. 2012. EPPO Plant Quarantine Information Retrieval System (PQR), version 5.0.5540. European and Mediterranean Plant Protection Organization.

Evenhuis, N.L. 2010. Checklist of Fijian Lepidoptera. Accessed April 12, 2012 from: http://hbs.bishopmuseum.org/fiji/checklists/lepidoptera.html.

Fibiger, M. and Skule, B. 2011. Fauna Europaea: Spodoptera littoralis. In O. Karsholt and E. J. Nieukerken (eds.) Fauna Europaea: Lepidoptera, Moths, version 2.4. Accessed April 10, 2012 from: http://www.faunaeur.org.

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. Descriptions and colour illustrations of 1183 species., [7+]273pp.; [col. frontis., 63 col. pl., 18 fig., 290 X 220 mm]; many ref.

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

111 Spodoptera littoralis Primary Pest of Grape 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.

112 Spodoptera litura Primary Pest of Grape 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 2009 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).

Eggs: Spherical, somewhat flattened, sculpted with approximately 40 longitudinal ribs, 0.4 to 0.7 mm (0.016 to0.028 in.) in diameter; pearly green, 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).

Larvae: Newly hatched larvae are tiny, blackish green with a distinct black band on the first abdominal segment. Fully grown larvae are stout and smooth with scattered short setae. Head shiny black, and conspicuous black tubercules each with a long hair on each segment. Color of fully grown larvae not constant, but varies from dark gray to dark brown, or black, sometimes marked with yellow dorsal and lateral stripes of unequal width. The lateral yellow stripe bordered dorsally with series of semilunar black marks. Mature larvae are 40 to 50 mm (1.57 to 1.97 in.). Two large black spots on first and eight abdominal segments (Hill, 1975; USDA, 1982; CABI, 2007).

113 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

Figure 1. Egg mass (left), larva (center), and adult (right). Photos courtesy of CABI, 2007.

Pupae: Reddish brown in color, enclosed inside rough earthen cases in the soil, 18 to 22 mm (0.71 to 0.87 in.) long, last abdominal segment terminates in two hooks (USDA, 1982; CABI, 2007).

Adults: Body whitish to yellowish, suffused with pale red. Forewings dark brown with lighter shaded lines and stripes. 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 to 18 mm (0.55 to 0.71 in.), wing span 28 to 38 mm (1.1 to 1.5 in.) (Hill, 1975; USDA, 1982).

See Schmutterer (1969), Cayrol (1972), and Brown and Dewhurst (1975) for additional information.

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 three to four days. A single female lays 1500 to 2500 eggs in about six to eight 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 (82.4 °F). 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, 1979). According to Yamanaka et al. (1975) the female continues to lay eggs in egg masses over a period of five days at 25°C (77°F).

114 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

Maximum fecundity for S. litura was observed at 27°C (80.6°F)under 12 hours per 24 hours of light (100 foot candle light) (Hasmat and Khan, 1977, 1978). Temperatures between 24 and 30°C (75.2 and 86°F) were also favorable for fecundity and fertility. At 33 and 39°C (91.4 and 102.2°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). Parasuram 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 (46.4°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 (51.44°F) threshold from emergence to oviposition. The upper developmental threshold temperature of all stages was 37°C (98.6°F); 40°C (104°F) was lethal.

Maheswara Reddy (1983) showed that the majority of mating occurred between 23.30 and 00.30 hrs 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 mating per males was 10.3 and per female was 3.1 (Ahmed et al., 1979). Ohbayashi et al. (1973) showed two peaks in mating behavior at 23.00 (3 hours after initiation of a dark period) and a minor peak at 3:00 (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 (4.72 in.) deep. Across soil types, most larvae pupated at a 4 cm (1.57 in.) 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).

115 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

Grape: Larvae scrape the leaf tissue and cause ‘drying of the leaves’ (Balasubramaniam et al., 1978). The larvae damage the growing berries and cause defoliation. Balikai et al. (1999) also showed that later instar larvae cut the rachis of grape bunches and petioles of individual berries during the night hours leading to fruit drop. During the day, the larvae move to the lower portion of the leaf vines and the crevices of the soil. The larvae use the main stem for climbing from the soil level during dusk.

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

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 and spider mites (Pillai et al., 1993).

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.

116 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

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, the 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. Economically important hosts are provided below.

Major Hosts Abelmoschus esculentus (okra), Acacia mangium (brown salwood), Allium cepa (onion), Amaranthus (grain amaranth), Arachis hypogaea (peanut), Beta vulgaris var. vulgaris (sugarbeet), Boehmeria nivea (ramie), Brassica spp., Brassica oleracea var. botrytis (cauliflower), Brassica oleracea var. capitata (cabbage), Camellia sinensis (tea), Capsicum frutescens (chilli), Castilla elastica subsp. elastica (castilloa rubber), Cicer arietinum (chickpea), Citrus spp., Coffea spp. (coffee), Colocasia esculenta (taro), Corchorus spp. (jutes), Corchorus olitorius (jute), Coriandrum sativum (coriander), Crotalaria juncea (sunn hemp), Cynara cardunculus (artichoke), Erythroxylum coca (coca), Fabaceae (leguminous plants), Foeniculum vulgare (fennel), Fragaria x 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 spp. (poppies), Paulownia tomentosa (paulownia), Phaseolus spp. (beans), Piper nigrum (black pepper), Poaceae (grasses), Psophocarpus tetragonolobus (winged bean), Raphanus sativus (radish), Ricinus communis (castor bean), Rosa spp. (roses), Sesbania grandiflora (agati), Solanum lycopersicum (tomato), Solanum melongena (aubergine, eggplant), Solanum tuberosum (potato), Sorghum bicolor (sorghum), Syzygium aromaticum (clove), Tectona grandis (teak), Theobroma cacao (cocoa), 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.

117 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

Known Distribution Asia: Afghanistan, Andaman Islands, Bangladesh, Bonin Islands, Brunei, Cambodia, China, Christmas Island, Cocos Islands, India, Indonesia, Iran, Japan, Korea, Laos, Lebanon, Malaysia, Maldives, Myanmar, Nepal, Nicobar Islands, Oman, Pakistan, Philippines, Singapore, Sri Lanka, Syria, Taiwan, Thailand, and Vietnam. Europe: France. Russia. Africa: Ghana and Reunion. North America: United States (Hawaii). Oceania: American Samoa, Australia (including Tasmania), Austral Islands, Belau, Caroline Islands, Cook Islands, Federated states of Micronesia, Fiji (including Rotuma), French Polynesia (including Marquesas Islands and Tuamotus), Guam, Kiribati (including Line Islands), Mariana Islands, Marshall Islands, New Caledonia (including Loyalty Islands), New Zealand (including Kermadec Islands), Niue, Norfolk Island, Northern Mariana Islands, Palau, Papua New Guinea, Phoenix Islands, Pitcairn Islands (including Henderson Island), Samoa, Society Islands, Solomon Islands, Tonga, Tuvalu, Vanuatu, Wake Island, and the Wallis and Futuna Islands (CABI, 1993; Obeng-Ofori and Sackey, 2003; EPPO, 2012).

This species was found in 2010 in the United Kingdom and is considered transient and under eradication (EPPO, 2012).

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, California, 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 (precluded) in many areas of the United States.

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination. The trap is Plastic Bucket Trap. The lure is effective for 84 days (12 weeks).

The Lure Product Name is “Spodoptera litura Lure”.

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

Method Notes: This trap is also known as the unitrap. The trap has a green canopy, yellow funnel, and white bucket and is used with a dry kill strip. For instructions on using the trap, see Brambila et al. (2010).

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

118 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

Lure Notes: Place S. litura and S. littoralis lures in different traps and separate at least 20 meters (65 feet).

Though the lures for Spodoptera littoralis and S. litura are composed of the same two compounds (Z,E,9,11-14:AC and Z,E,9,12-14:AC), the compounds are loaded into the lure dispensers in different amounts depending on the target species. Therefore, it is necessary to use the specific lure for each of the two targets.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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 (6.56 ft.) above the ground level caught significantly more male S. litura than those placed at higher of lower heights (ranging from 0.5 to 4.0 m (1.64 to 13.1 ft.). Ranga Rao et al. (1991) suggest trap placement at 1 m (3.28 ft.).

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 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).

119 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of S. litura is by morphological identification. S. litura is difficult to distinguish 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).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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/pests-and-diseases/Pest/Main/136292.

Easily Confused Pests 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.

Balasubramanian, M., Murugesan, S., and Parameswaran, S. 1978. Beware of cutworm on grapes. Kisan World 5(6): 51-52.

120 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

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.

Brambila, J., Jackson, L., and Meagher, R. 2010. Plastic bucket trap protocol. http://caps.ceris.purdue.edu/webfm_send/398.

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. 2003. Distribution maps of pests, Spodoptera litura (Fabricius) Lepidoptera: Noctuidae. Series A: Map No. 61.

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.

EPPO. 2012. EPPO Plant Quarantine Information Retrieval System (PQR), version 5.0.5540. European and Mediterranean Plant Protection Organization.

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.

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

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.

121 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

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.

Obeng-Ofori, D. and Sackey, J. 2003. Field evaluation of non-synthetic insecticides for the management of insect pests of okra Abelmoschus esculentus (L.) Moench in Ghana. Ethiop. J. Sci. 26(2): 145-150.

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.

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 longetivity 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.

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

122 Spodoptera litura Primary Pest of Grape Arthropods Rice cutworm Moth

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

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].

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.

123 Adoxophyes orana Secondary Pest of Grape Arthropods Summer fruit tortrix Moth

Secondary Pests of Grape (Truncated Pest Datasheet)

Adoxophyes orana

Scientific Name Adoxophyes orana Fischer von Roeslerstamm

Synonyms: Adoxophyes reticulana, Capua reticulana, Cacoecia reticulana, Capua orana, Tortrix ornana, Tortrix reticulana, Capua congruana, Adoxophyes tripsiana, Adoxophyes fasciata, Adoxophyes congruana, Acleris reticulana.

Common Name Summer fruit tortrix, reticulated tortrix, apple peel tortricid

Type of Pest Moth

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

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2006 through 2012

Pest Description Eggs: Yellowish and deposited in masses (Fig. 1). After hatching, the transparent egg shells remain present.

Larvae: (Fig. 1) Greenish with light hairs and warts. The head is light brown to yellow (sometimes somewhat spotted) as is the thoracic shield and the anal shield. The anal comb is very fine and long with light colored Figure 1. Eggs, larva, and adult A. teeth. The thoracal legs are brown to black. orana (female top, male bottom). The head is long and wide. Abdominal and Photos courtesy of R. Coutin/OPIE. anal prolegs are greenish.

124 Adoxophyes orana Secondary Pest of Grape Arthropods Summer fruit tortrix Moth

Pupae: The pupae of A. orana are initially light brown, but become dark brown towards the time of emergence of the adult moth. The length is between 8 and 11 mm (0.31 to 0.43 in.). The posterior margin of abdominal segments two to eight of the pupae contains very small bristles. These bristles cannot be distinguished with a regular magnifying glass and are hence visible as a line. The specific fork-shape of wing veins seven and eight is already visible in the pupal stage.

Adults: A very specific characteristic of A. orana is the fork-shaped structure of the wing veins seven and eight. The forewing of the female is rather dull grayish brown, while in the male the coloration is brighter and is a yellowish brown (Fig. 1). Male wingspan is 15 to 19 mm (0.59 to 0.75 in.); while female is18 to 22 mm (0.71 to 0.87 in.). Sexual dimorphism is pronounced; antenna of male shortly ciliate, forewing with broad costal fold from base to about one-third, markings usually conspicuous, contrasting with paler ground color; female usually larger, antenna minutely ciliate, forewing without costal fold, with darker general coloration and less contrasting markings (Bradley et al., 1973).

Symptoms/Signs External feeding will be visible on leaves and fresh growth of twigs. Feeding will deform leaves and create areas with necrosis (dead tissue). Leaves may appear wilted, yellow, shredded, or dead. Leaves are likely to be rolled or folded and held together with silk webbing. Feeding on new growth of twigs will Damage to apple epidermis leave lesions. If the insect is feeding in Figure 2. showing “gnawed” appearance (top) flowers, external feeding damage and silk and damage to pear foliage and fruit webbing will be evident. In all areas where the (bottom). Photos courtesy of R. insect has fed, frass should also be visible. Coutin/OPIE.

Summer generation larvae feed extensively and severely damage fruit (Fig. 2). Feeding on fruits or pods causes scabs or pitting, and frass may be present. On fruit crops, larvae prefer to feed sheltered under a leaf bound to fruit and silk.

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination.

125 Adoxophyes orana Secondary Pest of Grape Arthropods Summer fruit tortrix Moth

Any of the following Trap Product Names in the IPHIS Survey Supply Ordering System may be used for this target: 1) Paper Delta Trap, 2 sticky sides, Brown 2) Paper Delta Trap, 2 sticky sides, Green 3) Paper Delta Trap, 2 sticky sides, Orange

The Lure Product Name is “Adoxophyes orana Lure.” The lure is effective for 84 days (12 weeks).

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

Method Notes: Trap should be used with ends open. Trap color is up to the State and does not affect trap efficacy.

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

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Trapping: Several monitoring techniques have been developed and applied to A. orana. The most effective approach involves sex pheromone- baited traps. The sex pheromone is a blend of (Z)-9-tetradecenyl acetate and (Z)-11-tetradecenyl acetate. These two compounds are most attractive to males in a 9:1 blend of (Z)-9:(Z)-11 isomers; E- isomers of either compound had a strong inhibitory effect (Davis et al., 2005). The 9:1 pheromone blend is available commercially as Adoxomone (Murphy PheroconTM Summer Fruit Tortrix Moth Attractant) for use with Pherocon 1C traps [Zoecon Corp].

Visual survey: Visual sampling and beat sampling may also be used to inspect plants for eggs and larvae. Eggs may be observed on the stems and leaves; late instars may be found in the crown on new shoot growth; and pupal cocoons may be found in leaves, on stems, or in mummified pods/seeds. Both methods are time consuming. Visual sampling or beat sampling are not commonly recommended (Davis et al., 2005).

Not recommended: As an alternative to pheromone traps, Robinson light traps with 125W mercury vapor bulbs, 125 W black light bulbs, or 100W flood lights can be used. While sex pheromone traps attract males of a targeted species, light traps non- selectively draw in many flying insects.

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 have a low to moderate risk rating for A. orana establishment based on host availability and climate within the continental United States. Areas of the southeastern

126 Adoxophyes orana Secondary Pest of Grape Arthropods Summer fruit tortrix Moth

United States, California, Illinois, Indiana, Maryland, Missouri, Oklahoma, and Texas have the highest risk of A. orana establishment.

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation requires a morphological identification. Adoxophyes orana may occur in mixed populations with other morphologically similar species, including other Adoxophyes species. Final identification is by dissection of male genitalic structures.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Because of their very secretive nature, leafrollers are difficult to detect. Distinguishing between males and females of adult Adoxophyes is difficult in general (Balachowsky 1966). According to Yasuda (1998), the extensive color and pattern variation of the forewing and morphological resemblance among Adoxophyes species have created difficulties in the identification of the species. A. orana very closely resembles two U.S. species, Adoxophyes furcatana and A. negundana, but there are slight differences in male genitalia. Any identification should be confirmed by an appropriately trained entomologist.

References Bradley, J.D., Tremewan, W.G., and Smith, A. 1973. British Tortricoid moths. Cochylidae and Tortricidae: Tortricinae. Ray Society, London.

Davis, E.E., French, S., and Venette, R.C. 2005. Mini Risk Assessment – Summer Fruit Tortrix Moth, Adoxophyes orana (Fischer von Roslerstamm, 1834) [Lepidoptera: Tortricidae]. http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/aoranapra.pdf.

127 Aleurocanthus spiniferus Secondary Pest of Grape Arthropods Orange spiny whitefly Whitefly

Aleurocanthus spiniferus

Scientific Name Aleurocanthus spiniferus Quaintance

Synonyms: Aleurocanthus citricolus, Aleurocanthus rosae, Aleurocanthus spiniferus var. intermedia Aleurodes citricola, and Aleurodes spinifera.

Common Name Orange spiny whitefly, citrus mealy wing

Type of Pest Whitefly

Taxonomic Position Class: Insecta, Order: Hemiptera, Family: Aleyrodidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2006 through 2009; Additional Pests of Concern List- FY2012

Pest Description Aleurocanthus spiniferus is a whitefly pest that is not known to occur in the continental United States. The pest has been reported to occur in Guam and Hawaii. Although A. spiniferus is primarily a pest of citrus and rose, it has been reported to occur on other hosts including peach (Prunus persica), black cherry (Prunus serotina), pear (Pyrus spp.), grape (Vitis vinifera), and guava (Psidium spp.) (EPPO, 1997; Jeffers, 2009; Figure 1. A. spiniferus eggs and Gyeltshen et al., 2011). nymphs. Nymphal instars 1, 2, and 4 (pupae, 1.2 mm in length). The Whiteflies have six developmental stages: egg, white, waxy filaments are typical of crawler (1st instar), two sessile nymphal instars the species. Photo courtesy of nd (2 and 3rd instars), the pupa (4th instar) and CABI, 2009. adult. Identification of the Aleyrodidae is largely based upon characters found in the pupal (4th instar) stage (Gyeltshen et al., 2011).

Eggs: The elongate-oval eggs (0.2 mm; 0.008 in. long) are yellow when first laid and then darken to charcoal gray or black; each is attached to the leaf by a short pedicel.

128 Aleurocanthus spiniferus Secondary Pest of Grape Arthropods Orange spiny whitefly Whitefly

Larvae: The six-legged, dusky, elongate first-instar larvae [0.3 x 0.15 mm (0.01 x 0.006 in.] have two long and several shorter, slender dorsal glandular spines. All subsequent immature stages are sessile, have non-functional leg stubs, and possess numerous, dark dorsal spines on which a stack of exuviae of earlier instars may occur. The second instar [0.4 x 0.2 mm (0.016 x 0.008 in.)] is a dark brown to charcoal convex disc with yellow markings, while the third instar [0.87 x 0.74 mm (0.034 x 0.029 Figure 2. Adult A. spiniferus. Photo in.)] is usually black with a rounded, courtesy of M.A. van den Berg. greenish spot on the anterior part of the www.bugwood.org. abdomen and obvious dorsal spines. In the fourth immature stage or 'pupa', females are larger (1.25 mm (0.05 in.) long) than males (1 mm (0.04 in.) long). This stage is black, has numerous dorsal spines, and is often surrounded by a white fringe of waxy secretion (Fig. 1). This is the stage required for identification purposes.

Adults: Winged; the females (1.7 mm (0.07 in.) long) are larger than the males (approximately 1.33 mm (0.05 in.) long). The wings are dark gray at ecdysis (Fig. 2), sometimes developing a metallic blue-gray sheen later; lighter markings on the wings appear to form a band across the insect. The body is orange to red initially; the thorax darkens to dark gray in a few hours. The limbs are whitish with pale yellow markings. Figure 3. Sooty mold on citrus Symptoms/Signs leaves. Photo courtesy of M.A. van Both adults and larvae damage plants. den Berg. www.bugwood.org.

Primarily, orange spiny whitefly affects host plants by sucking the sap but it also causes indirect damage by producing honeydew and subsequently promoting the growth of sooty mold. Sticky honeydew deposits accumulate on leaves and stems and usually develop black sooty mold fungus (Fig. 3), giving the foliage (even the whole plant) a sooty appearance. Sooty mold can lead to reduction of respiration and photosynthesis interfering with normal leaf function (USDA, 1982). In heavy infestations, sooty mold can develop on the fruit, lowering the quality (USDA, 1982). Ants may be attracted by the honeydew.

Inspect for spiral egg masses and larvae on the underside of leaves. The colorful adult may be found on tender terminal growth (USDA, 1982). Infested leaves may be

129 Aleurocanthus spiniferus Secondary Pest of Grape Arthropods Orange spiny whitefly Whitefly distorted. The insects are most noticeable as groups of very small, black spiny lumps on leaf undersides.

Heavy infestations can lead to loss of vitality and eventually tree mortality if left unchecked (USDA, 1982).

Survey CAPS-Approved Method*: The CAPS-approved survey method is a visual survey.

Note: A trap and lure combination has been described in the literature but this method has not been fully evaluated. A sex pheromone has been identified for this species and is used in conjunction with a yellow sticky card for control and monitoring in China (Wang, personal communication; Zhang et al., 2010). The lure components are not described in the literature and the only sources for the lure are Chinese companies (Wang, personal communication). This lure is not currently available for purchase through the PPQ Trap and Lure Ordering Database. The pheromone is being investigated by other PPQ programs. More information will be posted here when it is available.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Surveys should be focused where the greatest risk for establishment occurs. Based on the current distribution of A. spiniferus in Africa, Asia, Europe, North America, and Oceania, the pest may be able to establish in U.S. Plant Hardiness Zones 8 to 12 (Jeffers, 2009). The pest could possibly become a pest in heated glasshouses in other areas in the United States (EPPO, 1997). A recent host analysis by USDA-APHIS-PPQ- CPHST indicates that many states in the United States have a low to moderate risk from A. spiniferus establishment based on host availability. Areas of Texas and Florida, however, have the greatest risk for establishment. Alabama, Arkansas, Georgia, Louisiana, Mississippi, have the greatest risk of A. spiniferus establishment.

Trapping: In China, A. spiniferus is trapped for control purposes using a sex pheromone and yellow sticky card (Wang, personal communication; Zhang et al., 2010).

Visual: Aleurocanthus spiniferus is most often found on citrus and roses. Examine plants, especially shrubs or trees, closely for signs of sooty mold or sticky honeydew on leaves, stems, and fruit, or for ants running about. A heavy infestation gives trees an almost completely black appearance. Sooty mold can reduce respiration and photosynthesis of plants and can cause plants and fruit to become unsightly and unsalable (EPPO, 1997). Foliage that is badly contaminated may drop while fruit set may be reduced with heavy infestations (EPPO, 1997). Look for distorted leaves with immature stages of A. spiniferus on the undersides. Larvae often form dense colonies with up to several hundred larvae on one leaf (EPPO, 1997).

130 Aleurocanthus spiniferus Secondary Pest of Grape Arthropods Orange spiny whitefly Whitefly

Leaves infested by A. spiniferus are mainly found on the lower parts of the host plants with infested leaves tending to be aggregated on the plant (EPPO, 2002). Good light conditions are essential for detection; in poor light, a powerful flashlight is helpful. A large hand lens may be necessary to aid in recognition of the dorsal spines on immature stages (CABI, 2009).

Surveys may be carried out at any time of the year, however adults will not be found during winter (USDA, 1982). Adults may periodically be found on the terminal growth (USDA, 1982), and are active fliers when disturbed (EPPO, 1997).

Movement by this species is dependent on movement of host plants as adult flight is not a major means of long-range dispersal (EPPO, 1997). Spread can occur on movement of nursery stock and infested fruits (Gyeltshen et al., 2011). This species has been intercepted several times over the years at United States ports of entry. Most interceptions occurred on host plant leaves in baggage (AQAS, queried May 5, 2011).

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of A. spiniferus requires a morphological identification. A. spiniferus is similar to many other species within the Aleurocanthus genus. Several similar species of Aleurocanthus also occur on citrus, including A. citriperdus and A. woglumi. These species differ from each other only in microscopic characters of the “pupa” (fourth instar) and require expert preparation and identification to distinguish them reliably. The main characteristic difference between orange spiny whitefly and citrus black fly, A. woglumi that can be observed in the field, is that the white wax fringe that surrounds their pupal case margins is generally twice as large for the orange spiny whitefly.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: A diagnostic protocol for positively identifying A. spiniferus through examination of the pupal case can be found in EPPO (2002).

Molecular diagnostics to differentiate A. spiniferus from closely related whitefly species have recently been studied (Xun et al., 2009).

Additional images are available at http://www.eppo.org/QUARANTINE/aleurocanthus_spiniferus_IT/first_record.htm.

Easily Confused Pests A. spiniferus is similar to many other species within the Aleurocanthus genus.

References

131 Aleurocanthus spiniferus Secondary Pest of Grape Arthropods Orange spiny whitefly Whitefly

AQAS. 2011. Total interception data for Aleurocanthus spiniferus. Agricultural Quarantine Activity Systems. Accessed May 5, 2011 from: https://mokcs14.aphis.usda.gov/aqas/HomePageInit.do.

CABI. 2009. Crop protection compendium: global module. Commonwealth Agricultural Bureau International, Wallingford, UK. http://www.cabi.org/compendia/cpc/.

Gyeltshen, J., Hodges, A., and Hodges, G.S. 2011. Orange spiny whitefly, Aleurocanthus spiniferus (Quaintance) (Insecta: Hemiptera: Aleyrodidae). http://edis.ifas.ufl.edu/pdffiles/IN/IN61800.pdf or http://entomology.ifas.ufl.edu/creatures/citrus/orange_spiny_whitefly.htm.

EPPO. 1997. Data Sheets on Quarantine Pests: Aleurocanthus spiniferus. European and Mediterranean Plant Protection Organization (EPPO). http://www.eppo.org/QUARANTINE/insects/Aleurocanthus_spiniferus/ALECSN_ds.pdf.

EPPO. 2002. Diagnostic Protocols for Regulated Pests, Aleurocanthus spiniferus. EPPO Bulletin 32(2): 255-259.

Jeffers, L. 2009. New Pest Advisory Group Report. Aleurocanthus spiniferus Quaintance: Orange Spiny Whitefly Hemiptera/ Aleyrodidae. http://www.aphis.usda.gov/plant_health/cphst/npag/downloads/SampleReports/Aleurocanthus- spiniferusReport.pdf.

Martin, J.H. 1987. An identification guide to common whitefly pest species of the world (Hompotera: Aleyrodidae). International Journal of Pest Management 33/34: 298-322.

USDA. 1982. Pests Not Known to Occur in the United States or of Limited Distribution, No. 14: Orange spiny whitefly.

Wang, B. Pheromone for Aleurocanthus spiniferus. Personal communication to L. D. Jackson on June 3, 2011 from B. Wang (USDA-APHIS-PPQ-CPHST).

Xun, L., FangHao, W., and GuiFen, Z.. 2009. SCAR marker for rapid detection of the spiny whitefly, Aleurocanthus spiniferus (Quaintance) (Homoptera: Aleyrodidae) [abstract]. Acta Entomologica Sinica 52 (8): 895-900.

Zhang, Y-D., He, Z.-Q., Liu, Y.-H., Mao, J.-H. and Wang, H. 2010. Primary research on effect of application of sex pheromone clad plate to control Aleurocanthus spiniferus in organic tea garden [abstract]. Southwest China Journal of Agricultural Sciences 23(6). (Abstract).

132 Eudocima fullonia Secondary Pest of Grape Arthropods Fruit piercing moth Moth

Eudocima fullonia

Scientific Name Eudocima fullonia Clerck

Synonyms: Othreis fullonia, Noctua dioscoreae, Ophideres fullonia, Ophideres obliterans, Ophideres princeps, Othreis phalonia, Othreis pomona, Phalaena fullonica, Phalaena fullonica, Phalaena phalonia, and Phalaena Pomona.

Common Names Fruit piercing moth, fruit-sucking moth

Type of Pest Moth

Taxonomic Position Class: Insecta Order: Lepidoptera Family: Noctuidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2006 through 2012

Pest Description This species can be a major pest of Citrus spp. in many areas, but it will also attack other fruits including banana, fig, guava, kiwifruit, longan, lychee, mango, stone fruit, persimmon and ripening papaw (Astridge and Fay, 2005).

Eggs: Hemispherical, just over 1 mm (0.04 in.) in diameter, and greenish-white to creamy-yellow when laid. Delicate surface sculpturing can be seen with the aid of a microscope. The brownish head capsule of the developing larva becomes obvious beneath the shell a few hours before hatching.

Larvae: The newly hatched larvae are 4 to 5 mm (0.16 to 0.2 in.) long, a bright translucent green in color, and Figure 1. Larva of E. fullonia. Photo inconspicuously banded by brown spots and courtesy of CABI, 2009. hairs. The head capsule is 0.5 mm (0.02 in.) wide. Second instars are a uniform dull black, with two developing, paired, lateral orange eyespots. Larvae molt four or five times during development. Final instars can reach about 60 mm (2.36 in.) in length, with a head capsule of 4.5 mm (0.18 in.). Mature larvae are a velvety brown to black (Fig. 1) or pale yellow to green. There are

133 Eudocima fullonia Secondary Pest of Grape Arthropods Fruit piercing moth Moth fine powdery white spots along the entire length of the body and two conspicuous, paired, lateral eyespots on the second and third abdominal segments. In dark larvae, the eyespots are peripherally white (above) and orange (below), with a central black area surrounding a pale blue core. When resting, larvae hold the posterior part of the body upwards, while the anterior part is curled with the head tucked under (Fig. 1). If disturbed, larvae may rear and 'spit' digestive juices. Larvae move with a semi-looping action.

Pupae: The post-feeding larva forms a silken cocoon among the leaves of the larval host plant and attaches itself within the cocoon at the anal end. The pupa is about 30 mm (1.18 in.) long, a glistening Figure 2. Adult female fruit piercing brown-black, and can be sexed at this moth. Photo courtesy of CABI, 2009. stage using differences in the position of the genital grooves.

Adults: Adults have 7 to 10 cm (2.76 to 3.94 in.) wingspans, with mottled brown, gray, green, or silvery white forewings. The color patterns of the forewings are sexually dimorphic. Males have leaf-like forewings of red-brown to purplish-brown. There is an inconspicuous, irregular spot centrally placed near the anterior margin. In females, the forewings are more variegated and striated than in males. The color varies between purplish-brown and grayish-ochre, often flecked with green and white. There is a distinct dark, roughly triangular mark in a similar position to the spot in the male forewing. The hind wings are characterized by the orange-yellow color, extensively bordered by a black and hatched area and a central black mark (kidney shaped or round). These are often exposed when the moth is feeding. The thorax is a purplish-brown and the abdomen orange-yellow. An individual moth can spend several hours feeding Figure 3. Male (left) and female (right) E. fullonia on a green mandarin from the one fruit, but would generally attack a number of fruit on a single night. fruit. Photo courtesy of CABI, 2007.

Adults rest with the forewings held tent- like over the body. When feeding, the forewings are held out exposing the bright hindwings (Fig. 3).

Symptoms/Signs For most moth pests, the larvae are the damaging stage. The fruit piercing moth differs in this aspect, because it is the adult moth that is the damaging stage, and

134 Eudocima fullonia Secondary Pest of Grape Arthropods Fruit piercing moth Moth

the larvae are essentially not harmful. The mouth parts of the moth are about 2.5 cm (1 in.) long and strong enough to penetrate through tough-skinned fruit. Once the moth has punctured the skin of the fruit, a process that usually takes a few seconds, it feeds upon the juices of the fruit (Fig. 3). The proboscis is pushed deeper as the juices become exhausted in that particular site; normally only ripened or well matured fruits are attacked unless these are unavailable (Baptist, 1944). Feeding occurs at night. Fruit flesh damaged by this moth becomes soft and mushy differing from fruit damaged by fruit flies, which is more liquid. On pear, Cave and Lightfield (1997) state that E. fullonia is considered an external feeder.

A round, pinhole-sized puncture is made in fruits. The hole serves as an entry point for pathogens and can result in early fruit drop. The latter is an obvious sign of fruit piercing moth activity in citrus. A small cavity is left in the fruit in the feeding site. The area of the fruit around the cavity will be dry and spongy. It may be bruised beneath the skin (Astridge and Fay, 2005). The fruit piercing moth is a known vector of Oospora citri, a fungus that rots the fruit and has a penetrating odor that attracts this moth. Other microorganisms that Figure 4. A damaged fruit showing gain entrance into the fruit and cause fruit rot around the piercing moth rotting include Fusarium spp., feeding site. Photo courtesy of CABI, Colletotrichum spp., and several types of 2009. bacteria. When moths are abundant, green fruit is attacked, causing premature ripening and dropping of fruits. On oranges, a green fruit turns yellow at the site of the piercing and fungi soon develop within the wound.

Survey CAPS-Approved Method*: The CAPS-approved survey method is visual survey.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Surveys should be focused where the greatest risk for establishment occurs. A recent host analysis by USDA-APHIS-PPQ-CPHST indicates that most states in the United States are at low risk for E. fullonia establishment based strictly on host availability. Portions of California, Florida, Maryland, Massachusetts, New Jersey New York, Pennsylvania, Vermont, and West Virginia have a moderate risk, however.

135 Eudocima fullonia Secondary Pest of Grape Arthropods Fruit piercing moth Moth

Visual survey: Adult moths are likely to be found on mature fruit several weeks before harvest. The most effective way to monitor for fruit piercing moths is to inspect the crop by flashlight after sundown beginning a few weeks before harvest (Davis et al., 2005). Moths are most active in the first few hours of the night. The large, red-glowing eyes of the moths are easily seen. Check trees/vines in the two outer rows of an orchard, particularly on the leeward side. Most damage occurs in the peripheral rows. Surveys were typically initiated 30 minutes after sundown and lasted one hour. Foliage of host plants may be inspected for larvae and other life stages (Davis et al., 2005).

In some fruits, such as lychees, detection of fruit piercing moth damage can be difficult. The slightest sign of weeping can be an indication, and when the fruit is squeezed, the juice will squirt out. The damage site will be flaccid and flattened in appearance and lack the firm, rounded flesh of intact fruit. Many farmers opt to place freshly picked fruit in a cool store at high humidity, which facilitates detection of damage after one day.

Survey site and selection: Areas where host material is found should be targeted for surveys; these can include orchards, nurseries, residential areas and other areas where host plants are used as ornamentals.

Time of year to survey: In Fiji, generations are continuous throughout the year (Martin Kessing and Mau, 1993). The larval populations increase in June and peak in August (Davis et al., 2005). After this, adults emerge with females laying eggs from June to October (Kumar and Lal, 1983).

Trapping: No pheromones or semiochemicals have been identified for E. fullonia. Combinations of attractants have been developed and incorporated into sugared-agar baits. Field trials with a range of these experimental baits in Clementine and hybrid mandarins in north Queensland showed many more attacks on the best baits (85% of moths) than on fruit up to the early harvest phase. Baits were less competitive, but still very attractive, when nearly all fruit reached the ripe stage. Recent studies have resulted in the selection of a suitable toxicant to incorporate in the baits, while the means to present the attractants so that they have a practical field life are still being pursued.

In an experiment by Reddy et al. (2007), researchers found that E. fullonia was significantly more attracted to fruit puree with agar and phytogel than to fruit puree with agarose. Fruit baits were ranked with banana being the most preferred, followed by banana and orange. These were significantly more attractive than kiwi, apple, pineapple, pear, papaya, mango, grapefruit, tomato, and green grape. The least attractive were star fruit, plum, and sour sop which were no more attractive than the controls (Reddy et al., 2007).

This species is attracted to ultraviolet lights making blacklight traps a potential trapping method (Martin Kessing and Mau, 1993). However, this method is not specific to this species.

136 Eudocima fullonia Secondary Pest of Grape Arthropods Fruit piercing moth Moth

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of E. fullonia requires a morphological identification.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Confirmation of E. fullonia requires a morphological identification.

A short description of the adult can be found in Baptist (1944). Maddison (1982) also has short descriptions of the life stages.

Easily Confused Pests Adults of E. fullonia closely resemble species such as Eudocima homaena and Eudocima jordani. All species of Eudocima cause similar damage. Separation of species involves detailed microscopic examination. Fruit discoloration could be attributed to fruit fly damage, but the size of the hole left at the damage site will clarify whether fruit-piercing moths were involved.

References Astridge, D. and Fay, H. 2005. Fruit piercing moths in rare fruit. Queensland Government, Department of Primary Industries and Fisheries.

Baptist, B.A. 1944. The fruit-piercing moth (Othreis fullonica L.) with special reference to its economic importance. The Indian Journal of Entomology 6(1/2): 1-13.

Cave, G.L. and Lightfield, J.W. 1997. Importation of fragrant and Ya pear fruit from China into the United States, a supplemental pest risk assessment. USDA-APHIS.

Davis, E.E., French, S., and Venette, R.C. 2005. Mini Risk Assessment- Fruit Piercing Moth: Eudocima fullonia Green [Lepidoptera: Noctuidae]. http://www.aphis.usda.gov/plant_health/plant_pest_info/pest_detection/downloads/pra/efulloniapra.pdf.

Kumar, K. and Lal, S.N. 1983. Studies on the biology, seasonal abundance, and host-parasite relationship of fruit sucking moth Othreis fullonia (Clerck) in Fiji. Fiji Agricultural Journal 45: 71-77.

Maddison P.A, 1982. Fruit-piercing moth. South Pacific Commission Advisory Leaflet, 14. New Caledonia: South Pacific Commission.

Martin Kessing, J.L. and Mau, R.F. 1993. Othreis fullonia (Clerk) Pacific fruit-piercing moth. University of Hawaii Department of Entomology, Honolulu, Hawaii. http://www.extento.hawaii.edu/kbase/crop/Type/othreis.htm.

Reddy, G.B.P., Cruz, Z.T., and Muniappan, R. 2007. Attraction of fruit-piercing moth Eudocima phalonia (Lepidoptera: Noctuidae) to different fruit baits. Crop Protection 26: 664-667.

137 Eutetranychus orientalis Secondary Pest of Stone Fruit 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 – 2006 through 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 (0.006 in.) Figure 1. Eggs (left) and adult (right ) of E. orientalis. (Avidov and Harper, Photos courtesy of Pedro Torrent Chocarro. 1969).

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

138 Eutetranychus orientalis Secondary Pest of Stone Fruit Arthropods Citrus brown mite Mite

reddish brown. The protonymph is pale-brown to light-green, with legs shorter than the 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; two pairs of anal setae; three 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.

139 Eutetranychus orientalis Secondary Pest of Stone Fruit Arthropods Citrus brown mite Mite

appear to be covered in a 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.

CAPS-Approved Method*: Visual survey is the method to survey for E. orientalis.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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 in most states in the continental United States pest establishment is unlikely. Risk for E. orientalis establishment based on climate and host availability is low in Arizona, California, and Nevada, Texas, and Utah. Risk is low to moderate in Florida.

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.

Hall (1992) discusses sampling strategies for spider mites in orange groves. The author’s sampling method consisted of examining 16 leaves per tree, five trees within a small area of trees, and three 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 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

140 Eutetranychus orientalis Secondary Pest of Stone Fruit Arthropods Citrus brown mite Mite

when processed the next day. Dhorria and Butani (1984) collected forty random leaves (10 leaves per tree) from each almond 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.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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.

Walter et al. (1995) provide a key to differentiate known Australian species of Eutetranychus, including E. orientalis.

Easily Confused Pests 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 (Tetranychus urticae) 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.

141 Eutetranychus orientalis Secondary Pest of Stone Fruit Arthropods Citrus brown mite Mite

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.

Walter, D.E., Halliday, R.B., and Smith, D. 1995. The Oriental Red Mite, Eutetranychus orientalis (Klein) (Acarina: Tetranychidae), in Australia. J. Aust. Ent. Soc. 34: 307-308.

142 Oxycarenus hyalinipennis Secondary Pest of Grape Arthropods Cottonseed bug True bug

Oxycarenus hyalinipennis

Scientific Name Oxycarenus hyalinipennis Costa

Synonyms: Aphanus hyalinipennis and Aphanus tardus var. hyalipennis

Common Name(s) Cotton seed bug, cotton stainer, dusty cotton stainer, dusky cotton bug, dusky cottonseed bug, Egyptian cotton seed bug

Type of Pest Bug

Taxonomic Position Class: Insecta, Order: Hemiptera, Family: Lygaeidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2008 through 2012

Pest Description Eggs: Oval 0.28 x 0.95 mm (0.01 x 0.04 in.), longitudinally striated, pale yellow becoming pink.

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.

Adults: (Fig. 1) Newly emerged individuals are pale pink, but rapidly turn black. Length of male about 3.8 mm (0.15 in.); female 4.3 mm (0.17 in.). Male abdomen terminates in round lobe, while the female’s is truncate. The Figure 1. Oxycarenus hyalinipennis insects have three tarsal joints and a pair of adult, dorsal and side view. Photos ocelli. Second antennal segments are usually courtesy of Natasha Wright, Florida in part pale yellow. Hemelytra hyaline and Department of Agriculture and usually whitish; clavus, base of corium, and Consumer Services, costal vein more opaque than rest. Setae of www.bugwood.org. three different types: More or less erect, stiff

143 Oxycarenus hyalinipennis Secondary Pest of Grape Arthropods Cottonseed bug True bug setae, which are blunt at tip and terminate in four to seven small teeth; normal, straight, tapering setae; and very thin, curved, flat-lying setae (USDA, 1983). 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. On cotton, the lint in which the bugs have been present is stained pinkish, sometimes with a trace of green, and contaminated with crushed fragments of the insect. Cotton seeds appear undamaged on the outside; internally, the embryos are shriveled and discolored (USDA, 1983).

O. hyalinipennis has been reported on several other hosts, including stone fruit, 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).

O. hyalinipennis has been observed sucking the fruits of grapes (Avidov and Harpaz, 1969) as well as several types of fruit trees (plum, pear) (USDA, 1983). In Israel, O. hyalinipennis has been recorded causing damage to dates, figs, avocado, and persimmon (Nakache and Klein, 1992). Avidov and Harpaz (1969) state that infestations in Israel can occur in apricot, peach, and persimmon, and to a lesser extent in apple, pear, quince, and grapevine. Damage can be due to feces, pungent odors (caused by crushing of adults or nymphs) or toxic saliva (Avidov and Harpaz, 1969; Nakache and Klein, 1992; Sweet, 2000). The feeding damage can appear as greasy spots that exude light colored gum (Avidov and Harpaz, 1969).

Larvae are only known to complete development if a host plant within the order Malvales is present (USDA-APHIS, 2010). Fruits are not known to be true hosts, but they can still be damaged by adults looking for moisture (USDA-APHIS, 2010). Sweet (2000) suggests that O. hyalinipennis feeds on these other plants to obtain moisture so the true hosts will not be damaged by their toxic saliva. This way, the true hosts will continue to develop more valuable food seeds later on (Sweet, 2000).

Survey CAPS-Approved Method*: Visual survey is the approved method to survey for O. hyalinipennis.

The CPHST Pest Datasheet includes a detailed visual survey protocol for this pest in cotton. This information is also available in the cotton commodity-based reference manual.

144 Oxycarenus hyalinipennis Secondary Pest of Grape Arthropods Cottonseed bug True bug

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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. Interceptions of O. hyalinipennis have occurred on fruits and vegetables including apple, avocado, corn, dates, figs, grapes, peach, okra, pineapple, and pomegranate, as well as hibiscus (USDA, 2009). 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. There is no pheromone or lure currently available for use in trapping O. hyalinipennis.

Surveys should be focused where the greatest risk for establishment occurs. According to Holtz (2006), O. hyalinipennis could potentially complete four to seven 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 have areas that are at low to moderate risk for O. hyalinipennis establishment based on climate and host availability. 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). This pest was recently found in 2010 in Monroe County, Florida (FDACS, 2010) and also in Puerto Rico and the U.S. Virgin Islands (USDA-APHIS, 2010). The pest was only ever detected at two locations in the lower Keys and has been identified nowhere else throughout Monroe County, Florida in spite of repeated surveys.

Survey site selection: Surveys should be conducted in high risk areas. “In Florida, this may include cultivated or wild cotton fields in southern Florida closer to the Caribbean islands where it is currently known to be established. Areas with regular traffic from countries with known infestations that may carry hitchhiker bugs should also be targeted for regular surveys” (USDA-APHIS, 2010).

Time of year to survey: Surveys should be carried out when the host plants are in seed. Surveyors for cotton should examine crops when host plants have newly matured bolls and dry seeds (Derksen et al., 2009). For early detection surveys, surveying during the quiescent period of the host is not recommended. This is due to the cryptic nature of O. hyalinipennis (USDA-APHIS, 2010).

A sampling protocol has been developed for surveying for this pest in cotton. There is also another set of survey procedures developed by APHIS that is found in the New Pest Response Guidelines for O. hyalinipennis (USDA-APHIS, 2010). This manual includes information on detection surveys as well as delimiting and monitoring surveys for if the pest is found in the United States.

145 Oxycarenus hyalinipennis Secondary Pest of Grape 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. Sweep netting is not recommended unless the pest has a high likelihood of being found (USDA-APHIS, 2010). 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).

Trees that adults can commonly be found on include Ficus, Acacia, and some Eucalyptus. Adults prefer rougher bark to smoother bark. Adult colonies can be found from near ground level to 6 to 7 m (19.7 to 23.0 ft.) (Kirkpatrick, 1923).

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.

Ultraviolet lights: “UV-light traps are not recommended for surveying for the cotton seed bug except in cases where there is a need to confirm eradication or enhance detection of a known population. UV-light traps are not pest specific, and consequently are cumbersome and time-consuming for sampling and identification purposes. In addition, it is unclear whether or not UV-light traps would be an effective monitoring tool for the cotton seed bug. Kirkpatrick (1923) demonstrated positive phototropism in laboratory experiments; however, when Kirkpatrick placed light traps at night in the direct path that the cotton seed bug was known to use between a tree and nearby field where they were coming from, no individuals were captured. It was concluded that the cotton seed bug did not migrate at night, and was not attracted to light at night” (USDA-APHIS, 2010). “Conversely, Nakache and Klein (1992) noted that the cotton seed bug was strongly attracted to light at night in Israel. Additional research regarding the efficacy of UV-light traps is needed” (USDA-APHIS, 2010).

Key Diagnostics/Identification CAPS-Approved Method*: Morphological examination of adults is needed to confirm identification. A field screening aid is available for O. hyalinipennis on the CAPS website at http://caps.ceris.purdue.edu/webfm_send/529. Final identification should be confirmed by dissecting and examining adult male internal structures (Brambila, 2010).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

146 Oxycarenus hyalinipennis Secondary Pest of Grape Arthropods Cottonseed bug True bug

Literature-Based Methods: The key diagnostic involves morphological examination of adults.

A technical description of O. hyalinipennis can be found in Samy (1969). The egg and nymph stage are described in Sweet (2000).

Easily Confused Pests A similar oxycarenid, Metopoplax ditomoides, is exotic to the United States but currently found in Oregon (Lattin and Wetherill, 2002; USDA-APHIS, 2010). The anterior part of the head of M. ditomoides is more rounded versus acute (USDA-APHIS, 2010).

References Avidov, Z., and Harpaz, I. 1969. Plant Pests of Israel. Jerusalem, Israel University Press.

Derksen, A., Griffiths, K., and Smith, T.R. 2009. 2009 Florida CAPS Oxycarenus Survey Report. Florida Cooperative Agricultural Pest Survey Program Report No. 2009-05-OH-01.

FDACS (Florida Department of Agriculture and Consumer Services). 2010. New cotton pest identified in Florida Keys. Florida Department of Agriculture and Consumer Services Press Release, April 23, 2010.

Holtz, T. 2006. Qualitative analysis of potential consequences associated with the introduction of the cottonseed bug (Oxycarenus hyalinipennis) into the United States. USDA-APHIS.

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) (Heteroptera: Lygaeidae). Journal of Chemical Ecology 10: 641-649.

Lattin, J.D. and Wetherill, K. 2002. Metopoplax ditomoides (Costa), a species a Oxycarenidae new to North America (Lygaeoidea: Hemiptera: Heteroptera). Pan-Pacific Entomologist 78:63–65.

Nakache, Y. and Klein, M. 1992. The cotton seed bug, Oxycarenus [sic] hyalinipennis, attacked various crops in Israel in 1991. Hassadeh 72: 773-775.

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: Oxycarenidae) 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.

147 Oxycarenus hyalinipennis Secondary Pest of Grape Arthropods Cottonseed bug True bug

Sweet, M.H. 2000. Seed and Chinch Bugs (Lygaeoidea). In C. W. Schaefer and A. R. Panizzi (eds.). Heteroptera of Economic Importance. CRC Press, Boca Raton. pp143–264.

USDA. 1983. Pest not known to occur in the United States or of limited distribution, No. 38: Cottonseed bug.

USDA. 2009. Significant Pest Bulletin Cottonseed Bug. United States Department of Agriculture, Animal and Plant Health Inspection Service. 1 p.

USDA–APHIS. 2010. Cotton seed bug New Pest Response Guidelines, Oxycarenus hyalinipennis. USDA–APHIS–PPQ–Emergency and Domestic Programs–Emergency Management, Riverdale, Maryland. http://www.aphis.usda.gov/import_export/plants/manuals/emergency/downloads/nprg- cotton_seed_bug.pdf

148 Planococcus lilacinus Secondary Pest of Grape Arthropods Oriental mealybug Mealybug

Planococcus lilacinus

Scientific Name Planococcus lilacinus Cockerell

Synonyms: Dactylopius coffeae, D. crotonis, Planococcus crotonis, P. deceptor, P. tayabanus, Pseudococcus coffeae, P. crotonis, P. deceptor, P. lilacinus, P. tayabanus, Tylococcus mauritiensis

Common Name(s) Oriental mealybug, cacao mealybug, coffee mealybug

Type of Pest: Mealybug

Taxonomic Position Class: Insecta, Order: Homoptera, Family: Pseudococcidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2006 through 2009; Additional pest of concern.

Pest Description Descriptions of the egg, nymph, male pupa, and adult male are lacing except for a brief description of the life history by van der Goot (1917).

Adult females: In the field, the adult females of P. lilacinus may be easily distinguished from P. citri by the much more globose shape of P. lilacinus (Fig. 1, Fig. 2). Beneath the pink to purple wax coating, the body is yellowish. The mid-dorsal line is fairly wide but indistinct. Illustrations of external features are available in Le Pelley (1968).

The mounted female is broadly oval to Figure. 1. P. lilacinus (2.7 mm) on rotund, length 1.2 to 3.1 mm (0.047 to 0.12 cocoa pod. Photo courtesy of in.), width 0.7 to 3.0 mm (0.03 to 0.018 in.). CABI, 2007. Margin of body with a complete series of 18 pairs of cerarii, usually all with stout conical setae. Legs stout: hind trochanter + femur 210 to 315 µm long, hind tibia + tarsus 210 to 275 µm long, ratios of lengths of hind tibia + tarsus to hind trochanter + femur 0.77 to 0.97; translucent pores present on hind

149 Planococcus lilacinus Secondary Pest of Grape Arthropods Oriental mealybug Mealybug

coxae and tibiae. The inner edges of ostioles are strongly sclerotized. Circulus large and quadrate, width 105 to 200 µm. Cisanal setae noticeably longer than anal ring setae. Anal lobe cerarii each situated on a moderately sized, well-sclerotized area (Cox, 1989).

Venter: Multilocular disc pores occurring on median area only, present around vulva as single or double rows across posterior borders of median areas of abdominal segments IV to VII and usually in a single row across anterior edge of segment VII (although the latter is sometimes reduced to a few pores). A few pores are sometimes present on anterior edges of median areas Figure 2. Adult female P. lilacinus. (left); of abdominal segments V and VI. winged male (right). From Kantheti, 1994; original photograph by Dr Dorsum: Multilocular disc pores and tubular K. S. Jayarama). Each unit in the scale ducts absent. Setae very long, stout, and represents 0±5 mm. flagellate (a character which distinguishes P. lilacinus from many other Planococcus species). Length of longest setae on abdominal segments VI or VII is 50 to 140 µm.

Symptoms/Signs There is very little information on the symptoms of attack by P. lilacinus in the available literature, although it has been said to cause severe damage or is listed as being a serious or the main pest of coffee, tamarind, custard apples, coconuts, cocoa, and citrus (CABI, 2007).

Symptoms on coconuts and cocoa are described as button nut shedding, drying up of inflorescence, and the death of tips of branches. Dense colonies form conspicuous patches on fruits; copious honeydew excretion may result in sooty mold development near colonies and the attraction of attendant ants. Fruits have been reported to have an abnormal shape and drop Figure 3. P. lilacinus on prematurely. mango fruit. Photo courtesy of Mango Information Network. Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods:

150 Planococcus lilacinus Secondary Pest of Grape Arthropods Oriental mealybug Mealybug

Visual Survey: In the field, P. lilacinus may be detected by thoroughly inspecting its normal habitats such as fruits, growing plant tips, shoots, and roots (CABI, 2007). On plants such as pomegranates, Annona spp., and Citrus spp., infestations on fruits, which tend to be heavy, can easily be detected (Fig. 3). Quarantine procedures should include thorough inspection of fruits, plant parts, and seedlings of suspect host plants using a hand lens. Males were said to pupate on the underside of leaves and to be scarce (van der Goot, 1917).

On cocoa, P. lilacinus is attended by several ant species, including Dolichoderus bituberculatus (commonly referred to as the black ant) in Java and Oecophylla longinoda, Technomyrmex detorquens and Odontomachus haematodus in Sri Lanka. In the Philippines, P. lilacinus is attended by Anoplolepis longipes [Anoplolepis gracilipes], an ant which on cocoa in Java tends to displace D. bituberculatus.

Surveys should be focused where the greatest risk for establishment occurs. A recent host analysis by USDA-APHIS-PPQ-CPHST indicates that most states in the United States are at low risk for P. liliacinus establishment based on host availability. Areas Alabama, California, Florida, and Georgia have the highest risk for establishment of P. liliacinus.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Reliable identification requires detailed study of slide- mounted adult females. P. lilacinus has often been mistaken in the field for other Planococcus spp. on cocoa and coffee, such as P. citri and P. kenyae, because of its yellowish body color beneath the wax and the presence of a median dorsal stripe extending from the first thoracic to the mid-abdominal region. It can, however, be distinguished by its more globose body shape and by having a much wider, but indistinct, median dorsal stripe.

The slide-mounted female is also quite distinct by the combination of stout legs, long dorsal setae and reduced number of multilocular disc pores, which occur mainly in small numbers in the posterior abdominal segments. A Lucid tool on mealybugs has been recently developed (see http://www.sel.barc.usda.gov/ScaleKeys/ScaleInsectsHome/ScaleInsectsMealybugs.ht ml).

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

Cox, J.M. 1989. The mealybug genus Planococcus (Homoptera: Pseudococcidae). Bulletin of the British Museum (Natural History), Entomology, 58(1):1-78.

Le Pelley, R.H. 1968. Pests of coffee. London and Harlow, UK: Longmans, Green and Co Ltd.

151 Planococcus lilacinus Secondary Pest of Grape Arthropods Oriental mealybug Mealybug van der Goot, P. 1917. De zwarte cacao-mier (Dolichoderus bituberculatus, Mayr.) en haar Beteekenis voor de cacaocultuur op Java, Meded. Proefst. Midden-Java, Salatiga, 25:142pp.

152 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

Thaumatotibia leucotreta

Scientific Name Thaumatotibia leucotreta Meyrick

Synonyms: Cryptophlebia leucotreta

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

Type of Pest: Moth

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

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

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 Figure 1. Larvae of T. leucotreta. recorded as feeding on over 50 different Photo courtesy of T. Grove and W. plant species. The generalist feeding Styn. www.bugwood.org. 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).

153 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

Eggs: Eggs are flat, oval (0.77 mm (0.03 in.) long by 0.60 mm (0.02 in.) wide) shaped discs with a granulated surface. The eggs are white to cream colored when initially laid, then changing to reddish color before the black head capsule of the larvae becomes visible under the chorion prior to eclosion (Daiber, 1979a).

Larvae: First instar (neonate) larvae approximately 1 to 1.2 mm (0.04 to 0.05 in.) 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 (0.47 to 0.71 in.) long, with a brown head capsule and first thoracic segment (Fig. 1). The last abdominal segment bears an anal comb with two to seven 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, Figure 2. Adult false codling moth. Photo respectively (Daiber, 1979b). courtesy of CABI, 2007.

Pupae: Prepupa and pupa form inside a lightly woven silk and soil cocoon formed by the fifth instar larvae on ground. Length is 8 to 10 mm (0.31 to 0.39 in.) and sexual determination through morphological differences on pupal case is possible (Daiber, 1979c)

Adult: Adult body length 6 to 8 mm (0.24 to 0.31 in.), wingspan of female and male moth is 15 to 20 mm (0.59 to 0.79 in.) and 15 to 18 mm (0.59 to 0.71 in.), respectively. Body brown, thorax with 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 sport 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).

Symptoms/Signs In general, the habit of internal feeding by FCM larvae (Fig. 3A) 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.

154 A

Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

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 A found around a fresh penetration hole or a mass of frass may be found around older penetration holes (Fig. 3B). Sometimes, however, frass is not visible. The area around the penetration hole can become sunken and brown as damaged tissue decays (Fig. 3C) (Johnson, date unknown).

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 B vulnerable to fungal organisms and scavengers. There is sometimes a scar visible on infested fruit (Stibick, 2006).

Corn: Larvae damage corn by entering the ear from the husk through the silk channel (Stibick, 2006).

Cotton: Larval penetration of cotton bolls facilitates entry of other microorganisms that can rot and destroy the boll. The cultivars Edranol, C 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 (0.55 to 0.75 in.) 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). Figure 3. A) FCM larva inside Avocado: Moths lay eggs superficially on the fruit grape fruit. B) A larval penetration of avocado. Larvae hatch and hole near the stalk with frass develop, and can enter through the skin. Larvae exuding from it as the larva feeds are unable to develop in avocado fruit. However, inside the fruit. C) Fruit damage their entrance creates lesions that lessen the around the penetration hole in a marketability of fruit. Lesions develop into a raised grape where a FCM larva entered crater on the fruit surface, with an inconspicuous the fruit. Photos courtesy of hole in the center where the larva has entered. Shelley Johnson, University of Granular excreta can also be seen (Stibick, 2006). Stellenbosch, South Africa.

155 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

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 frass. 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 (USDA, 1984; Stibick, 2006).

Survey CAPS-Approved Method*: The CAPS-approved method is a trap and lure combination. The approved trap is a wing trap.

Either of the following Trap Product Names in the IPHIS Survey Supply Ordering System may be used for this target: 1) Wing Trap, Paper 2) Wing Trap, Plastic

The Lure Product Name is “Thaumatotibia leucotreta Lure.” The lure is effective for 56 days (8 weeks).

Trap Spacing: When trapping for more than one species of moth, separate traps for different moth species by at least 20 meters (65 feet).

Method Note: The wing trap and the diamond trap are both effective traps for Thaumatotibia leucotreta. In order to standardize data and trap procurement, it is preferable that states use the wing trap. However, if states find reason to use the diamond trap, it is acceptable for negative data reporting. Diamond traps will not be available through the IPHIS Survey Supply Ordering System.

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

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Surveys should be focused in areas of high risk. A recent risk analysis by USDA- APHIS-PPQ-CPHST shows that portions of Alabama, Arkansas, California, Florida, Georgia, Louisiana, Mississippi, North Carolina, South Carolina, and Texas are at the greatest risk from T. leucotreta. Establishment of T. leucotreta is precluded in the northern United Stated based on climate and host range.

156 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

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 one trap per four hectares and traps should be no closer than 150 to 200 m (492 to 656 ft.) 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 eight 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 layed singly, which makes them 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 (0.039 in.) exit

157 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

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 (600 ft., 182.9 m) 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.

For early detection surveys in grape, vineyards 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 a frequency of one trap per four hectares and traps should be no closer than 150 to 200 m (492 to 656 ft.) to each other (Johnson, date unknown). Traps should be inspected weekly. Grapes 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.

Key Diagnostics/Identification CAPS-Approved Method*: Morphological. Confirmation of T. leucotreta is by morphological identification of adult specimens.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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.

158 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

See Padil website for additional FCM images, including diagnostic characters (http://www.padil.gov.au/pests-and-diseases/Pest/Main/136292).

A new identification tool, Tort AI – Tortricids of Agricultural Importance, is available at http://idtools.org/id/leps/tortai/ from CPHST’s Identification Technology Program. This tool contains larval and adult keys, fact sheets, an image gallery, molecular search capacity, and more. Thaumatotibia leucotreta is included in this tool.

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), however, 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 all other 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 (Bradly et al., 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: Olethreutinae. Vol. 2. London, England: British Museum (Natural History)

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

Couilloud, R. 1988. Cryptophlebia (Argyoploce) leucotreta (Meyrick). Lepidoptera, Tortricidae, Olethreutinae. Coton et Fibres Tropicales 43: 319-351.

Daiber, C. 1978. A survey of male flight of the false codling moth, Cryptophlebia 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.

Daiber, C.C. 1979b. A study of the biology of the false codling moth [Cryptophlebia leucotreta (Meyr.)]: the larvae. Phytophylactica 11, 141-144.

159 Thaumatotibia leucotretra Secondary Pest of Grape Arthropod False codling moth Moth

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. 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.

Gunn, D. 1921.The false-codling moth (Argyroploce leucotreta Meyr.). Science Bulletin No. 21. Pretoria, South Africa: The Government Printing and Stationary office.

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.

Möhr, J.D. 1973. Light trap studies with the false codling moth. Citrus and Sub-tropical Fruit Journal: 20- 22.

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.

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.

USDA. 1984. Pests not known to occur in the United States or of limited distribution, No. 48: False codling moth. USDA- APHIS-PPQ.

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.

160 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Pests of Grape Requested by the CAPS Community

Homalodisca vitripennis

Scientific Name Homalodisca vitripennis

Synonyms Homalodisca coagulata, Homalodisca triquetra, Phera vitripennis, Phera coagulata, Tettigonia coagulata

Common Names Glassy-winged Sharpshooter

Type of Pest Leafhopper

Taxonomic Position Class: Insecta, Order: Hemiptera, Family: Cicadellidae

Reason for inclusion in manual Request of the CAPS Community, PPQ Program Pest

Pest Description Eggs: Females lay their “sausage-shaped” eggs in masses of about 10 to 15 in the epidermis of the lower surface of young, fully developed leaves. The glassy-winged sharpshooter (GWSS) eggs measure approximately 2.5 mm (0.098 in. ) long and 0.53 mm (0.021 in.) wide (Boyd and Hoddle, 2007). When it is first laid, the egg mass appears as a greenish blister on the leaf (Fig. 1). The female covers the inserted egg mass with a secretion that resembles white chalk and is more visible than the leaf blister. Shortly after the eggs hatch, the leaf tissue begins to turn brown. The dead leaf tissue remains as a permanent brown scar (Fig. 2). Figure 1. Freshly laid “blister-like” eggs of H. coagulata. Photo courtesy of The University of Nymphs: Nymphs are the California.

161 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

immature insects that will later on become adults. Nymphs (Fig. 2) look similar to the adults except they are smaller, wingless, uniform greenish gray in color, and have prominent red bulging eyes. Nymphs range in size from 0.07 inches (2 mm) to nearly ½ inch (13 mm) long. The nymphs are much more active than the adults and are more likely to jump away when approached.

Adults: Adults (Fig. 3) are about 1.5 to 2.0 cm (0.59 in. and 0.79 in.) in length and are relatively large for the sharpshooter family. They are generally dark brown to black when viewed from the top or side. The abdomen is whitish or yellow. The head is brown to black and covered with numerous white to yellowish spots (Nielson, 1968). The wings of the sharpshooter are translucent brown with red veins. The structure of H. coagulata female genetalia is described in detail by Hummel et al. (2006).

Biology and Ecology Sanderson (1905) observed that H. coagulata has two to three generations per year in Texas, and the adults hibernated in ‘rubbish’ on the ground near food plants. Turner and Pollard (1959) using yellow sticky traps to study H. coagulata in Georgia, found that the insect had two full generations per year, followed by a partial third generation. Bivoltine patters of H. coagulata occur in Florida (Alderz and Hopkins, 1979) and southern California (Blua et al., 1999).

Adults are present and must feed throughout the year. In California, egg-laying activities are either Figure 2. Older eggs absent or reduced to very low levels during the darken, hatch, and winter months of December, January, and February. appear as a brown scar During this same period, the number of (top) and GWSS nymphs overwintering adults also decreases. Mating occurs (bottom). Photos courtesy in the spring and summer. Egg-laying resumes in of Ken Peek, Alameda late February and continues through May. The first County Dept. of generation completes development from late May to Agriculture and the late August. Adults from this generation lay egg University of California masses from mid-June through late September, which give rise to overwintering adults (CDFA, 2006).

162 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Symptoms/Signs Even though the GWSS is large enough to be seen with the naked eye, it is very A

inconspicuous in nature. The brown coloration of the insect blends very well with the color of the twigs where it is usually found, and it hides by moving to the other side of the twig or branch when it detects movement or is otherwise approached or disturbed.

GWSS eggs are laid together on the underside of leaves, usually in groups of 10 to 12. The egg masses appear as B small, greenish blisters. The eggs are covered with a white material scraped from deposits on the female forewings. This white powder is termed brochosomes, consisting of intricately structured hydrophobic particles. These blisters are easier to observe after the eggs hatch, when they appear as tan to brown scars on the leaves.

When a large number of sharpshooters are in a tree they constantly draw fluids Figure 3. Adults of H. coagulata. The out of the branches and excrete them profile of the sharpshooter (A) and out the other end giving the appearance the characteristic red veins in the of “rain” coming off the tree. The excreta wings. (B). Photos courtesy of CDFA. also can cause a whitewashed appearance on leaves, fruit and even on the sidewalk under it. Leaves or fruit coated with a whitish, powdery material may indicate that there has been heavy GWSS feeding on that plant (Fig. 4). There are other ways to create this whitewashed appearance, mineral deposits from water is one common way. White spots on plants are not always a sign that sharpshooters are present, but if you see it, closer inspection may reveal the sharpshooter hiding on Figure 4. Whitewashed appearance to fruit the stems or branches of the due to a GWSS infestation. Photo courtesy plant. of The University of California.

163 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Because the sharpshooter can remove a large amount of water from the xylem tissue, during drought conditions plants may become weakened if the water is not replenished.

Pest Importance The GWSS is a polyphagous pest native to the southeastern region of the United States. It is usually not a serious pest in the area of its native distribution. It was first reported in California in 1989 and has since spread throughout southern California. GWSS feeds on the plant’s xylem fluid and can acquire and transmit the Xylella fastidiosa bacterium (Fournier et al., 2006). Strains of this xylem-limited bacterial pathogen are responsible for several plant diseases including Pierce’s disease, almond leaf scorch, oleander leaf scorch, citrus variegated chlorosis, phony peach disease, oak leaf scorch, and etc. The bacterium replicates in the mouthparts of the insect (Hill and Purcell, 1995), and the pathogen is lost during the molting process (Purcell and Finlay, 1979). However, if an insect acquires X. fasitidiosa as an adult, it can retain the pathogen for the remainder of its life (Severin, 1949). In microscopic studies of X. fastidiosa-infected glassy-winged sharpshooters, bacteria were observed in the cibarium and apodemal groove of the diaphragm (Brlansky et al., 1982, 1983). These data suggest a noncirculative mechanism of transmission (Purcell and Finlay, 1979).

The GWSS has an extremely broad host range of over 200 species (CDFA, 2001), and a single individual will feed on a variety of plant species during its lifetime. The high mobility (distances greater than 90 m, 295.88 ft.) of this insect (Blackmer et al., 2004), and its use of a large number of host plant species, provides this vector with ample exposure to multiple X. fastidiosa strains during its lifetime (Costa et al., 2006). Almeida et al (2005) showed that H. coagulata can acquire X. fastidiosa from and inoculate the bacterium into dormant grape plant. Transmission to dormant plants during the winter is a potential problem in vineyards adjacent to citrus groves or other habitats with large overwintering populations of H. coagulata.

In Texas, the single greatest threat to the production of susceptible grape cultivars is Pierce's Disease. The disease has caused millions of dollars in losses to the state's wine industry since 1990 and the problem has escalated in the past five years. Also within the last few years, the GWSS has established itself in southern California where it also poses a potentially serious threat to the entire wine and table grape industry in that region of the country. H. coagulata currently is considered to be the most significant insect pest threatening the California grape industry (Purcell and Saunders, 1999).

Known Hosts Over 200 host plants in 35 families are known to serve as hosts for H. coagulata (CDFA, 1991). Major agricultural, ornamental, and weed hosts are listed below. For a complete listing see: http://pi.cdfa.ca.gov/pqm/manual/htm/454.htm#a.

Hosts: Amaranthus spp. (pigweed), Acacia spp. (wattles), Ambrosia spp. (ragweed), Abelmoschus esculentus (okra), Canna spp. (canna), Capsicum spp. (chilli pepper), Cercis spp. (red bud), Chenopodium spp. (lambsquarter), Chamaedorea spp.(palms),

164 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Chrysanthemum spp. (chrysanthemum), Citrus spp. (citrus), Coleus spp. (coleus), Crassula spp. (crassula), Dianthus spp. (dianthus), Elaeagnus spp. (elaeagnus), Erigeron spp. (fleabane), Eucalyptus spp. (eucalyptus), Euonymus spp. (euonymus), Ficus spp. (fig), Fraxinus spp. (ash), Gladiolus spp. (gladiolus), Hedera spp. (ivy), Helianthus spp. (sunflower), Hibiscus spp. (hibiscus), Ilex spp. (holly), Ipomoea spp. (morning glory), Jasminum spp. (jasmine), Juniperus spp. (juniper), Lactuca spp. (lettuce), Lagerstroemia spp. (crape myrtle), Lantana spp. (shrub verbena), Laurus nobilis (sweet bay), Ligustrum spp. (privet), Liriope spp. (giant turf lily), Lonicera spp. (honeysuckle), Malus spp. (apple), Morus spp. (mulberry), Nephrolepis spp., (sword fern), Nerium spp. (oleander), Olea spp. (olive), Persea spp. (avocado), Philodendron spp. (philodendron), Phlox spp. (phlox), Phoenix spp. (date palm), Photinia spp. (photinia), Pinus spp. (pine), Pittosporum spp. (pittosporium), Prunus spp. (peach, apricot, cherry), Quercus spp. (oak), Rhus spp. (sumac), Rosa spp. (rose), Rubus spp. (blackberry), Salix spp. (willow), Schinus spp. (pepper tree), Solanum spp. (potato, eggplant), Solidago spp. (goldenrod), Sonchus spp. (sonchus), Strelitzia spp. (bird of paradise), Thuja spp. (arborvitae), Tradescantia spp. (spiderwort), Ulmus spp. (elm), Viburnum spp. (viburnum), Vinca spp. (periwinkle), Vitis vinifera (grape), Washingtonia spp. (Washington palm), Wisteria spp. (wisteria), Yucca spp. (yucca), Xylosma spp. (xylosma), and Zea spp. (corn).

Known Vectors (or associated insects) The GWSS is a known vector of several strains of Xylella fastidiosa (see pest importance section),

Known Distribution Turner and Pollard (1959) describe the range of H. coagulata as a strictly southern species native to the United States, abundant from the latitude of Augusta, Georgia to Leesburg, Florida, having a western boundary of Val Verde and Edwards counties in Texas. Current distribution data shows that the sharpshooter is present to some extent in northern Mexico and south Florida as well. The described native range includes Alabama, Arizona, Arkansas, Florida, Georgia, Louisiana, Oklahoma, Mississippi, Missouri, North Carolina, South Carolina, and Texas (NAPPO, 2005; Overall et al., 2010).

Sorensen and Gill (1996) noted that the range of H. coagulata has extended to include several counties in southern California, most likely having been introduced to the state through the nursery industry. The infested counties in California include: Los Angeles, Orange, Riverside, San Bernadino, San Diego, and Ventura Counties. Limited infestations of GWSS also occur in areas of Fresno, Kern, Imperial, Sacramento, Santa Barbara, Santa Clara, Solano, and Tulare Counties (CDFA, 2006). In May 2004, the GWSS was identified from Hawaii (Hawaii Department of Agriculture, 2004).

North America: Mexico and the United States. Oceania: Cook Islands, Easter Island, French Polynesia (Tahiti), (Logarzo et al., 2006; CABI, 2007; IPPC, 2007).

165 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Potential Distribution within the United States GWSS is abundant in the southeastern United States and infestations occur in California and Hawaii.

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

Literature-Based Methods: Surveys in California are currently being conducted using a combination of sticky traps and visual surveys (CDFA, 2006).

Note: Sticky cards will not detect sharpshooters in the juvenile or egg stages. Thus, utilizing both sticky traps and visual survey is the best way to find new infestations.

Trapping: Yellow sticky traps are commonly used for surveillance and detection of adults in orchards, although H. coagulata only shows slight, if any, preference for yellow compared to other colors (CABI, 2007). Panels measuring a minimum of 5” x 9” are the trap of choice for GWSS. The flight temperature threshold for GWSS is approximately 18 °C (65 °F). Trapping will not be effective during periods when temperatures are lower than this threshold. Trapping season begins no earlier than March 1 and ends October 31, depending on local conditions. Trap densities from 1 to 5 traps per sq. mile are used for areas with 25 to 501 residences per square mile (CDFA, 2006). Rural areas with less than 25 homes per square mile should not be trapped. The number of traps should be doubled in the area within ¼ mile of high-risk nurseries (those which receive plant material from GWSS-infested areas). Irrigated areas with a diversity of plants which include multiple preferred hosts should be selected whenever possible. Traps should be placed near lights when possible. Preferred hosts should always be selected for trap deployment. Good hosts include the following:

• Spring: Crape myrtle, citrus, privet, photinia, grape, mulberry, xylosma, oleander, pittosporium, and euonymus.

• Summer: Crape myrtle, citrus, privet, photinia, grape, mulberry, xylosma, oleander, ornamental plum, pear, peppertree, red bud, pittosporium, and euonymus.

• Fall: Citrus, crap myrtle, oleander, olive, red bud, photinia, privet, and xylosma.

Traps should be placed primarily in the outer canopy of host trees (e.g., citrus) and highly visible. Traps should be serviced every two weeks and replaced/relocated every six weeks to another host at least 500 feet (152.4 m) away. For grape, traps should be placed from the edge to within 30 feet (9.1 m) of vineyards for early detection (University of California and USDA, date unknown). Traps may be placed on trellises and above the canopy or poles/stakes can be used to suspend the yellow panel traps just above the grape canopy. Deployment in perimeter rows or along heavily traveled routes should be avoided. The traps should be hung perpendicular to the vine so that

166 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

both sides of the trap are visible (Yolo County, 2004). Traps should be moved as vine growth and development occurs to prevent traps from being obscured by foliage. One sticky trap per 10 acres is recommended but placement in the area is dependent upon terrain and surrounding GWSS host plants (particular attention should be given to areas bordered by citrus). Additionally if a winery is present at the vineyard location, place traps at the edge adjacent to the winery or landscape plantings. For grapes, replacement of traps every two weeks is recommended.

Visual surveys: Visual surveys can be conducted by examining the terminal leaves, petioles, and stem in citrus and grapes. Surveys should begin no earlier than June 1 and end no later than October 31. GWSS infestations can also be determined by examining the underside of plant leaves or recently matured foliage (older foliage should be avoided) for the presence of adults, nymphs, nymphal cast skins, egg masses, and egg scars. Inspections for egg masses and nymphs are best restricted to known hosts. Old egg scars are the easiest to detect since egg deposition sites are visible on both leaf surfaces. Newly laid eggs, in contrast, are only visible on the underside of the leaves. Consequently, a representative sample of leaves should be turned over and examined with for egg masses. Backlighting against a sunny sky will also aid in finding egg masses (CDFA, 2006).

When searching for active life stages stages (nymphs and adults) search the plant stems. On trees, the GWSS usually selects shoots that are growing upward (vertically oriented as opposed to horizontal twigs. New flush growth and southern exposures are also preferred (CDFA, 2006). Adults are the easiest life stage to detect, because they are highly visible when flying around or between host plants. Flight activity is most pronounced during the late morning and afternoon hours.

Insect nets: Visual searches of the host plants can be enhanced by using insect nets (aerial and sweep and beat sheets). The effectiveness of these devices is largely dependent on the type and density of GWSS life stages present. At low densities and during cooler times, nets and beat sheets may be used to agitate foliage causing adults to take flight.

On warm nights, H. coagulata is attracted to black and incandescent lights (CABI, 2007).

Note: Detailed survey methods for GWSS are given at www.cdfa.ca.gov/.../2012_Survey_and_Delimitation_Protocols.pdf. Refer to this document for additional clarification and detail.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: The morphology of the adult has been used primarily to differentiate species of Homalodisca and other sharpshooters. If an egg mass is

167 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

discovered, considerable time and cost were required to rear the insects to the adult stage for morphological identification.

Predator gut analysis studies are often conducted to assess the efficacy of a potential biological control agent. Fournier et al. (2006) developed an ELISA technique to detect the egg stages of H. coagulata and H. liturata (the smoke tree sharpshooter) and to a lesser extent the adult stage of gravid females in the gut of predators. De Leon et al. (2006) used sequence characterized amplified region (SCAR) makers to develop markers that are H. coagulata and H. coagulata/H. liturata specific for use in predator gut content examinations. The authors stated, however, that these markers would be useful in identifying any life stage of H. coagulata and/or H. liturata.

Smith (2005) showed that the amino acid sequences of H. liturata and H. coagulata had only a single substitution that was unique (75th amino acid of alignment = serine in H. liturata and asparagine in H. coagulata). Though the majority of the genetic variation between the two species was neutral, the author felt that it could still be useful to develop a molecular diagnostic tool to discriminate between nymphs of H. liturata and H. coagulata.

Additionally, Costa et al. (2006) describe a PCR-based method to detect and differentiate strains (Pierce’s disease and oleander leaf scorch) of X. fastidiosa present in individual GWSS adults.

Easily Confused Pests Adults of H. coagulata appear dark brown to black in their overall appearance. The abdomen ranges in color from white to yellow. The head is brown to black and covered with numerous ivory to yellowish spots. These spots help distinguish glassy-winged sharpshooter from a close relative, the smoke tree sharpshooter (H. liturata, also listed as H. lacerata), which is native to the desert region of southern California and has pale, wavy lines instead of spots (Gill, 1995; CABI, 2007). The smoke tree sharpshooter is also slightly smaller (Gill, 1995). Immature stages are very difficult to distinguish (Smith, 2005).

References Alderz, W.D., and Hopkins, D.L. 1970. Natural infectivity of two sharpshooter vectors of Pierce’s disease of grapes in Florida. Journal of Economic Entomology 72: 916-919.

Almeida, R., Wistrom, P.P., Hill, B.L., Hashim, J., and Purcell, A.H. 2005. Vector transmission of Xylella fastidiosa to dormant grape. Plant Disease 89: 419-424.

Blackmer, J.L. Hagler, J.R., Simmons, G.S., and Canas, L.A. 2004. Comparative dispersal of Homalodisca coagulata and Homoalodisca liturata (Homoptera: Cicadellidae). Environmental Entomology 33: 88-99.

Blua, M.J., Phillips, P.A., and Redak, R.A. 1999. A new sharpshooter threatens both crops and ornamentals. California Agriculture 53(2) 22-25.

168 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Boyd, E.A., and Hoddle, M.S. 2007. Host specificity testing of Gonatocerus spp. egg-parasitoids used in a classical biological control program against Homalodisca vitripennis: A retrospective analysis for non- target impacts in southern California. Biological Control 43: 56-70.

Brlansky, R.H., Timmer, L.W., and Lee, R.F. 1982. Detection and transmission of a gram-negative, xylem-limited bacterium in sharpshooters from a citrus grove in Florida. Plant Disease 66: 590-592.

Brlansky, R.H., Timmer, L.W., French, W.J. and McCoy, R.E. 1983. Colonization of the sharpshooter vectors, Oncometopia nigricans and Homalodisca coagulata, by xylem-limited bacteria. Phytopathology 73: 530-535.

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

CDFA (California Department of Agriculture). 1991. CDFA official host list for glassy-winged sharpshooter. http://pi.cdfa.ca.gov/pqm/manual/htm/454.htm#a.

CDFA. 2006. Glassy-winged sharpshooter statewide survey and delimitation protocols as of 2002 (revised March 2006). http://www.cdfa.ca.gov/pdcp/Documents/Survey-Delimitation%20Protocols%2007- 08.pdf. Revised in 2012: www.cdfa.ca.gov/.../2012_Survey_and_Delimitation_Protocols.pdf.

Costa, H.S., Guzman, A., Hernandez-Martinez, R., Gispert, C., and Cooksey, D.A. 2006. Detection and differentiation of Xylella fastidiosa strains acquired and retained by glassy-winged sharpshooters (Hemiptera: Cicadellidae) using a mixture of strain-specific primer sets. Journal of Economic Entomology 99(4): 1058-164.

De Leon, J.H., Fournier, V., Hagler, J.R., and Daane, K.M. 2006. Development of molecular diagnostics markers for sharpshooters Homalodisca coagulata and Homalodisca liturata for use in predator gut content examinations. Entomolgia Experimentalis et Applicata 119: 109-119.

Fournier, V., Hagler, J.R., Daane, K.M., de Leon, J.H., Groves, R.L., Costa, H.S. and Henneberry, T.J. 2006. Development and application of a glassy-winged and smoke-tree sharpshooter egg-specific predator gut content ELISA. Biological control 37: 108-118.

Gill, R. J. 1995. New agricultural pest for Southern California, glassy-winged sharpshooter, Homalodisca coagulata. California Plant Pest and Disease Report 14: 67-68.

Hawaii Department of Agriculture. 2004. New Pest Advisory: Glassy-winged Sharpshooter. http://www.hawaii.gov/hdoa/pi/ppc/test-area/npa-1/NPA04-02-GWSS.pdf.

Hill, B.L., and Purcell, A.H. 1995. Acquisition and retention of Xylella fastidiosa by an efficient vector, Graphocephala atropunctata. Phytopathology 87: 1197-1200.

Hummel, N.A., Zalom, F.G., and Peng, C.Y.S. 2006. Structure of female genetalia of glassy-winged sharpshooter, Homalodisca coagulata (Say) (Hemiptera: Cicadellidae). Arthropod Structure and Development 35: 111-125.

IPPC. 2007. Notification of glassy-winged sharpshooter occurrence in the Cook Islands. IPPC Official Pest Report, No. CK-1/1. Rome, Italy: FAO. https://www.ippc.int/IPP/En/default

Logarzo, G.A., De Leon, J.H., Triapitsyn, S.V., Gonzalez, R.H., and Virla, E.G. 2006. First report of a proconiine sharpshooter, Anacuerna centrolinea (Hemiptera: Cicadellidae), in Chile, with notes on biology, host plants, and egg parasitoids. Annals of the Entomological Society of America 99(5): 879-883.

NAPPO. 2005. Glassy-winged sharpshooter detected in Arizona. North America Plant Protection Organization’s Phytosanitary Alert System.

169 Homalodisca vitripennis Requested by the CAPS Community Arthropod Glassy-winged sharpshooter Leafhopper

Nielson, M.W. 1968. The leafhopper vectors of phytopathogenic viruses (Homoptera: Cicadellidae) taxonomy, biology, and virus transmission. USDA Technical Bulletin 1382.

Overall L.M., Rebek, E.J., Wayadande, A.C. 2010. New distribution records of the glassy-winged sharpshooter, Homalodisca vitripennis (Germar), in Oklahoma (abstract). Southwestern Entomologist, 35(2):193-195.

Purcell, A.H., and Finlay, A.H. 1979. Evidence for noncirculative transmission of Pierce’s disease bacterium by sharpshooter leafhoppers. Phytopathology 69: 393-395.

Purcell, A.H., and Saunders, S.R. 1999. Glassy-winged sharpshooters expected to increase plant disease. California Agriculture 53: 26-27.

Sanderson, E.D. 1905. Miscellaneous cotton insects in Texas. United States Department of Agriculture, Farmer’s Bulletin No. 223, 18-21.

Severin, H.H.P. 1949. Transmission of the virus of Pierce’s disease of grapevines by leafhoppers. Hilgardia 19: 190-206.

Sorensen, J.T and Gill, R.J. 1996. A range extension of Homalodisca coagulata (Say) (Hemiptera: Clypeorrhyncha: Cicadellidae) to southern California. Pan-Pacific Entomologist 72: 160-161.

Turner, W.F. and Pollard, H.N. 1959. Life histories and behavior of five insect vectors of phony peach disease. USDA Technical Bulletin No.1188.

University of California and USDA. Date unknown. Managing the glassy-winged sharpshooter in vineyards. http://info.ucr.edu/gwss/GWSS_brochure.pdf.

Yolo County. 2004. Glassy-winged sharpshooter survey guidelines. http://www.yolocounty.org/org/AG/gwss.htm.

170 Cernuella virgata Secondary Pest of Grape Mollusk Maritime garden snail Mollusks

Primary Pests of Grape (Full Pest Datasheet)

None at this time

Secondary Pests of Grape (Truncated Pest Datasheet)

Cernuella virgata

Scientific name Cernuella virgata Da Costa

Synonyms: Cernuella virgatus, Cernuella variabilis, Cernuella virgata ssp. variegata, Helicella maritime, Helicella variabilis, Helicella virgata, Helix virgata

Common Name(s) Maritime garden snail, Mediterranean snail, Mediterranean white snail, striped snail, vineyard snail, white snail

Type of Pest Mollusk

Taxonomic Position Class: Gastropoda, Order: Stylommatophora (Eupulmonata), Family: Hygromiidae (Helicidae)

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2003 through 2011; Additional pest of concern (as Cernuella spp.)

Pest Description The shell of C. virgata is globose- depressed and white or yellowish- white in color with dark-brown bands or spots (Fig. 1, 2). Snail size is 6 to 19 mm (0.24 to 0.75 in.) high x 8 to 25 mm (0.31 to 0.98 in.) Figure 1. Banding of C. virgata. Photo courtesy wide. Shell size and banding of Tenby Museum patterns are reported to vary widely

171 Cernuella virgata Secondary Pest of Grape Mollusk Maritime garden snail

geographically throughout Southeastern Australia (Baker, 1988). Size has been demonstrated as inversely proportional to population density (Baker, 1988). C. virgata is considered polymorphic; banded and unbanded (more common) morphs have been found throughout Australia. Relative frequencies of each morph are likely correlated with site-specific factors such as predator pressure (Baker, 1988). The maritime gardensnail is relatively small and is characterized by prominent spiral banding on the shell (Fig. 1).

Symptoms/Signs C. virgata is found atop plants during summertime (Fig. 3) and may also be found feeding on new growth earlier in the season. These snails aestivate on plant heads and stalks, which contaminates crops and clogs machinery. Areas previously infested with Figure 2. C. virgata. Photo courtesy of L. Poggiani. snails can prevent the re- www.lavalledelmetauro.it. establishment of a site as pastureland as livestock often reject slime-contaminated hay and forage (Baker, 2002).

Survey CAPS-Approved Method*: Visual (Floyd, 2008).

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Visual survey: C. virgata is a conspicuous crop pest that hides during the day. Surveys are best carried out at night using a flashlight or in the morning or evening following a rain event. It is Figure 3. Multiple easily seen, and attacked plants exhibit extensive rasping and C. virgata on tree defoliation. Like other mollusks, it can also be detected by signs trunk. Photo of ribbon-like excrement and slime trails on plants and buildings. courtesy of L. Poggiani, A recent risk analysis by USDA-APHIS-PPQ-CPHST shows that www.lavalledelmet portions of North Dakota are at the greatest risk from this mollusk auro.it. based on host availability. Portions of Arkansas, California, Colorado, Delaware, Idaho, Illinois, Indiana, Iowa, Kansas, Kentucky, Louisiana, Maryland, Michigan, Minnesota, Missouri, Mississippi, Montana, Nebraska, New York, North Carolina, North Dakota, Ohio, Oklahoma, Oregon, Pennsylvania, South Dakota,

172 Cernuella virgata Secondary Pest of Grape Mollusk Maritime garden snail

Tennessee, Texas, Washington, and Wisconsin are also at moderate risk based on host availability.

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation requires a morphological identification. All specimens should be submitted to Patrick Marquez (Western Region) or John Slapcinsky (Eastern Region). Both Domestic Identifiers are able to identify (even immature specimens) to the species level for this genus.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: A morphological identification is required. C. virgata is a relatively small snail (up to 15 mm (0.59 in.) in diameter) characterized by prominent spiral banding on the shell.

C. virgata closely resembles the white Italian snail (Theba pisana) in appearance and pest status. C. virgata can be differentiated from T. pisana by more pronounced spiral banding. Also, the umbilicus (hole about which the shell spirals) appears as a circular hole rather than being partially obscured as in the white Italian snail (CABI, 2007).

References Baker, G.H. 1988. The life history, population dynamics and polymorphism of Cernuella virgata (Mollusca: Helicidae). Australian Journal of Zoology 36: 497-512.

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

Floyd, J. 2008. New Pest Response Guidelines: Temperate Terrestrial Gastropods. USDA-APHIS-PPQ- Emergency and Domestic Programs, Riverdale, Maryland.

173 Lissachatina fulica Secondary Pest of Grape Mollusk Giant African Snail

Lissachatina fulica

Scientific Name Lissachatina fulica Bowdich

Synonyms: Achatina fulica

Common Names Giant African snail, African giant snail, Kalutara snail

Type of Pest Mollusk

Taxonomic Position Figure 1. The large size of the giant African snail. Photo courtesy of USDA- Class: Gastropoda, Order: Pulmonata, APHIS. Family: Achatinidae

Reason for inclusion in manual CAPS Target: AHP Prioritized Pest List – 2003 through 2012

Pest Description Lissachatina 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 x 5 mm (0.16 x 0.20 in.) in diameter, usually pale yellow, laid in clutches of 100 to 400 (Fig. 3) (USDA, 1982).

Juveniles: Similar to adults, but have a thinner, more brittle, translucent shell. Upon emergence, the juvenile shell is approximately 4 mm (0.16 in.) long (Denmark and Poucher, 1969; USDA- APHIS, 2005). Growth increases at a rate of 10 mm (0.39 in.) 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 Figure 2. Adult giant African snail. Photo parietal callus are white or bluish-white courtesy of Matt Ciomperlik, USDA APHIS with no trace of pink (Bequaert, 1950). PPQ

174 Lissachatina fulica Secondary Pest of Grape Mollusk Giant African Snail

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 concave. Snails with a lesser concaved columella tend to be 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, Lissachatina 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 L. fulica feeding 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. L. fulica has been shown to be a health hazard by transmitting the rat lungworm, Angiostrongylus cantonensis, which causes eosinophicillic meningoencepahlitis in humans. L. 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).

L. 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

175 Lissachatina fulica Secondary Pest of Grape Mollusk Giant African Snail 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. In September 2011, L. fulica was again found in Miami-Dade County Florida. An eradication effort is ongoing for this newly discovered population.

Symptoms/Signs Information specific to grape is not available. L. 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. L. 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. 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 L. 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 L. fulica.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

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. L. 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.

Surveys should occur in areas that are at greatest risk for establishment of L. 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, Oregon, South Carolina, Texas, and Washington are at low to moderate risk from L. fulica. Risk of L. fulica establishment is either low or unlikely (precluded) 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.

176 Lissachatina fulica Secondary Pest of Grape Mollusk Giant African Snail

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 Lissachatina fulica remains active at a range of 9 to 29°C (48 to 84°F). Lissachatina 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 slugs; 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 slug 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 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 L. fulica is by morphological identification. Identification should be verified by a malacologist at National Identification Services.

177 Lissachatina fulica Secondary Pest of Grape Mollusk Giant African Snail

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: L. 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 (7.87 in. ) the shell is more commonly in the range of 5 to 10 cm (1.97 to 3.94 in.). 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., 1960. Some snails and slugs of quarantine significance to the United States. Plant Quarantine ARS 82-1. Agricultural Research Service, USDA, Washington, D.C.

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.

178 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode Nematodes

Primary Pests of Grape (Full Pest Datasheet)

Xiphinema italiae

Scientific Name Xiphinema italie Meyl.

Synonyms: Xiphinema arenarium, X. bulgariensis, and X. conurum.

Common Name(s) Dagger nematode

Type of Pest Nematode

Taxonomic Position Class: Adenophorea, Order: Dorylaimida, Family: Longidoridae

Reason for Inclusion in Manual National Threat

Pest Description Females of this dagger nematode are about 3 mm (0.12 in.) long. They have an odontostylet (Fig. 1) 87 to 112 µm long and an odontophore about 54 to 68 µm long. Their tail is elongate-conoid with slight dorsal and ventral constrictions towards the terminus. Males are rare.

From Martelli et al. Figure 1: A Xiphinema spp. using its stylet to penetrate a (1966) and Cohn (1977) host root. Photo courtesy of North Carolina State University. Measurements: Neotype female: L= 2.4 mm; a= 84; b= 6.4; c= 32; c’= 4; V= 46; spear= 89 µm; spear extension= 64 µm. For 5 topotype females: L= 2.48 (2.3-2.8) mm; a= 80 (75-84); b= 6.8 (6.4-7.3); c= 34 (30-40); c’= 3.8 (3.2-4.3); V= 46 (45-46);

179 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

spear= 91 (87-96) µm; spear extension= 59 (54-64) µm. For 4 males from Israel and France: L= 3.06 (2.4-3.35) mm; a= 86.5 (79-101); b= 7.9 (6.4-9.2); c= 48.8 (47-50); c’= 2.25 (2-2.5); spear= 101.3 (93-102) µm; spear extension= 61.8 (56-65) µm.

From Cohn (1977): Female: Body slender, about 3 mm (0.12 in.) long; almost straight in the first two-thirds of the body, then ventrally arcuate. Body cuticle smooth, 2 to 3 µm thick near middle, conspicuously thickened (5.0-6.5 µm) on dorsal side of tail; finely striated transversely, most conspicuously in caudal region. Lateral body pores in a single row, clearly discernible up to base of oesophagus, beyond which they are more difficult to see. Lip region gently rounded, 10 to 12 µm across, set off by a slight constriction. Labial papillae clearly visible. Amphids post-labial, stirrup-shaped, apertures slit-like, occupying between one-half and two-thirds of body diameter.

Spear (odontostyle): Straight, hollow, needle-like, about 95 µm long and furcated at base; spear extension (odontophore) about 60 µm long with prominent tripartite basal flanges, measuring 8 to 11 µm across. Basal ring of spear guiding sheath about 81 µm from anterior end. Nerve ring, 11 to 13 µm thick, located behind base of spear extension; hemizonid about 170 µm from anterior end and hemizonion, often obscure, 213 to 235 µm from anterior end. Anterior part of oesophagus slender, coiled, with a triangular mucro, 2 to 3 µm long in oesophageal wall about 20 µm behind base of spear extension and pointing toward the head. Posterior part of oesophagus cylindrical, strongly muscular, measuring about 100 x 18 µm; nucleus of dorsal oesophageal gland near the anterior extremity of the bulb. Oesophago-intestinal valve often indistinct, conical, about 8 x 5 µm.

Vulva: A transverse slit, slightly pre-median (V=43 to 49), leading to a vagina, which extends one-third into the body. Gonads paired, opposed, reflexed at oviduct. Z-organ absent. Uterus differentiated to form an elongated narrow section and a short dilated pouch, the two parts sometimes appear to be separated by a sphincter, which is not always distinct. The oviduct is separated from the uterine pouch by a prominent sphincter and connects with the largest oocyst of the ovary. Uterine eggs, measuring about 112 x 25 µm, have been observed. Rectum oblique, 1 to 1.5 anal body widths long.

Tail shape: Variable even within the same populations, most commonly elongate-conoid with slight dorsal and ventral constrictions towards the terminus; sometimes bluntly conoid, and even almost subdigitate. Tail length 2.8 to 4.3 times longer than anal body width (2.3 to 2.8 times in South African specimens), with 2 to 3 pairs of caudal papillae.

Males: Rare, not essential for reproduction and, apparently, not functional. Body shape similar to that of female, but tail somewhat shorter, more conoid, with a smaller c’ value, which ranges from 2.0 to 2.3 in the Mediterranean specimens and from 1.8 to 1.9 in the South African ones (Heyns, 1974). Spicules arcuate, measuring 44 to 48 µm along their axis; lateral guiding pieces of South African specimens 13 to 14 µm long with dorsal

180 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

projections. An adenal pair of supplements present and one or two additional vetromedian ones. Copulatory muscles strongly developed.

Juveniles: The four larval stages can be recognized by a progressive increase in length of body and of functional and replacement spears. The c’value also decreases with age due to the greater development of the caudal region in diameter than in length.

LARVAL N BODY FUNCTIONAL REPLACEMENT C’ STAGE LENGTH SPEAR (µM) SPEAR (µM) (MM)

L1 4 1.44 (1.35- 62.1 (60-64) 74.7 (74-76) 4.35 (4.3- 1.5) 4.4) L2 6 1.86 (1.75- 70.6 (69-72) 86.2 (84-89) 4.08 (3.9- 1.95) 4.3) L3 6 2.23 (2.2-2.3) 82.5 (82-84) 96.6 (92-99) 3.9 (3.9-4.0) L4 5 2.55 (2.49- 85.3 (82-87) 101.5 (99-104) 3.8 )3.7-3.9) 2.6)

Biology and Ecology Xiphinema spp. are large nematodes that live free in the soil. They have long stylets, which enable them to feed deep within roots. Many species concentrate on root tips where their feeding can stop extension of growth and induce formation of root tip galls, containing enlarged, sometimes multinucleate, cells. X. italiae feeds along young roots but not necessarily at the tips. Terminal root galls, though sometimes present, are not conspicuous in infected roots. Besides causing direct damage, these nematodes can transmit damaging viruses (Brown et al., 1993). Populations are highest in the top 30 cm (11.8 in.) of soil, with a threefold drop in the 30 to 60 cm (11.8 to 23.6 in.) layer and continuous decrease with increased depth (Cohn, 1969)

At 28°C (82°F), X. italiae requires six months to complete its life cycle (SON, n.d., Cohn and Mordechai, 1970). It failed to complete its lifecycle even after 12 months at constant temperatures of 24°C (75°F) or below. X. italiae is closely associated with sandy-loam or loamy-sand soils containing low percentages of fine particle fractions, with acid soils, and those with normal organic matter content (2.5 to 4.5%) (Navas and Arias, 1986; Tzortzakakis et al., 2006).

Symptoms/Signs It has ectoparasitic feeding habits and feeds along feeder roots causing root discoloration (darkening) and stunting. Plants also have fewer feeder roots. The cortex of infected roots disintegrates in several areas and, where, nematode infestation is heavy, entire roots are sometimes black and decayed (Cohn, 1970).

Pest Importance Because of their economic importance as plant pathogens and as vectors of viral disease of plants, Xiphinema spp. have been receiving increased attention by the

181 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

scientific community. Grapevine fan leaf virus (GFLV), which is potentially vectored by X. italiae, is responsible for a progressive degeneration of grapevines that occurs in most vineyards worldwide. GFLV causes serious economic losses by substantially reducing yield by up to 80% and affecting fruit quality. According to Wang et al. (2003), GFLV is the ‘most economically important nepovirus of grape world-wide’.

Known Hosts Definite proof of nematode multiplication has been obtained under controlled conditions only on grapevine and sour orange (Cohn and Mordechai, 1969). The nematode has been recovered in the rhizosphere of other plants, referred to here as associated hosts.

Major Hosts: Vitis spp. (grape)

Poor Host: Citrus aurantium (sour orange)

Associated Hosts: Prunus amygdalus (almond), Prunus armeniaca (apricot), Eucalyptus spp., Morus spp. (mulberry), Olea europaea (olive), Prunus persica (peach), Prunus domestica (plum), and conifers (SON, n.d.; Vovlas and Avgelis, 1988).

Known Vectors (or associated insects) Xiphinema italiae was shown by Cohn et al. (1970) to vector grape fanleaf virus (GFLV), a serious pathogen of grape, in Israel. In Israel, both larvae and adult females transmitted three different isolates of GFV within four to eight weeks after acquisition. It has not been shown to be a vector of GFLV in other countries, however, and its vector status needs to be confirmed (Catalano et al., 1992; Weiland Ardaiz and Perez, 1995).

Known Distribution Xiphinema italiae is a Mediterranean dagger nematode.

Africa: Algeria, Egypt, Libya, Nigeria, South Africa, and Tunisia. Asia: Israel and Turkey. Caribbean: Cuba. Europe: Bulgaria, Cyprus, France, Greece, Hungary, Italy, Portugal, Romania, Russia, Serbia, Slovakia, and Spain (SON, n.d.; Cohn, 1969; Romascu, 1971; Heyns, 1974; Lima, 1973; Chavez and Garaert, 1977; Polinovskij, 1979; Antoniou, 1981; Romanenko, 1985; Navas and Arias, 1986; Siddiqui et al., 1987; Vovlas and Avgelis, 1988; Peneva and Choleva, 1992; Khan et al., 1993; Liskova et al., 1993; Philis, 1993; Dias-Silveira and Herrera, 1995; Lamberti et al., 1996; Nagy, 1999; Knoetze et al., 2000; Barsi and Lamberti, 2003; Wang et al., 2003; Tzortzakakis et al., 2006; Teliz et al., 2007; Kumari and Liskova, 2009).

The record from Cameroon (Chavez and Geraert, 1977) is now thought to be X. savanicola (Luc and Aubert, 1985).

182 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

Potential Distribution within the United States This nematode has the potential to occur in all grape-growing regions of the United States.

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

Literature-Based Methods: Soil sampling: Philis (1993) sampled for Xiphinema index, but also found X. italiaea and X. pachtaicum using the following procedure in Cyprus: 1) samples were collected with a 10-cm (3.94-in.) diameter sampling tube (auger) to a depth of 20 to 25 cm (7.87 to 9.84 in.) in the root zone (all samples contained feeder roots). In each village area, six to seven vineyards were sampled at random with preference given to old vineyards. From each field, three to four separate samples, each of 600 to 800 g, were collected, put into polyethylene bags, and then into a cooler box with ice. Samples were stored in a refrigerator until required for processing, which always occurred within four to five days of collection. Note: A higher sampling rate would be required for early detection sampling (e.g., a protocol similar to that used in the national potato cyst nematode survey). Vovlas and Avgelis (1988) collected soil at a 20 to 40 cm (7.87 to 15.75 in.) in vineyards in Greece.

Extraction of Nematodes: Note: (From Robbins and Brown, 1991) “Longidorus, Paralongidorus, and larger Xiphinema species are much more common in North America than reported. Nematode extraction methods used in most North American laboratories are inappropriate for recovering very large and/or long nematodes such as Longidorus, Paralongidorus, and Xiphinema species. Frequently, these nematodes will be discarded with soil debris retained on the coarse screen during the extraction process. Also, these nematodes move sluggishly and only slowly migrate down through support meshes used with the Baermann funnel or mist extraction techniques. Furthermore, when sugar centrifugation extraction is used for recovering the larger longidorid nematodes, higher sugar density-gradients are required than routinely used in most North American laboratories for recovering small vermiform nematodes. Methods suitable for extracting virus vector nematodes, including the larger members of the Longidoridae, have been comprehensively described by Brown and Boag (1988)”.

Nematodes are extracted from soil by a modified sieving and decanting method (Brown and Boag, 1988). Protocols are given for Longidorids, to which X. italiae belongs. A slightly different version of the procedure is used for Trichodorids. Soil (200 g) is stirred thoroughly by hand to suspend all particles in 3 liters of tap water in a bucket. The resulting suspension is washed quickly after waiting 15 seconds into another bucket through a sieve of 2.0 mm (0.08 in.) aperture and the suspension is carefully stirred again. After 15 seconds of sedimentation, the suspension is slowly decanted through a 150 µm sieve. After 15 seconds of sedimentation, the suspension is slowly decanted through a 95 µm sieve. The residue on the sieve is carefully washed from the opposite

183 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

side of the sieve with a gentle jet of tap water over a smaller bucket and after 10 seconds the suspension is again gently washed through the same sieve. The process is repeated until the residue is without soil particles and the water at washing is clean. At the end of the last washing process, the residue is concentrated at one part of the sieve by bowing and the residue is rinsed from the sieve to a glass beaker. After sedimentation (24 hours), live individuals of X. italiae can be hand-picked directly from the suspension under a stereomicroscope. Individuals are generally preserved in 1 M NaCl and send to an approved diagnostic laboratory. Kumaria and Liskova (2009) used a slightly modified version of this protocol by starting with 500 g of soil and waiting 10 seconds after stirring. Tzortzakakis et al. (2006) used this same method.

Flegg (1967) used a modified version of Cobb’s decanting and sieving technique. Bulked soil samples were carefully mixed after being gently crumbled by hand to a maximum particle size of about 1 cm diameter. To 250 ml water in a calibrated beaker sufficient soil is added to raise the level to 450 ml providing a standard measure of 200 ml soil. The resulting slurry was poured into a 1 liter polypropylene beaker, which was then topped up with water. For most loam soils soaking for half an hour was sufficient, but heavy clays needed an hour and this time was adopted as the standard. During the soaking period, the slurry was stirred two or three times with a circular Perspex plate, of diameter just less than that of the beaker, bored with many conical holes and agitated rapidly up and down over the sediment. The resulting suspension was washed through a 2 mm (0.08 in.) aperture sieve into a 5 liter plastic bucket, and after topping up with water was swirled thoroughly by hand to suspend all particles. After a 25 second sedimentation time, the supernatant fluid was decanted through a bank of three 150 µm aperture sieves. The residue on the sieves was washed thoroughly with a gentle jet of water and collected in a beaker. The residue in the bucket was then topped up with water, again thoroughly swirled, and after 15 second settling, decanted through the same bank of sieves. After washing, the residue on the sieve was added to that collected previously. The sievings were then gently shaken to suspend all particles before pouring through a 90 µm aperture nylon screen supported by a polyethylene ring, which was immediately placed on a Baermann funnel containing enough water just to submerge the screen surface and debris. After 20 hours, about 25 ml of water containing the extracted nematodes can be run off from the funnel for examination under a stereoscope microscope at 25 or 50 X magnification.

Philis (1993) used a modification of the Flegg (1967) method for nematode extraction with final separation made through a 115 µm nylon sieve in 9-cm (3.54-in.) diameter petri dishes containing sufficient water to just cover the sieve surface. Samples were thoroughly mixed and an aliquot of 250 g was processed. The sieves were left for 24 hours and care was taken to ensure that the temperature during the extraction process did not exceed 23°C (73°F).

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

184 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

Literature-Based Methods: Morphological: Martelli et al. (1966) redescribe X. italiae and Cohn (1977) provides a morphological summary of several key taxonomic publications of X. italiae.

Identification of nematodes based on morphology and morphometrics is time- consuming and difficult due to the overlapping of many characteristics. Additionally Xiphinema species can be found together in the same field and often occur at low densities. If only a single or a few individuals are available for morphological analysis, identification can be quite difficult. The major characters for separating species of Xiphinema consist primarily of the following female features: 1. General morphology and body measurements (body length, ‘a’ ratio, stylet length, shape of lip region, etc.); 2. Reproductive system (position of vulva, single or double genital tract and state of regression of anterior gonad, presence or absence of Z organ); and 3. Tail structure (c ratio, tail length/anal body diameter (c’), tail shape) (Cohn and Sher, 1972). The structure of the female gonad is of the utmost importance in the taxonomy of the genus.

Loof and Luc (1990) provide a morphological key of 172 valid species of Xiphinema (excluding the X. americanum group), which includes X. italiae. Because of the large number of species, the key is split into eight groups on the basis of development of the anterior female genital branch, uterine differentiation, and tail shape. X. italiae belongs to group 7 where both female genital branches are equal, without uterine differentiation, and tail elongate to conical.

Molecular: Knoetz et al. (2000) used PCR-RFLP to distinguish between Xiphinema index, X. italiae, X. elongatum, X. americanum, and X. diffusum in South Africa.

Wang et al. (2003) developed a PCR-based method to distinguish Xiphinema spp. (X. index, X. diversicaudatum, X. vittenezi, and X. italiae) from grapevines in China. This method allows for the amplification of crushed nematodes without a separate DNA extraction step. It proved sensitive enough to accurately detect and characterize a single nematode at any developmental stage. This method can also be used in a multiplex test, which allows detection of single individuals belonging to several species mixed in the same sample.

Kumari and Liskova (2009) used the Wang et al. (2003) PCR method to validate their morphological determinations for X. italiae from Slovakia, and the authors also found that no PCR product was obtained in their water control or with DNA from X. brevicollum, X. dentatum, X. diversicaudatum, X. pachtaicum, X. simile, and X. vuittenezi. These authors also sequenced four independently evolving molecular makers (cox1, ITS2, D2/D3, and 18S) for X. italiae.

Hubschen et al. (2004) conducted a study to confirm the specificity, sensitivity, and reliability of the Wang (2003) primers for X. diversicaudatum, X. index, and a range of closely related longidorid species that are typically encountered in vineyard soil. A weak PCR product (3 of 8 replicate PCR reactions) was observed with one non-target organism, Longidorus elongates. The authors found the primer be sufficiently specific,

185 Xiphinema italiae Primary Pest of Grape Nematode Dagger Nematode

sensitive, and reliable to be used for phytosanitary/quarantine issues related to the viticulture industry in Europe.

Easily Confused Pests Other Xiphinema spp., especially X. index and X. mediterraneum, that occur on grape can be confused with X. italiae. In addition, other Xiphinema spp. that occur in group seven (Loof and Luc, 2000) can be morphology confused with X. italiae.

References Antoniou, M. 1981. A nematological survey of vineyards in Cyprus. Nematol. Medit. 9: 133-137.

Barsi, L., and Lamberti, F. 2003. Morphometrics of adults and juvenile stages of three longidorid nematodes (Nematoda:Dorylaimida) from Vojvodina Province, Northern Serbia. Nematol. Medit. 31: 65- 85.

Brown, D.J.F., and Boag, B. 1988. An examination of methods used to extract virus-vector nematodes (Nematoda: Longidoridae and Trichodoridae) from soil samples. Nematol. Medit. 16: 93-99.

Brown, D.J.F., Dalmasso, A., and Trudgill, D.L. 1993. Nematode pests of stone fruits and vines. In: Plant Parasitic Nematodes in Temperate Agriculture. K. Evans, D.L. Trudgill, and J.M. Webster (eds.). CAB International, Wallingford. Pp: 427-462.

Catalano, L., Savino, V., and Lamberti, F. 1992. Presence of grapevine fanleaf nepovirus in populations of Longidorid nematodes and their vectoring capacity. Nematol. Medit. 20: 67-70.

Chavez, E., and Geraert, E. 1977. Observations sur quelques Mononchides et Dorylaimides du Cameroun et du Zaire (Vermes-Nematoda). Revue Zool. Afrc. 3: 766-780.

Cohn, E. 1969. The occurrence and distribution of species of Xiphinema and Longidorus in Israel. Nematologica 15: 179-192.

Cohn, E. 1970. Observations on the feeding and symptomatology of Xiphinema and Longidorus on selected host roots. Journal of Nematology 2(2): 167-173.

Cohn, E. 1977. Xiphinema italiae. CIH Description of Plant Parasitic Nematodes. Commonwealth Institute Helminthology. St. Albans U.K. Set 7 no. 95.

Cohn, E., and Mordechai, M. 1969. Investigations of the life cycle and host preference of some species of Xiphinema and Longidorus under controlled conditions. Nematologica 15: 295-302.

Cohn, E., and Mordechai, M. 1970. The influence of some environmental and cultural conditions on rearing populations of Xiphinema and Longidorus. Nematologica 16: 85-93.

Cohn, E., and Sher, S.A. 1972. A contribution to the taxonomy of the genus Xiphinema Cobb, 1913. Journal of Nematology 4(1): 36-64.

Cohn, E., Tanne, E., and Nitzany, F.E. 1970. Xiphinema italiae, a new vector of grapevine fanleaf virus. Phytopathological Notes 60: 181-182.

Dias-Silveira, M.F., and Herrera, J.O. 1995. Principales problemas nematologicos en Cuba. In: Proceedings of Congresso International de Nematologia Tropical, Rio Quente, Brazil. Pp. 161-171.

Flegg, J.J.M. 1967. Extraction of Xiphinema and Longidorus species from soil by a modification of Cobb’s decanting and sieving technique. Ann. Appl. Biol. 60: 429-437.

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Heyns, J. 1974. The genus Xiphinema in South Africa II. X. elongatum-group (Nematoda: Dorylaimida). Phytophylactica 6: 249-260.

Hubschen, J., Kling, L., Ipach, U., Zinkernagel, V., Bosselut, N., Esmenjaud, D., Brown, D.J.F., and Neilson, R. 2004. Validation of the specificity and sensitivity of species-specific primers that provide a reliable molecular diagnostic for Xiphinema diversicaudatum, X. index, and X. vuittenezi. European Journal of Plant Pathology 110: 779-788.

Khan, F.A., Erinle, J.D., and Chindo, P.S. 1993. Survey of plant parasitic nematodes associated with grapevine in four northern states of Nigeria and observations on grapevine fanleaf virus. J. Afr. Zool. 107: 505-510.

Knoetze, R., Burger, J.T., and Meyer, A.J. 2000. Discrimination of some Xiphinema species from South Africa by rDNA-RFLP analysis. African Plant Protection 6(2): 25-30.

Kumari, S., and Liskova, M. 2009. Molecular confirmation of Xiphinema italiae Meyl, 1953 (Nemtatoda: Longidoridae) from the Slovak Republic. Helminthologia 46(2): 1310134.

Lamberti, F., Agostinelli, A., and Radicci, V. 1996. Longidorid nematodes from northern Egypt. Nematol. Medit. 24: 307-339.

Lima, M.B. 1974. Xiphinema italiae Meyl., 1953 (Nematoda: Dorylaimida) found around roots of grapevine in Portugal. Agronomia Lusitana 35(3): 273-273.

Liskova, M., Lamberti, F., Sabova, M., Valocka, B., and Agostinelli, A. 1993. First record of some species of Longidorid nematodes from Slovakia. Nematol. Medit. 21: 49-53.

Loof, P.A.A., and Luc, M. 1990. A revised polytomous key for the identification of species of the genus Xiphinema Cobb, 1913 (Nematoda: Longidoridae) with exclusion of the X. americanum-group. Systematic Parasitology 16: 35-66.

Luc, M, and Aubert, V. 1985. On the distribution of Xiphinema italiae Meyl, 1953 and X. savanicola Luc and Southey, 1980 (Nematoda: Longidoridae). Revue Nematol. 8(1): 85-86.

Martelli, B., Cohn, E., and Delmasso, A. 1966. A redescription of Xiphinema italiae Meyl., 1953 and its relationship to Xiphinema arenarium Luc et Dalmasso, 1963 and Xiphinema conurum Siddiqi, 1964. Nematologica 12: 183-194.

Nagy, P. 1999. New faunistic record for Longidorus attenuatus Hooper, 1961 and Xiphinema italiae Meyl. 1953 (Nematoda: Longidoridae) in Hungary. Novenyvedelem 35(1): 15.

Navas, A., and Arias, M. 1986. On the distribution and ecology of Xiphinema index and X. italiae in Spain. Nematol. Medit. 14: 207-215.

Peneva, V., and Choleva, B. 1992. Nematodes of the family Longidoridae from forest nurseries in Bulgaria. II. The genus Xiphinema Cobb, 1913. Helminthology 32(1): 46-58.

Philis, J. 1993. Distribution and ecology of Xiphinema index in Cyprus. Nematol. Medit. 21: 139-142.

Polinovskij, A.I. 1979. Species composition and geographic distribution of nematodes of the family Longidoridae (Nematoda: Dorylaimida) on grapevines of the Moldavian SSR, USSS. Izv. Akad. Nauk Mold. SSR. Ser. Biol. Chim. Nauk, 11: 37-48.

Robbins, R.T.. and Brown, D.J.F. 1991. Comments on the taxonomy, occurrence and distribution of Longidoridae (Nematoda) in North America. Nematologia 37(4): 395-419.

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Romanenko, N.D. 1985. Distribution of Longidoridae nematodes in fruit and berry plantations in the USSR. In: Progressive methods of soft fruit cultivation and breeding. Sbornki Nauch. Truc. Moscov. Pp. 112-120.

Romascu, E. 1971. Xiphinema americanum Cobb, 1913 and Xiphinema italiae Meyl, 1953 (Nematoda: Dorylaimida), new vine pests. Inst. Cercet. Pentru. Prot. Plant An.7: 203-109.

Siddiqui, Z.A., Rashid, A., Farooqi, N., and Bisheya, F. 1987. A survey of plant parasitic nematodes associated with citrus in Libya and trials on chemical control. Indian J. Nematol. 17: 76-80.

SON. n.d. Society of Nematologists. Xiphinema italiae. http://nematode.unl.edu/pest34.htm. Accessed July 13, 2010.

Teliz, D., Landa, B.B., Rapoport, H.F., Perez Camacho, F., Jimenez-Diaz, R.M., and Castillo, P. 2007. Plant-parasitic nematodes infecting grapevine in southern Spain and susceptible reaction to root-knot nematodes of rootstocks reported as moderately resistant. Plant Disease 91: 1147-1154.

Tzortzakakis, E.A., Pateras, D., and Charoulis, A. 2006. Occurrence of Xiphinema species in grapevine areas of Tyranavos with comments on the distribution of X. italiae in Greece. Helminthologia 43(3): 186- 187.

Vovlas, N., and Avgelis, A. 1988. Occurrence and distribution of Xiphinema species in vineyards of the Heraklion Province, Crete (Greece). Nematol. Medit. 16: 197-200.

Wang, X., Bosselut, N., Castagnone, C., Voison, R., Abad, P., and Esmenjaud, D. 2003. Multiplex polymerase chain reaction identification of single individuals of the Longidorid nematodes Xiphinema index, X. diversicaudatum, X. vuittenezi, and X. italiae using specific primers from ribosomal genes. Phytopathology 93: 160-166.

Weiland Ardaiz, C., and Perez Camacho, F. 1995. Nematode vectors of virus in the ‘Denomincacion de origen condada de Huelva. Spain. Acta. Hortic. 388: 31-35.

188 Meloidogyne mali Secondary Pest of Grape Nematode Apple root-knot nematode

Secondary Pests of Grape (Truncated Pest Datasheet)

Meloidogyne mali

Scientific name Meloidogyne mali Itoh, Ohshima, and Ichinohe

Synonyms: None

Common Name(s) Apple root-knot nematode

Type of Pest Nematode

Taxonomic Position Class: Nematoda, Order: Tylenchida, Family: Meloidogynidae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List - 2006 through 2009; Additional pest of concern

Pest Description Figure 1. Male (left) and female Measurements in µm from Itoh et al. (1969). (right) root knot nematode. The Measurements in parentheses indicate the range male nematode remains mobile of measurements observed in the character. after the final molt, whereas the female stays in place Females: Stylet length 15 (13-17), dorsal gland (sedentary), establishes a opening 5.5 (4-7) behind stylet base, excretory feeding site, matures, and lays port 23 (20-29) annules from anterior. Perineal eggs. Photo courtesy of North pattern oval, striae smooth, finely spaced, dorsal Carolina State University. arch low, flat, some pronounced transverse striae toward both ends of vulva, ventral arch flat, tail terminus forming circular striae. Phasmids large, “distinct striae always located bending downwards spanning phasmids.” Lateral fields with single or double incisures.

Males: Length 1447 (1270-1630), a=38 (31-44), c=40 (32-58), stylet length 20 (18-22), dorsal gland opening 8 (6-13) behind stylet base, excretory pore 20 (7-26) annules behind median bulb and one or two annules posterior to hemizonid, spicules 32 (28-35), gubernaculums 8.5 (7-10). Lateral field with four incisures areolated on tail portion.

189 Meloidogyne mali Secondary Pest of Grape Nematode Apple root-knot nematode

Second-stage juveniles: Length 418 (390- 450), a = 28.5 (27-31), c=13.3 (12-15), c’ = 3.7 (3-5), stylet length 14, dorsal gland opening 4.7 (4-6) behind stylet base. Tail length 31 (30-34), conoid, terminus irregular, rounded, unstriated. Rectum not swollen. Lateral field with four incisures, outer bands wider than inner band.

Symptoms/Signs Above ground, new primary shoot growth decreases in number and length. Annual gain in plant height, trunk thickness, and numbers of leaves are reduced. Secondary shoots increase in numbers and length. The above ground symptoms are not specific for M. mali and may be caused by other nematodes or organisms damaging the roots.

Below ground, M. mali produces the characteristic root-knot nematode galls on roots of host plants (Fig. 2). All stages are found associated with the root galls. Eggs, Figure 2. Characteristic root knot infective second-stage juveniles, and males galls caused by Meloidogyne spp. can also be found in the soil. Photo courtesy of AVRDC- The World Vegetable Center. Survey CAPS-Approved Method: Has not been evaluated at this time.

Literature-Based Methods: Soil sampling: Soil samples must be collected and analyzed by a taxonomic expert. In apple orchards, M. mali is most abundant in the top 25 cm of soil. A few nematodes occur 50 cm (19.69 in.) deep. Horizontally, the nematodes are most abundant in two zones, 20 to 40 cm (7.87 to 15.75 in.) and 120 to 160 cm (47.2 to 62.99 in.) around the tree. These patterns of nematode distribution are thought to be related to the development and distribution of apple roots in the soil (USDA, 1987).

A recent risk analysis by USDA-APHIS-PPQ-CPHST shows that portions of California, New York, Pennsylvania, Maryland, Massachusetts, New Jersey, Vermont, and West Virginia have a moderate risk-level based on host availability.

Key Diagnostics/Identification CAPS-Approved Method: Has not been evaluated at this time.

190 Meloidogyne mali Secondary Pest of Grape Nematode Apple root-knot nematode

Literature-Based Methods: Morphological: Males, females with head attached, and second-stage juveniles are needed for species determination. Males and juveniles should be relaxed by gentle heat before fixation in a formalin solution. Females can be fixed without heating. If galled roots are submitted for determination, males and second-stage juveniles should first be extracted by placing the roots in a moist atmosphere for several days. Extracted nematodes can then be relaxed and fixed.

Female M. mali have an oval cuticular perineal pattern with a flat dorsal arch as with M. arenaria. The species can be separated from M. arenaria by a characteristic double ridge of striae, which are present in the inner part of the lateral field.

Molecular: A PCR-RFLP technique has been developed to separate Japanese species of the genus Meloidogyne from Japan (including M. mali) using individual second-stage juveniles (Orui, 1998). The ten Meloidogyne species were best discriminated using either MseI or SspI or VspI+HinfI-digested patterns of the PCR product.

References Itoh, Y., Ohshima, Y., and Ichinohe, M. 1969. A root-knot nematode Meloidogyne mali n. sp. on apple tree from Japan (Tylenchida: Heteroderidae). Applied Entomology and Zoology 4(4): 194-202.

Orui, Y. 1998. Identification of Japanese species of the genus Meloidogyne (Nematoda: Meloidogynidae) by PCR-RFLP analysis.

USDA. 1987. Pests not known to occur in the United States or of limited distribution. No. 83: Apple Root- Knot Nematode.

191 Meloidogyne mali Secondary Pest of Grape Nematode Apple root-knot nematode

192 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows Plant Pathogens

Primary Pests of Grape (Full Pest Datasheet)

‘Candidatus Phytoplasma australiense’

Scientific Name ‘Candidatus Phytoplasma australiense’ Davis et al., 1997

Synonyms Phytoplasma solani australensis, Phytoplasma solani australiense

Common Name(s) Australian grapevine yellows, Australian lucerne yellows, Australian papaya dieback, cabbage tree sudden decline, coprosma lethal disease, cordyline sudden decline, cottonbush witches’ broom, gomphocarpus yellowing, Jaunisse de la Vigne, liquidamber yellows, mung bean witches’ broom, papaya dieback, paulownia yellows, periwinkle phyllody, phormium yellow leaf, pumpkin yellow leaf curl, strawberry green petal, strawberry lethal yellows, and yellow leaf disease.

Type of Pest Phytoplasma

Taxonomic Position Class: Mollicutes, Order: Acheloplasmatales, Family: Acheloplasmataceae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2009 through 2014

Pest Description Mollicutes are prokaryotes that have small genomes (530 to 1350 kbp), lack a cell wall, are pleomorphic, and have a low G + C content (23-29 mol%). Phytoplasmas belong to the class Mollicutes and are the proposed causative agents of diseases in several hundred plant species (McCoy et al., 1989). Phytoplasmas reside in the phloem tissue of the infected plant host and are transmitted by insect vectors, principally leafhoppers and planthoppers (White et al., 1998). Although phytoplasmas have been detected in affected plant tissues and insects with the use of technologies based on the transmission electron microscope, antibodies, and nucleic acids, they are unable to be cultured in vitro. Phytoplasmas cannot be morphologically or ultrastructurally distinguished from one another using either electron or light microscopy (CABI, 2007). Candidatus in scientific classification is a formal word that is placed before the genus and species name of bacteria that cannot be maintained in a Bacteriology Culture

193 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

Collection. Candidatus status may be used when a species or genus is well characterized but unculturable.

‘Candidatus Phytoplasma australiense’ (herein abbreviated as ‘Ca P. australiense’) is the phytoplasma associated with Australian grapevine yellows (AGY), papaya dieback (PD), and phormium yellow leaf diseases (PYL) in New Zealand and Australia (Davis et al., 1997). Molecular studies have shown that the phytoplasmas associated with AGY, PD, and PYL belong to the same species (Liefting et al., 1998). This phytoplasma is distinct from German grapevine yellows and stolbur. PYL is a lethal disease of New Zealand flax (Phormium tenax) and mountain flax (P. cookianum), which is only present in New Zealand (first found in 1908). It is transmitted by a planthopper, Zeoliarus (Oliarus) atkinsoni. PD can be a devastating disease in Queensland, Australia (first found in 1922). Australian grapevine yellows was first reported in Australia in 1975. No vectors have been identified for the latter two diseases.

Taxonomists classify phytoplasmas by directly comparing their 16S ribosomal ribonucleic acid (rRNA) sequences plus the 16S-23S rRNA intergenic spacer region (White et al., 1998). They group isolates that are > 97.5% in the same Candidatus species, unless significant biological or genetic properties suggest that they should be classified separately (Andersen et al., 2006). Andersen et al. (2006) reveal three distinct subgroups within ‘Ca. P. australiense’ by sequencing the tuf gene, which encodes the elongation factor Tu.

Figure 1. Symptoms associated with ‘Candidatus Phytoplasma australiense’ in grape. Mild, irregular chlorosis, leaves with backward curling, overlapping of leaves, tip death (left); necrosis (center); chlorosis along the veins (right). Photos courtesy of Fiona Constable (CABI, 2007).

Biology and Ecology The biology of ‘Ca. P. australiense’ is not completely understood. Like other phytoplasmas, it is an obligate intercellular parasite that occurs in the phloem sieve tubes of infected plants and the salivary glands of insect vectors (CABI, 2007). ‘Ca. P. australiense’ cells, like other phytoplasmas, are surrounded by a single-unit membrane, lack a rigid cell wall, and are pleomorphic in shape. When observed by transmission

194 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

electron microscopy, they appear as rounded to filamentous, pleomorphic bodies with a mean diameter of 200 to 800 nm (IRPCM, 2004). They are sensitive to antibiotics of the tetracycline group but not to penicillin.

The cixiic planthopper, Zeoliarus (Oliarus) atkinsoni and Zeoliarus oppositus, which occur in New Zealand, are the only known vectors of ‘Ca. P. australiense’. Z. atkinsoni has only been demonstrated to transmit the phytoplasma from Phormium plant to Phormium plant (Phormium yellows); while Z. oppositus is polyphagous and has been reported on many different plants (Cumber, 1953; Liefting et al., 1997; Beever et al., 2008). To date the insect vector responsible for the spread of ‘Ca. P. australiense’ in Australia is not known.

Symptoms/Signs Grape: Symptoms include yellow (chlorotic) and downward curled leaves that fall prematurely; reddening may be seen in red varieties (Fig. 1, 2). The chlorotic patches on affected leaves may become necrotic. Leaves of affected shoots can overlap one another. Shoots are stunted and unlignified. Abortion of flowering bunches early in the season has been observed (Constable et al., 2004). Any time from flowering, bunches may shrivel and fall (Magarey and Wachtel, 1986; CABI, 2007). Stems of affected shoots often take on a bluish hue (Constable et al., 2004). Only a few shoots on grapevine are usually affected, and inflorescence and fruit are generally only affected on symptomatic shoots. Later in the season, affected shoots tend Figure 2. Symptoms associated with Candidatus to be green and rubbery Phytoplasma australiense in grape. Irregular (CABI, 2007). The symptoms associated reddening. Photos courtesy of Fiona Constable with ‘Ca. P. australiense’ can be (CABI, 2007). influenced by the environment. Infected grapevines are less likely to show symptoms in summer than winter (CABI, 2007). Although the infected vines are likely to show symptoms year after year, the disease can go into remission and not express symptoms (CABI, 2007).

195 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

Solanaceous Hosts: In potato, upward rolling and purpling of the leaves were observed (Fig. 3a,b). The symptoms appeared similar to those of ‘zebra chip’, a disorder of potato recently found to be associated with ‘Candidatus Liberibacter solanacearum’ in New Zealand and the United States (Liefting et al., 2009).

a) b)

c) Figure 3. Symptoms associated with infection by ‘Candidatus Phytoplasma australiense’ in potato and Jerusalem cherry. a) ‘Ca. Phytoplasma australiense’ infected Ranger Russet potato plant. b) An up close view of the infected potato plant. c) Witches’ broom, foliar yellowing, and reduced leaf size in a Jerusalem cherry shoot on the right compared to a healthy shoot on the left. (Photos a and b courtesy of Lia Liefting. Photo c courtesy of Liefting et al., 2011).

In Jerusalem cherry (Solanum pseudocapsicum), symptoms include: witches’ broom, foliar yellowing, and reduced leaf size (Fig. 3c) (Liefting et al., 2011).

Other hosts: • In alfalfa (lucerne), symptoms range from a yellow to red discoloration of the leaves, a yellowish-brown root discoloration under the periderm, to plant death (Pilkington et al., 2003).

196 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

• In boysenberry, symptoms are obvious close to flowering when the lateral branches become stunted. Young leaves are smaller than normal and chlorotic. As the disease progresses, the older leaves become purple-bronze in color, particularly towards the margin. The fruit set as usual but remain small. New canes fail to grow and the plant is dead by the following winter (Liefting et al., 2011). • In celery, pink and yellow foliage and leaf deformation were observed (Liefting et al., 2011). • Coprosma lethal decline causes leaf reddening and bronzing, heavy leaf loss, dieback, and plant death (Beever et al., 2004). • Cordyline sudden decline may cause the death of mature cabbage trees. • On New Zealand flax, abnormal yellowing of the leaves, stunted growth, increased root death, phloem necrosis, and xylem gummosis of the rhizome vascular system are observed (CABI, 2007). • On papaya, symptoms include dieback, bending of the growing tip, bunching and chlorosis of the crown leaves, followed by necrosis of the young leaves and stem. Laticifer discoloration, particularly in the vicinity of the vascular tissue is evident. Plant death is observed within 2 to 3 weeks of first visible symptoms. • Paulownia yellows stunts plant growth, causes leaves to ‘yellow’, and reduces internode length and leaf size (Bayliss et al., 2005). • In pumpkin and strawberry plant growth is stunted and leaves turn yellow and curl (roll). • In red clover, diminished leaf size, pallor, rugosity, leaf deformation shoot proliferation, and severe stunting were observed (Saqib et al., 2006). • In swan plant (Gomphocarpus fruticosa), symptomatic plants exhibited abnormal foliar yellowing and slight upward rolling of the leaves. The symptoms eventually resulted in plant death (Liefting et al., 2011). • In sweetgum, the crown may have patchy chlorosis, chlorotic shoots with comparatively few leaves, dieback of apical and lateral branches, small leaves showing tip necrosis, and reduced fruit production (Habili et al., 2006).

Note: While phytoplasma infections are usually detrimental to plant growth, some plants exhibit minor symptoms or are symptomless.

Pest Importance Diseases caused by ‘Ca. P. australiense’ impact economically important food and ornamental crops. Researchers have documented vineyard losses as high as 13%. Severely affected grape vines can produce up to 54% less fruit than healthy grape vines (CABI, 2007; NPAG, 2007). Papaya dieback is responsible for annual plant losses of 10% and up to 100% during epiphytotics (epidemic among plants of a single kind over a wide area) in central and southern Queensland plantations (Glennie and Chapman, 1976; Guthrie et al., 1998). Australian lucerne yellows has caused a reduction in seed yield, which has led to the cutting or plowing-under of seed crops, resulting in estimated losses of $7 million annually (Pilkington et al., 1999).

197 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

The economic impact that ‘Ca. P. australiense’ could have on new host Solanum tuberosum is potentially significant. The total economic value of New Zealand’s potato industry in 2011 was estimated to be $382 million NZD ($300 million USD), with about a quarter of that coming from exports (Potatoes New Zealand, n.d.).

The economic impact that ‘Ca. P. australiense’ will have on its other newly identified hosts is potentially significant (Liefting et al., 2011). Already, phytoplasma-infected potato tubers have failed quality control checks at the processing factory. In boysenberry, fruit from infected plants are unmarketable, and the plants die within a year after symptoms are first noticeable. Although there have been no reports of the phytoplasma in commercially grown celery, the discoloration of the foliage that the phytoplasma induces in this crop would also render it unmarketable. The polyphagous feeding behavior of Zeolarius oppositus, the insect responsible for transmitting the phytoplasma into Cordyline and Coprosma, suggests that it may also be moving the phytoplasma into the new hosts described here. Z. oppositus is especially abundant in grasses and sedges that commonly grown around crops in New Zealand. These hosts may act as symptomless reservoirs of the phytoplasma, and along with weed hosts such as Jerusalem cherry, play an important role in the spread of the phytoplasma. The diversity of the new hosts described in the Liefting et al. (2011) paper emphasize the potential that ‘Ca. P. australiense’ has to spread into additional native plants and horticultural crops.

Some of the diseases caused by ‘Ca. P. australiense’ are ‘new’ diseases or recently described (see known hosts section). For example, cordyline sudden death became a concern in the late 1980s; apparently this phytoplasma jumped to other hosts, including cabbage trees. The ability to use new hosts is an important and threatening aspect of the pathogen (NPAG, 2007). Establishment of ‘Ca. P. australiense’ in the United States could impact trade because some countries, like Morocco, classify the pathogen as a dangerous quarantine pest (NPAG, 2007). It is also on the harmful organism listing for China, Japan, and Peru.

Known Hosts Research implicates ‘Ca. P. australiense’ as the cause of disease on the following hosts:

Asclepias physocarpa (balloon plant), Apium graveolens (celery), Carica papaya (papaya), Catharanthus roseus (periwinkle), Cicer arietinum (chickpea), Coprosma macrocarpa (coprosma), Coprosma robusta (coprosma), Cordyline australis (cabbage tree), Cordyline banksii (cabbage tree), Cucumis myriocarpus (paddy melon), Cucurbita maxima (pumpkin), Cucurbita moschata (pumpkin), Exocarpos (Exocarpus) cupressiformis (cherry ballart), Fragaria ananassa (strawberry), Fragaria virginiana (strawberry), Gomphocarpus fruticosa (also cited as G. fructicosus) (cottonbush; swan plant), Jacksonia scoparia (winged broom pea), Liquidambar (Liquidamber) styraciflua (sweetgum), Medicago sativa (alfalfa), Melilotus indicus (hexham scent), Paulownia fortunei (paulownia), Phaseolus vulgaris (bean), Phormium cookianum (mountain flax), Phormium tenax (New Zealand flax), Rubus loganobaccus (logan berry), Rubus ursinus

198 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

(boysenberry), Solanum pseudocapsicum (Jerusalem cherry), Solanum tuberosum (potato), Trifolium pratense (red clover), Vigna radiata (mung bean), and Vitis vinifera (grape) (EPPO, 1998; Liefting et al., 1998; Padovan et al., 1998, White et al., 1998; Schneider et al., 1999; Wood et al., 1999; Padovan et al., 2000; Andersen et al., 2001; Davis et al., 2003; Beever et al., 2004; Bayliss et al., 2005; Lucas, 2005; Saqib et al., 2005; Streten and Gibb, 2005; Streten et al., 2005; Habili et al., 2006; Saqib et al., 2006; CABI, 2007; Getachew et al., 2007; Liefting et al., 2009; CABI, 2011; Liefting et al., 2011).

Jones et al. (2005) identified ‘Ca. P. australiense’ on peach in Bolivia. This report is now thought to be a related-species and not ‘Ca. P. australiense’ (USDA-APHIS-PPQ- CPHST-PERAL, 2012; Davis, 2013). A ‘Ca. P. australiense’- related strain was found in China on Senna surattensis (sunshine tree) (Wu et al., 2012).

Known Vectors (or associated insects) The only known vectors of ‘Ca. Phytoplasma australiense’ are Zeoliarus (Oliarus) atkinsoni and Zeoliarus oppositus (not known in the United States) (Liefting et al., 1997; Beever et al., 2008). Other vectors are suspected but not currently confirmed (NPAG, 2007, CABI, 2011). The vector responsible for spread of ‘Ca. P. australiense’ in Australia is unknown.

Known Distribution ‘Ca P. australiense’ is present in New Zealand and Australia. A ‘Ca. P. australiense’- related strain was found in China on Senna surattensis (sunshine tree) (Wu et al., 2012).

Note: Two incorrect reports of 'Ca. P. australiense' occur in the literature: Nivum Haamir dieback of papaya in Israel (Gera et al., 2005) and yellow leaf roll of peach in Bolivia (Jones et al., 2005). In both cases, the 16S rRNA gene sequence did not fulfill the criteria to be classified as 'Ca. P. australiense' (USDA-APHIS-PPQ-CPHST-PERAL, 2012).

Pathway The only confirmed vectors of ‘Ca. Phytoplasma australiense’ are Zeoliarus (Oliarus) atkinsoni and Zeoliarus oppositus (Liefting et al., 1997; Beever et al., 2008). Neither of these pests has been found in the United States. It is possible for ‘Ca. Phytoplasma australiense’ to be moved within infected vectors, particularly the polyphagous vector Zeoliarus oppositus. There are no interceptions of these two leafhoppers at the species level to date in the United States (PestID, 2013). Zeoliarus is not currently listed in Pest ID (PestID, 2013).

The current pathway of ‘Ca. P. australiense’ infection in Solanum spp. remains unknown. One possible pathway is through Zeoliarus oppositus. Infected Z. oppositus was found in the same field as the infected potato and Jerusalem cherry (Liefting et al., 2011). Transmission of the phytoplasma into potato by Z. oppositus has not been demonstrated experimentally, but these planthoppers were observed on the potato

199 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

foliage and in one instance exhibited mating behavior (Liefting et al., 2011). It is possible that Jerusalem cherry could harbor the phytoplasma, which is then spread into the potato crop by the Zeoliarus planthopper.

Spread through infected plants for planting, such as nursery stock and cuttings, is also possible (CABI, 2011). This phytoplasma pathogen is not known to be seedborne. Vitis sp. and Solanum sp. (all propagules except seeds) are prohibited from all countries except for Canada when meeting the required import conditions (Plants for Planting Manual, 2012). There have been nine interception records (two destined for propagation) of Vitis sp. leaves, cutting, and fruit in baggage and permit cargo from Australia. There were an additional five interception records (two destined for propagation) of Vitis sp. stems and fruit in permit cargo, baggage, and quarters from New Zealand. There has also been four interception records (one destined for propagation) of Solanum sp. from New Zealand. There have been multiple interceptions (351 from Australia and 778 from New Zealand) of all currently known hosts of ‘Ca. Phytoplasma australiense’ with a total of 69 destined for propagation (PestID, 2013).

Coprosma, Cordyline, Paulownia, and Phormium spp. propagative material (PM) does not appear to be subject to regulations, and there appears to be an open pathway for this PM (Plants for Planting Manual, 2012). Shipments appear to be plant units, seed, and flasks. There were 3 and 34 shipments of shipments of Coprosma sp. PM from Australia and New Zealand, respectively, since January of 2003. There were an additional 53 and 115 shipments of shipments of Cordyline sp. PM and 43 and 249 shipments of Phormium sp. from Australia and New Zealand respectively during the same time period. There were 15 shipments of Paulownia sp. PM from Australia over the past ten years as well (Plants for Planting Manual, 2012).

Potential Distribution within the United States ‘Ca. P. australiense’ is considered an imminent threat that could be introduced into the United States with imported leaves, stems, and roots of infected plants (NPAG, 2007). A recent risk analysis by USDA-APHIS-PPQ-CPHST shows that portions of the southeastern United States are at the greatest risk from this phytoplasma based on host availability. Most of the United States has a low to moderate risk of ‘Ca. P. australiense’ establishment.

Survey CAPS-Approved Method*: Visual survey is the method to survey for ‘Ca. P. australiense’ by collecting symptomatic plant material. Be sure that each plant that is sampled exhibits shriveling of the fruiting cluster when sampling grape. Follow instructions in Phytoplasma sample submission for Cooperative Agricultural Pest Survey (CAPS) Program and Farm Bill Goal 1 surveys FY 2014.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

200 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

Literature-Based Methods: Visual survey: Conduct visual inspection for symptoms associated with the phytoplasma. Several of the known symptoms should be found together before suspecting a phytoplasma infection on grape (CABI, 2007). Symptoms begin to appear in late spring and increase in incidence until January/February in Australia (their summer). Beyond this time, AGY symptoms begin to disappear as symptomatic leaves and shoots fall from the vine (CABI, 2007). Phytoplasma DNA was detected from most plants when samples were collected in January and February (summer) compared to those sampled in Oct-Dec. (spring to early summer) and March-May (fall) in Australia (Gibb et al., 1999). The disease appears most often in Chardonnay and Riesling grapes but has also been reported in other cultivars (Magarey and Watchel, 1986).

Key Diagnostics/Identification CAPS-Approved Method*: Follow instructions in Phytoplasma sample submission for Cooperative Agricultural Pest Survey (CAPS) Program and Farm Bill Goal 1 surveys FY 2014.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: A 'universal' PCR assay has been developed that enables amplification of the 16S rRNA genes of phytoplasmas. Digestion of these PCR products with selected restriction enzymes provides a DNA fingerprint in the form of 16S rDNA fragment patterns that can be used to determine phytoplasma identity (Ahrens and Seemuller, 1992; Deng and Hiruki, 1991; Lee et al., 1993; Smart et al., 1996; Gibb et al., 1999). PCR assays have been developed to detect only the AGY phytoplasma in grape (Davis et al., 1997; Gibb et al., 1999).

The chromosome sequence of ‘Ca. P. australiense’ isolates from papaya and grapevine in Australia is available (Tran-Nguyen et al., 2008).

Easily Confused Pests AGY resembles flavéscence dorée, bois noir, Goldgelbe Vergilbung, leaf curl, berry shrivel and other grapevine diseases (tomato big bud and an uncharacterized disease also believed to be caused by a phytoplasma) (Magarey and Wachtel, 1985; Davis et al., 1997; Lee et al., 1998; Constable et al., 1998; Gibb et al., 1999). Detection of flavéscence dorée by cloned DNA probes was accomplished by Daire et al. (1992).

Mechanical disruption to the phloem of grape shoots can cause symptoms similar to those associated with ‘Ca. P. australiense’ infection (CABI, 2007). It is important to inspect symptomatic shoots for damage to the vascular tissue due to breakage, restrictions of the vascular tissue due to tendrils or string wrapping tightly around shoots, and damage to the vascular tissue by boring insects.

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References Ahrens, U., and Seemuller, E. 1992. Detection of DNA of plant pathogenic mycoplasma-like organisms by a polymerase chain reaction that amplifies a sequence of the 16S rRNA gene. Phytopathology 82(8): 828-832.

Andersen, M.T., Beever, R.E., Sutherland, P.W., and Forster, R.L.S. 2001. Association of ‘Candidatus Phytoplasma australiense’ with sudden decline of cabbage tree in New Zealand. Plant Disease 85: 462- 469.

Andersen, M.T., Newcomb, R.D., Liefting, L.W. and Beever, R.E. 2006. Phylogenetic analysis of ‘Candidatus Phytoplasma australiense’ reveals distinct populations in New Zealand. Phytopathology 96: 838-845.

Bayliss, K.L., Saqib, M., Dell, B., Jones, M.G.K., and St. J. Hardy, G.E. 2005. First record of ‘Candidatus Phytoplasma australiense’ in Paulownia trees. Australian Plant Pathology 34: 123-124.

Beever, R.E., Wood, G.A., Andersen, M.T., Pennycook, S.R., Sutherland, P.W., Forster, R.L.S. 2004. ‘Candidatus Phytoplasma australiense’ in Coprosma robusta in New Zealand. New Zealand Journal of Botany 42: 663-675.

Beever, R.E., Andersen, M.T., and Winks, C.J. 2008. Transmission of ‘Candidatus Phytoplasma australiense’ to Cordyline australis and Coprosma robusta. J. Plant. Pathol 90(S2): 459.

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

CABI. 2011, Crop Protection Compendium: Phytoplasma australiense (Australian grapevine yellows). Wallingford, UK: CAB International. Retrieved April 30, 2012, from http://www.cabi.org/cpc/?compid=1&dsid=39956&loadmodule=data.

Constable, F.E., Gibb, K.S., Moran, J.R., and Wilson, Y.M. 1998. Incidence of phytoplasma associated with yellows, restricted spring growth and late season leaf curl symptoms of grapevines. Australian Grape grower and Winemaker 409: 19-20.

Constable, F.E., Jones, J., Gibb, K.S., Chalmers, Y.M., and Symons, R.H. 2004. The incidence, distribution and expression of Australian grapevine yellows, restricted growth and late season leaf curl in selected Australian vineyards. Annals of Applied Biology 205-218.

Cumber, R.A. 1953. Investigations into yellow-leaf disease of Phormium. IV. Experimental induction of yellow-leaf condition on Phormium tenax Forstl by the insect vector Oliarus atkinsoni Myers (Hem. Cixiidae). NZ J. Sci. Technol. 34A(1): 31-40.

Daire, X., Boudon-Padieu, E., Berville, A., Schneider, B., and Caudwell, A. 1992. Cloned DNA probes for the detection of grapevine flavescence dorée mycoplasma-like organism (MLO). Ann. Appl. Biol. 121: 95-103.

Davis, R.E. 2013. Personal Communication. Email received 6/21/2013.

Davis, R.E., Dally, E.L., Gundersen, D.E., Lee, I.M., and Habili, N. 1997. ‘Candidatus Phytoplasma australiense,’ a new phytoplasma taxon associated with Australian grapevine yellows. International Journal of Systematic Bacteriology 47(2): 262-269.

Davis, R.I., Jacobson, S.C., De La Rue, S.J., Tran-Nguyen, L., Gunua, T.G., and Rahamma, S. 2003. Phytoplasma disease surveys in the extreme north of Queensland, Australia, and the island of New Guinea. Australian Plant Pathology 32: 269-277.

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Deng, S., and Hiruki, C. 1991. Amplification of 16S rRNA from culturable and non-culturable mollicutes. Journal of Microbiological Methods 14: 53-61.

EPPO. 1998. ‘Candidatus Phytoplasma australiense’. A document prepared by EPPO (European and Mediterranean Protection Organization). http://www.eppo.org/QUARANTINE/Alert_List/deleted%20files/bacteria/Phytoplasma_australiense.doc.

Gera, A., Mawassi, M., Zeidan, M., Spiegel, S., and Bar-Joseph, M. 2005. An isolate of ‘Candidatus Phytoplasma australiense’ group associated with Nivun-Haamir-Die Back. Plant Pathology 54: 560.

Getachew, M.A., Mitchell, A., Gurr, G.M., Fletcher, M.J., Pilkington, L.J., and Nikandrow, A. 2007. First report of a ‘Candidatus Phytoplasma australiense’ related strain in lucerne (Medicago sativa) in Australia. Plant Disease 91: 111.

Gibb, K.S., Constable, F.E., Moran, J.R., and Padovan, A.C. 1999. Phytoplasmas in Australian grapevines – detection, differentiation and associated diseases. Vitis 38(3): 107-114.

Glennie, J.D., and Chapman, K.R. 1976. A review of dieback – a disorder of the papaw (Carica papaya L.) in Queensland. Queensland Journal of Agriculture and Animal Science 33: 177-188.

Guthrie, J.N., White, D.T., Walsh, K.B., and Scott, P.T. 1998. Epidemiology of phytoplasma-associated papaya diseases in Queensland, Australia. Plant Disease 82: 1107-1111.

Habili, N., Farrokhi, N., and Randles, J.W. 2006. First detection of Candidatus Phytoplasma australiense in Liquidambar styraciflua in Australia. New Disease Reports 14: August 2006 – January 2007. http://www.bspp.org.uk/NDR/jan2007/2006-62.asp.

IRPCM Phytoplasma/Spiroplasma Working Team, P.T.G. 2004. ‘Candidatus Phytoplasma’, a taxon for the wall-less, non-helical prokaryotes that colonize plant phloem and insects. Int. J. Syst. Evol. Microbiol. 54: 1243-1255.

Jones, P., Arocha, Y., Antesana, O., Montilliano, E., and Franco, P. 2005. First report of an isolate of ‘Candidatus Phytoplasma australiense’ associated with a yellow leaf roll disease of peach (Prunus persicae) in Bolivia. Plant Pathology 54: 558.

Lee, I.M., Hammond, R.W., Davis, R.E., Gundersen, D.E. 1993. Universal amplification and analysis of pathogen 16S rDNA for classification and identification of mycoplasmalike organisms. Phytopathology 83(8): 834-842.

Lee, I.M., Gundersen-Rindal, D.E., Davis, R.E., and Bartoszyk, I.M. 1998. Revised classification scheme of phytoplasmas based on RFLP analyses of 16S rRNA and ribosomal protein gene sequences. International Journal of Systematic Bacteriology 48(4): 1153-1169.

Liefting, L.W., Beever, R.E., Winks, D.J., Pearson, M.N., and Forster, R.L.S. 1997. Planthopper transmission of Phormium yellow leaf phytoplasma. Australasian Plant Pathology 26: 148-154.

Liefting, L.W., Padovan, A.C., Gibb, K.S., Beever, R.E., Andersen, M.T., Newcomb, R.D., Beck, D.L., and Forster, R.L.S. 1998. Candidatus Phytoplasma australiense is the phytoplasma associated with Australian grapevine yellows, papaya dieback, and Phormium yellow leaf diseases. European Journal of Plant Pathology 104: 619-623.

Liefting, L.W., Verrakone, S., Ward, L.I., and Clover, G.R.G. 2009. First report of ‘Candidatus Phytoplasma australiense’ in potato. Plant Dis. 93(9): 969.

203 Ca. Phytoplasma australiense Primary Pest of Grape Phytoplasma Grapevine yellows

Liefting, L.W., Veerakone, S., and Clover, G.R.G. 2011. New hosts of ‘Candidatus Phytoplasma australiense’ in New Zealand. Australasian Plant Pathology. DOI 10.1007/s13313-011-0036-z. http://www.springerlink.com/content/q7r772452m66042l/fulltext.pdf.

Lucas, R. 2005. Managing Pests and Disease: A Handbook for New Zealand Gardeners. Craig Potten Publishing.

Magarey, P.A., and Wachtel, M.F. 1986. Grapevine yellows, a widespread, apparently new disease in Australia. Plant Disease 70: 694.

McCoy, R.E., Caudwell, A., Chang, C.J., Chen, T.A., Chykowski, L.N., Cousin, M.T., Dale, J.L., de Leeuw, G.T.N., Golino, D.A., Hackett, K.J., Kirkpatrick, B.C., Marwitz, R., Petzold, H., Sinha, R.C., Sugiura, M., Whitcomb, R.F., Yang, I.L., Zhu, B.M., and Seemuller E. 1989. Plant diseases associated with mycoplasma-like organisms. In: The Mycoplasmas, Vol. 5. Whitcomb, R.F., and Tully, J.G. (ed.). Academic Press, New York.

New Pest Advisory Group (NPAG). 2007. Candidatus Phytoplasma australiense. USDA-APHIS-PPQ- CPHST.

Padovan, A., Gibb, K., Persley, D. 1998. Phytoplasmas associated with diseases in strawberry. Australasian Plant Pathology 27: 280.

Padovan, A., Gibb, K., and Persley, D. 2000. Association of ‘Candidatus Phytoplasma australiense’ with green petal and lethal yellows diseases in strawberry. Plant Pathology 49: 362-369.

PestID. 2013. Pest Identification Database (PestID). United States Department of Agriculture, Animal and Plant Health Inspection Service, Plant Protection and Quarantine. Last accessed June 21, 2013. https://mokcs14.aphis.usda.gov/aqas/login.jsp.

Pilkington, L., Gurr, G.M., Fletcher, M.J., Nikandrow, A., and Elliot, E. 1999. Occurrence and severity of lucerne yellows diseases in Australian lucerne seed crops. Australasian Plant Pathology 28: 235-239.

Pilkington, L.J., Gibb, K.S., Gurr, G.M., Fletcher, M.J., Nikandrow, A., Elliot, E., van de Ven, R., and Read, D.M.Y. 2003. Detection and identification of a phytoplasma from lucerne with Australian lucerne yellows disease. Plant Pathology 52: 754-762.

Plants for Planting Manual. 2012. United States Department of Agriculture. Last updated June, 2012. http://www.aphis.usda.gov/import_export/plants/manuals/ports/downloads/plants_for_planting.pdf.

Potatoes New Zealand. n.d. http://www.potatoesnz.co.nz.

Saqib, M., Bayliss, K.L., Dell, B., Hardy, G. E. St., and Jones, M.G.K. 2005. First record of a phytoplasma-associated disease of chickpea (Cicer arientinum) in Australia. Australasian Plant Pathology 34: 425-426.

Saqib, M., Jones, M., and Jones, R. 2006. ‘Candidatus Phytoplasma australiense’ is associated with diseases of red clover and paddy melon in southwest Australia. Australian Plant Pathology 35(1): 283- 285.

Schneider, B., Padovan, A., De La Rue, S., Eichner, R., Davis, R., Bernuetz, A., and Gibb, K. 1999. Detection and differentiation of phytoplasmas in Australia: an update. Aust. J. Agric. Res. 50: 333-342.

Smart, C.D., Schneider, B., Blomquist, C.L., Guerra, L.J., Harrison, N.A., Ahrens, U., Lorenz, K.H., Seemuller, E., and Kirkpatrick, B.C. 1996. Phytoplasma-specific PCR primers based on sequences of the 16S-23S rRNA spacer region. Applied and Environmental Microbiology 62(8): 2988-2993.

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Streten, C., and Gibb, K.S. 2005. Genetic variation in Candidatus Phytoplasma australiense. Plant Pathology 54: 8-14.

Streten C., Conde, B., Herrington, M., Moulden, J., and Gibb, K. 2005. Candidatus Phytoplasma australiense is associated with pumpkin yellow leaf curl disease in Queensland, Australia, and the Northern Territory. Australian Plant Pathology 34: 103-105.

Tran-Nguyen, L.T., Kube, M. Schneider, B., Reinhardt, R., and Gibb, K.S. 2008. Comparative genome analysis of ‘Candidatus Phytoplasma australiense’ (subgroup tuf-Australia I; rp-A) and ‘Ca. Phytoplasma asteris’ strains OY-M and AY-WB. J. Bacteriol. 190(11): 3979-3991.

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Wu, W., Cai, H., Wei, W., Davis, R.E., Lee, I.-M., Chen, H., and Zhao, Y. 2012. Identification of two new phylogenetically distant phytoplasmas from Senna surattensis plants exhibiting stem fasciation and shoot proliferation symptoms. Annals of Applied Biology 160: 25-34.

205 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

‘Candidatus Phytoplasma vitis’

Scientific Name ‘Candidatus Phytoplasma vitis’ IRPCM 2004

Note: The provisional name ‘Candidatus Phytoplasma vitis’ has been suggested to denote a phytoplasma associated with flavescence dorée disease in Europe, but the name has yet to be formally published and accepted.

Common Name(s) FD, flavescence dorée phytoplasma, flavescence dorée de la Vigne, flavescencia dorada de la viña, flavescenza dorata de la vite, grapevine "flavescence dorée" phytoplasma, grapevine yellows.

Some older papers refer to the disease as grapevine bois noir. It was demonstrated, however, that this disease is caused by a distinct phytoplasma ‘Candidatus Phytoplasma solani’ or the bois noir phytoplasma (Daire et al., 1993; Quaglino et al., 2013).

Type of Pest Phytoplasma

Taxonomic Position Class: Mollicutes, Order: Acholeplasmatales, Family: Incertae sedis, Genus: “Candidatus Phytoplasma”

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List - 2014

Background Information Phytoplasmas, formerly known as mycoplasma-like organisms (MLOs), are pleomorphic, cell wall-less bacteria with small genomes (530 to 1350 kbp) of low G + C content (23-29%). They belong to the class Mollicutes and are the putative causal agents of yellows diseases that affect at least 1,000 plant species worldwide (McCoy et al., 1989; Seemüller et al., 2002). These minute, endocellular prokaryotes colonize the phloem of their infected plant hosts as well as various tissues and organs of their respective insect vectors. Phytoplasmas are transmitted to plants during feeding activity by their vectors, primarily leafhoppers, planthoppers, and psyllids (IRPCM, 2004; Weintraub and Beanland, 2006).

Although phytoplasmas cannot be routinely grown by laboratory culture in cell free media, they may be observed in infected plant or insect tissues by use of electron microscopy or detected by molecular assays incorporating antibodies or nucleic acids.

206 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

Since biological and phenotypic properties in pure culture are unavailable as aids in their identification, analysis of 16S rRNA genes has been adopted instead as the major basis for phytoplasma taxonomy. The provisional taxonomic status of ‘Candidatus’, used for incompletely described microorganisms, has been adopted for describing and naming distinct phytoplasmas (i.e., ‘Candidatus Phytoplasma’). Several species (i.e., ‘Ca. Phytoplasma’ species) have been named following established guidelines (IRPCM, 2004; Zhao et al., 2009; Harrison et al., 2011; Davis et al., 2013; Quagliano et al., 2013).

Phytoplasmas are classified in a system of groups and subgroups based upon DNA fingerprints (RFLP patterns) of 16S rRNA genes (16S rDNA) (Lee et al., 1998, 2000; Wei et al., 2008). Most of the 16S rDNA RFLP groups contain at least one phytoplasma species (Zhao et al., 2009). For example, ‘Candidatus Phytoplasma vitis’ is classified in group 16SrV, subgroups C & D (16SrV-C and 16SrV-D).

A new ‘Candidatus Phytoplasma’ species may be recognized if the nucleotide sequence of 1,200 bases of its 16S rRNA gene shares < 97.5 identity with that of all previously named ‘Candidatus Phytoplasma’ species (IRPCM, 2004). If a phytoplasma shares >97.5% nucleotide sequence identity of 16S rDNA with any previously named species, the subject phytoplasma may be named as a distinct new species if significant biological or genetic properties distinguish the phytoplasma from already named species (IRPCM, 2004).

Pest Description ‘Candidatus Phytoplasma vitis’ (herein abbreviated ‘Ca. P. vitis’) infection causes flavescence dorée (FD), a very serious grapevine yellows disease in vineyards and a quarantine pest in Europe. The most common damage associated with FD is the significant loss of grape and wine production due to the progressive decline of the plants. In most cases, especially in the more sensitive varieties, the infected grapevines die within a few years. FD is caused by several strains which belong to the 16SrV-C and –D phytoplasma phylogenetic subgroups. Strains classified in 16SrV-D subgroup are most widespread (Arnaud et al. 2007; Filippin et al., 2009).

Biology and Ecology Like other phytoplasmas, ‘Ca. P. vitis’ is an obligate intracellular parasite that occurs in the phloem sieve tubes of infected plants (CABI, 2007). In general, phytoplasmas are transmitted to plants in a circulative-propagative manner by phloem-feeding insect vectors. Their ingestion of sap from diseased plants is followed by an incubation/latency phase lasting for one to several weeks during which time these bacteria circulate, multiply, and parasitize various tissues and organs of their respective vectors. Once salivary glands have been colonized, vectors are then capable of transmitting phytoplasmas during any subsequent feeding activity for their remaining lifespan (Weintraub and Beanland, 2006; Gitau et al., 2009). Phytoplasmas are also transmitted by grafting of infected material.

207 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

The confirmed vector for FD is the leafhopper Scaphoideus titanus Ball (S. titanus), commonly known as the American grapevine leafhopper (Schvester et al., 1961). For FD, the incubation/latency period in S. titanus is 4 weeks. By feeding on an infected grapevine and then feeding on other grapevines, the vector spreads the disease through the vineyard. S. titanus is responsible for the rapid spread of FD in France and in Europe (Papura et al. 2009). Long-distance dissemination is, however, largely the result of the transport of infected propagative material by humans. S. titanus is native to the United States and was likely introduced in Europe when American rootstocks were imported as a part of the fight against downy mildew and phylloxera in the early twentieth century (Caudwell et al., 1983; Papura et al., 2012). Recent genomic studies have demonstrated that the phytoplasma strains responsible for FD originated in Europe (Arnaud et al., 2007, Malembic-Maher et al., 2011). It was only with the recent introduction from North America of the vector Scaphoideus titanus that FD was able to rapidly spread among the Vitis vinifera grapevines (INRA, 2013).

Recent studies carried out on the leafhopper S. titanus demonstrate that cold winter temperatures are not essential for terminating diapause although they do affect male– female ratios over time. The studies also showed that winter temperatures regulate hatching dynamics (Chuche and Thiery, 2012). These findings are very important because first hatchings can now be forecast and can be used to better control larvae before they have the chance to acquire the phytoplasma. The findings also allow predictions of the ability of the vector to colonize new wine-growing areas (INRA, 2013).

Like many other plant-eating insects, sucking insects in particular, S. titanus displays a strong attraction to the color yellow. It appears that the attraction is associated with young, developing leaves that are richer in nitrogen and thus of greater potential nutritional value for the insect. S. titanus is attracted by the color green to a lesser, but still significant, extent. Yellow and, to a larger degree, green are precisely the colors most present on grapevines. The insect’s attraction to these colors is crucial for the propagation of FD. Insects are more attracted to the yellowed leaves of white varieties infected with FD, thereby spurring a vicious cycle (INRA, 2013).

Choice tests carried out with the white varietal Baco 22A, a widely planted variety in Armagnac, France that is highly susceptible to FD, confirmed that plants infected with FD attract more S. titanus, both at larval and adult stages. The attractiveness of infected plants may in part serve to explain the rapid spread of the disease (INRA, 2013).

There is some disagreement as to whether or not Clematis vitalba and Alnus spp. (alder) are “natural hosts” for the FD phytoplasma. There is also disagreement as to whether or not Dictyophara europaea and Oncopsis alni are natural vectors of the FD phytoplasma. Phytoplasmas genetically very similar to the FD phytoplasmas (group 16SrV-C) have been found in both hosts and can rarely be transmitted to grapevine (Malembic-Maher/ Foissac personal communication; Maixner personal communication). Filippin et al. (2009) showed that an FD-related phytoplasma, referred to in the paper as FD, can be occasionally transmitted from infected Clematis vitalba plants to grapevines by the planthopper Dictyophara europaea, commonly known as the European lantern fly

208 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

(exotic to the United States). FD/FD-related genotypes were also detected in alders (Malembic-Maher et al., 2007; Ember et al., 2011; Mehle et al., 2011), and it was shown that the leafhopper Oncopsis alni is able to occasionally transmit alder yellows phytoplasmas (16SrV-C) to grapevine (Maixner et al., 2000). The FD phytoplasma by definition can be transmitted by S. titanus. The genetically similar phytoplasmas to FD in alder and clematis have not been demonstrated to be transmitted by S. titanus. Thus, these phytoplasmas will be referred to as FD-related in this document.

Symptoms/Signs There has not been a demonstrated correlation between different symptoms and early or late infection of the plant. Some plants can express symptoms early in the season; while others express symptoms later. The cultivar, the climate, and the age of the plant are factors that can induce differences in symptoms expression. The FD phytoplasma occurs in vine tissue with an irregular distribution and at a low titre (Boudon-Padieu, 2002), which can make detection of the disease more difficult. Infection of plants by S. titanus typically occurs during mid-to-late summer, but the disease symptoms are usually not expressed until the following year (Boudon-Padieu, 2002).

In general, symptoms resemble those of other grapevine yellows diseases (“leaf rolling, discoloration of lamina and veins, partial or total lack of reserve accumulation (lignification) with flexuous canes, and decline of a part or the entire vine stock”) (Boudon-Padieu, 2002).

Shoots of susceptible cultivars fail to ripen and are thin, rubbery, and hang down. The infected shoots become brittle and many small, black pustules develop along their length; buds may become necrotic. In more resistant cultivars, the nodes of infected shoots ripen but some of the internodes do not (Defra, 2000). During the winter, infected shoots become black. These shoots may die or survive to produce some spring growth depending upon the plant cultivar, plant age, and climatic conditions (Defra, 2000).

Red-fruited cultivars develop reddish discolorations of the leaves and leaf veins (Fig 1a- b). White-fruited cultivars develop a metallic-yellow discoloration on areas of leaf exposed to the sun (Fig 1c-d). Small, discrete, yellowish spots appear along the main veins. These spots enlarge and merge to form yellow bands which gradually spread over the leaf to produce discolored, angular areas delimited by leaf veins. The centers of the discolored areas become dry and brittle. Infected leaves often roll downward but remain on the plant longer than healthy leaves, because they are less affected by autumn frosts. They can, however, be more easily detached by the wind. Blossoms dry out and may fall from vine. Grape bunches become brown and shriveled (Fig 1e); the stalks dry out and the grapes are easily dislodged (Defra, 2000).

In Clematis vitalba, symptoms of FD-related phytoplasmas may include discoloration, rolling and wilting of leaves (Fig. 2), although the relationship between the presence of phytoplasma and leaf symptoms is not clear. PCR detection showed positive results only in DNA extracts for clematis plants sampled in autumn months and never in

209 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

a) b)

c) d)

e) Figure 1: a) a “Cabernet

Sauvignon” red grape plant

infected with ‘Ca. P. vitis’ shows

reddening of leaves. b) a

symptomatic leaf of another

infected red grape cultivar c) an

infected “Sémillon” white grape

cultivar shows yellowing of

leaves. d,e) Drying of the grapes

and shriveling of the fruits in

infected “Sémillon” and “Cabernet

Sauvignon” cultivars. Photo c

courtesy of Michael Maixner,

Julius Kühn-Institut (JKI). Photos

a, b, d, and e courtesy of of S.

Malembic-Maher, INRA

210 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

samples collected in July and August. This could reflect a low concentration or, indeed, absence of the pathogen early in the season (Angelini et al., 2004). A study by Filippin et al. (2009) showed that a FD-related phytoplasma is widespread in C. vitalba, with approximately 36% of plants surveyed throughout Europe testing positive.

Approximately 60 to 80% of alder plants (Alnus sp.) in Europe are infected with an FD-related phytoplasmas. Alders, however, are rarely symptomatic (Foissac, personal communication) (Fig.3).

One of the difficulties in detecting the disease is that symptoms do not always appear every year and may only be present on Figure 2: Infected one shoot or on a small number of shoots. In addition, Vitis symptomatic Clematis vinifera varieties are not equally susceptible to FD and may not vitalba by FD-related present symptoms with equal intensity. Rootstocks which are phytoplasma in hybrids of various American Vitis sp. present very discreet Hungary. Photo symptoms or no symptoms at all. They nevertheless are courtesy of S. carriers of the disease (Caudwell et al., 1994; INRA, 2013). Malembic-Maher, INRA Bordeaux. Pest Importance FD has greatly negative economic impacts among grapevine yellows causing phytoplasmas (Hren et al., 2007). FD is recognized by the European and Mediterranean Plant Protection Organization (EPPO) as a quarantine pest (EPPO, 2012). FD causes symptoms that are detrimental to infected grapevines. Depending on the intensity of infection the yields may decrease dramatically. Their vitality is affected, yields are reduced, and the quality of wine is decreased by high acid and low sugar contents of infected fruiting clusters. When no control of the vector has been undertaken, Figure 3: Infected (FD-related the number of infected vines may increase phytoplasma) but non-symptomatic steadily about 10 times every year and may alders next to a vineyard in Germany. reach 80 to 100% within a few years. The Photo courtesy of S. Malembic-Maher, economic viability of maintaining a vineyard INRA Bordeaux. ceases when the productive plants are less than 25% of the total plants in a vineyard (CABI, 2013a).

211 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

Wine production in 2006 represented 5% of the value of total European Union (EU) agricultural output. France, Spain, Italy, and Portugal, all countries known to have FD, are home to roughly 92% of all vineyard acres in the EU (Pappalardo et al., 2012).

FD is viewed as a threat to the grape industry in the United States if it were to become established. One study estimates the total 2004 value of wine, grapes, and grape products to the U.S. economy to be $90 billion (MKF, 2006).

FD is listed as a harmful organism by the following 16 countries: Brazil, Chile, China, Costa Rica, Israel, Japan, Mexico, Morocco, New Caledonia, Japan, Russia, Thailand, Timor-Leste, Turkey, Ukraine, and Uruguay (PCIT, queried July 18, 2013). If the pest were found in the United States potential trade impacts with these countries could result.

Known Hosts Major host: Vitis spp. (EPPO, 2012).

All Vitis vinifera varieties are sensitive to FD but may not express symptoms of the disease to the same degree (Boudon-Padieu, 2002). Vitis rootstocks are also susceptible to infection.

Other possible hosts: Alnus glutinosa, Alnus incana, and Clematis vitalba (Arnaud et al. 2007, Filippin et al., 2009; Mehle et al., 2011).

Recent evidence points to these hosts as possible alternate hosts of FD phytoplasma strains (Filippin et al., 2009; Mehle et al., 2011).

Experimental hosts: Chrysanthemum carinatum, Trifolium repens, and Vicia faba (EPPO, 2012).

Known Vectors (or associated insects) Primary vector: Scaphoideus titanus (CABI, 2013a, b).

Other Potential Vectors: Dictyophara europaea, Orientus ishidae, and Oncopis alni (Filippin et al., 2009; Mehle et al., 2011). Dictyophara europaea is indicated by some authors to be a natural vector (Filippin et al., 2009). Others consider that all three of these potential vectors can harbor FD-related phytoplasmas but that they are not confirmed vectors of FD (Malembic-Maher/Foissac personal communication; Maixner personal communication).

Experimental vectors: Anoplotettix fuscovenosus, Euscelidius variegatus, Euscelis incisus (all leafhoppers from the family Cicadellidae) (Bressan et al., 2006).

212 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

Known Distribution Europe: Austria, Croatia, France (including Corsica), Hungary, Italy, Macedonia, Portugal, Serbia, Slovenia, Spain (including the Balearic Islands), and Switzerland (CABI, 2013a).

Recent studies have shown that 60 to 80% of alders (Alnus sp.) in Europe are infected with FD-related phytoplasma, and they are rarely symptomatic. Therefore, the range of this phytoplasma and related phytoplasmas in Europe is probably wider than currently reported (Foissac, personal communication).

A sample of Scaphoideus titanus collected in New York, United States was reported to carry a phytoplasma related to ‘Ca. P. vitis’ (Maixner et al., 1993), but ‘Ca. P. vitis’ has not been confirmed there. Surveys conducted in Virginia and in different provinces of Canada led to the detection of 16SrI and 16SrIII phytoplasmas in grapevine but no 16SrV types (Beanland et al., 2006; Olivier et al., 2009).

Pathway There have been 82 shipments of Vitis sp. propagative material from known host countries since 2003 (AQAS, queried July 12, 2013). Shipment sizes ranged from 1 gram to 1797 plant units. These shipments are likely comprised of a mixture of seed, plants, and cuttings based on the units of measure used. Vitis sp. (except seeds), however, are currently prohibited for import into the United States except from Canada (USDA, 2012). ‘Ca. P. vitis’, like other phytoplasmas, is not known to be transported through seed. Thus, this pathway appears to be unlikely. In contrast, Vitis sp. plant material destined for propagation has been intercepted from known host countries 33 times in the past ten years, indicating that people are bringing plant material into the United States from areas known to have FD (AQAS, 2013).

Dormant perennials of Clematis sp. are allowed entrance into the United States (USDA, 2012). There have been 57 shipments of Clematis sp. propagative material from known host countries since 2003. Shipment sizes ranged from 1 gram to 121,481 plant units. This appears to be a potential open pathway for FD-related phytoplasmas.

There have been shipments of Alnus sp. propagative material from FD containing countries (Hungary (17) and Italy (11since 2003). These were likely shipments of seed. Shipment sizes ranged from 2 kilograms to 139 kilograms with one indicating 25 plant units. There have been no reported interceptions of Alnus sp. propagative material since 2003 from known host countries (AQAS, queried July 17, 2013). Effective July 6, 2009, the entry of Alnus sp. plants for planting (except seed) was prohibited to prevent the introduction and dissemination of Phytophthora alni into the United States (USDA, 2012).

The potential vector D. europaea has been intercepted 7 times from Italy and Turkey on imported tile since January, 2003 (AQAS, 2013). Oncopis alni is currently listed in the AQAS database and no interceptions have been reported to date. These potential vectors are not known to be present in the United States. Orientus ishidae (Japanese

213 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

leaf hopper) is listed in AQAS as non-reportable, because it occurs in the United States (as of 1955 in New Jersey, New York, Maryland, Pennsylvania, New Hampshire, Washington D.C., Ohio, and Connecticut). This insect and potential vector could be allowed entry and contain FD-related phytoplasmas . There are also 26 native Oncopis spp. in the United States (McKamey personal communication). It is possible that one of these natives could vector FD or FD-related phytoplasmas if the pathogen were introduced.

Potential Distribution within the United States The potential for spreading of ‘Ca. P. vitis’ if established in the United States is considerable. There is a wide range of known and potential hosts (Fig. 4). In addition, the confirmed vector S. titanus is native to North America and present in at least 35 states and five Canadian Provinces (CABI, 2013a). While the potential vector Dictyophara europaea is not known to be present in North America, it has been intercepted in shipments of ceramic and marble tile from Italy, a country known to have ‘Ca. P. vitis’ (AQAS, 2013).

Survey CAPS-Approved Method*: The CAPS-approved survey method is to collect symptomatic plant tissue by visual survey.

Follow instructions in Phytoplasma sample submission for Cooperative Agricultural Pest Survey (CAPS) Program and Farm Bill Goal 1 surveys FY 2014.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Key Diagnostics CAPS-Approved Method: Molecular: Follow instructions in Phytoplasma sample submission for Cooperative Agricultural Pest Survey (CAPS) Program and Farm Bill Goal 1 surveys FY 2014.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/

Literature-Based Methods: Serological: According to EPPO (2007): ELISA with polyclonal and monoclonal antibodies has been used for detection of flavescence dorée in the vector and grapevine (Boudon-Padieu et al., 1989; Caudwell and Kuszala, 1992; Osler et al., 1992; Irimia et al., 2010). According to EPPO (2007), this method relies on “availability of antibodies which are not sold commercially and has been replaced in practice by PCR, which is versatile, specific and sensitive.” Irimia et al. (2010), however, state that the antibodies are available commercially. Neogen Europe and Sediag indicate that they have ELISA materials available commercially. This ELISA procedure has not been tested for sensitivity, specificity, or for regulatory use.

214 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

a)

b) c)

Alnus glutinosa Alnus incana

d) Present Absent Figure 4: Distribution of known and potential hosts of ‘Ca. P. vitis’, a) Vitis sp. (values from low to high indicate host acreage by county), b) Alnus glutinosa, c) Alnus incana, and d) Clematis vitalba distribution in the United States. (b-d courtesy of USDA, http://www.plants.usda.gov. courtesy USDA-APHIS-PPQ- Clematis vitalba a CPHST.

215 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

Molecular: Potentially infected samples can be analyzed for the presence of FD with a real-time PCR procedure as described in Hren et al. (2007), Angelini et al. (2007), or Pelletier et al. (2009). The Pelletier et al. (2009) detection method is the French official method validated by their Plant Protection Services. It is also now used in Croatia, Hungary, and Spain.

Further molecular characterization may then be performed on the positive samples by PCR followed by RFLP or sequencing of the secY gene or other non- ribosomal markers as described in Angelini et al. (2001, 2003) and Arnaud et al. (2007).

Easily Confused Pests Flavescence dorée resembles Australian grapevine yellows (AGY, ‘Candidatus Phytoplasma australiense’), bois noir, leaf curl, berry shrivel and other grapevine diseases (North American grapevine yellows, tomato big bud and an uncharacterized disease also believed to be caused by a phytoplasma) (Magarey and Wachtel, 1985; Davis et al., 1997; Lee et al., 1998; Constable et al., 1998; Gibb et al., 1999). There is no way to visually distinguish FD from bois noir (‘Candidatus Phytoplasma solani’) and other grapevine yellows diseases (Foissac, personal communication). Molecular methods are used to distinguish various grapevine yellows diseases.

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216 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

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Constable, F.E., Gibb, K.S., Moran, J.R., and Wilson, Y.M. 1998. Incidence of phytoplasma associated with yellows, restricted spring growth and late season leaf curl symptoms of grapevines. Australian Grape grower and Winemaker 409: 19-20.

Daire, X., Boudon-Padieu, E., Berville, A., Schneider, B., and Caudwell, A. 1992. Cloned DNA probes for the detection of grapevine flavescence dorée mycoplasma-like organism (MLO). Ann. Appl. Biol. 121: 95-103.

Davis, R.E., Dally, E.L., Gundersen, D.E., Lee, I.M., and Habili, N. 1997. ‘Candidatus Phytoplasma australiense,’ a new phytoplasma taxon associated with Australian grapevine yellows. International Journal of Systematic Bacteriology 47(2): 262-269.

Davis R., Zhao Y., Dally E.L., Lee I.-M., Jomantiene R., and Douglas S.M. 2013. ‘Candidatus Phytoplasma pruni’, a novel taxon associated with X-disease of stone fruits, Prunus spp.: multilocus characterization based on 16S rRNA, secY, and ribosomal protein genes. International Journal of Systematic and Evolutionary Microbiology 63:766-776.

217 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

Defra. Central Science Laboratory. 2000. EC listed disease: Grapevine flavescence dorée (Ref. no. QIC/54). Department for Environment, Food and Rural Affairs, Central Science Laboratory.

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Ember; I., Acs, Z., Salar, P., Danet, J.L., Foissac, X., Kolber, M., and Malembic-Maher, S. 2011. Survey and genetic diversity of phytoplasmas from the 16SrV-C and -D subgroups in Hungary. Bulletin of Insectology. 64: 33-34. International Phytoplasmologist Working Group, Neustadt, Sept. 2011.

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Filippin, L., Jović, J., Cvrković, T., Forte, V., Clair, D., Toševski, I., Boudon-Padieu, E., Borgo, M. and Angelini, E. 2009. Molecular characteristics of phytoplasmas associated with Flavescence dorée in clematis and grapevine and preliminary results on the role of Dictyophara europaea as a vector. Plant Pathology 58: 826-837.

Foissac, Xavier. 2013. INRA, Bordeaux, France. Personal communication.

Gibb, K.S., Constable, F.E., Moran, J.R., and Padovan, A.C. 1999. Phytoplasmas in Australian grapevines – detection, differentiation and associated diseases. Vitis 38(3): 107-114.

Gitau, C.W., Gurr, G.M., Dewhurst, C.F., Fletcher, M.J., and Mitchell, A. 2009. Insect pests and insect- vectored diseases of palms. Australian Journal of Entomology 48: 328-342.

Harrison, N.A., Gundersen-Rindal, D., and Davis, R.E. 2011. Genus I. “Candidatus Phytoplasma” gen. nov. IRPCM.

Hren, M., Boben, J., Rotter, A., Kralj, P., Gruden, K., and Ravnikar, M. 2007. Real-time PCR detection systems for ‘flavescence doree’ and ‘bois noir’ phytoplasmas in grapevine: comparison with conventional PCR detection and application in diagnostics. Plant Pathology 56: 785–796.

INRA. 2013. Grapevine Flavescence Doree. Retrieved from, http://www.inra.fr/en/Scientists-Students/Plant-biology/All-reports/Grapevine-flavescence-doree.

Irimia, N., Ulea, E., Balau, A.M., 2010. Detection of pathogen Flavescence dorée phytoplasma in some grapevine varieties using ELISA test. Lucrări Ştiinţifice 53: 187-190.

IRPCM Phytoplasma/Spiroplasma Working Team – Phytoplasma Taxonomy Group (IRPCM). 2004. ‘Candidatus Phytoplasma’, a taxon for wall-less, non-helical prokaryotes that colonize plant phloem and insects. International Journal of Systemic and Evolutionary Microbiology 54: 1243-1255.

Lee, I.M., Gundersen-Rindal, D.E., Davis, R.E., and Bartoszyk, I.M. 1998. Revised classification scheme of phytoplasmas based on RFLP analyses of 16S rRNA and ribosomal protein gene sequences. International Journal of Systematic Bacteriology 48(4): 1153-1169.

Lee, I.M., Davis, R.E., Gundersen-Rindal, D.E. 2000. Phytoplasma: Phytopathogenic Mollicutes. Ann. Rev. Microbiol. 54: 221-255.

Magarey, P.A., and Wachtel, M.F. 1985. Grapevine yellows, a widespread, apparently new disease in Australia. Plant Disease 70: 694.

218 Candidatus Phytoplasma vitis Primary Pest of Grape Phytoplasma Flavescence dorée

Marzorati, M., Alma, A., Sacchi, L., Pajoro, M., Palermo, S., Brusetti, L., Raddadi, N., Balloi, A., Tedeschi, R., Clementi, E., Corona, S., Quaglino, F., Attilio Blanco, P., Beninati, T., Bandi, C., and Daffonchio, D. 2006. A Novel Bacteroidetes Symbiont Is Localized in Scaphoideus titanus, the Insect Vector of Flavescence Doree in Vitis vinifera. Applied And Environmental Microbiology 72(2): 1467– 1475.

Maixner, M., Reinert, W., and Darimont, H. 2000. Transmission of grapevine yellows by Oncopsis alni (Schrank) (Auchenorryncha: Macropsinae).- Vitis 39(2): 83-84.

Maixner, Michael. 2013. Julius Kühn Institute, Siebeldingen, Germany. Personal communication.

Malembic-Maher, Sylvie, and Foissac, Xavier. 2013. INRA, Bordeaux, France. Personal communication.

Malembic-Maher, S., Salar, P., Vergnes, D., and Foissac, X. 2007. Detection and diversity of “flavescence dorée” related phytoplasmas in alders surrounding infected vineyards in Aquitaine - France. In: First International Phytoplasmologist Working Group meeting, pp. 329-330. Bologna: Bulletin of Insectology.

Malembic-Maher, S., Salar, P., Filippin, L., Carle, P., Angelini, E., and Foissac, X. 2011. Genetic diversity of European phytoplasmas of the 16SrV taxonomic group and proposal of 'Candidatus Phytoplasma rubi'. International Journal of Systematic and Evolutionary Microbiology. 61: 2129-2134.

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Mehle, N., Seljak, G., Rupar, M., Ravnikar, M., and Dermastia, M. 2010. The first detection of a phytoplasma from the 16SrV (elm yellows) group in the mosaic leafhopper Orientus ishidae. New Disease Reports 22: 11.

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Olivier, C.Y., Lowery, D.T., Stobbs, L.W., Vicent, C., Galka, B., Saguez, J., Bittner, L., Johnson, R., Rott, M., Masters, M., and Green, M. 2009. First report of aster yellow phytoplasmas ('Candidatus Phytoplasma asteris') in Canadian grapevines. Plant Disease 93(6) : 669.

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Datasheet Reviewers: Dr. Robert Davis, USDA Agricultural Research Service, Beltsville, MD, Sylvie Malembic-Maher and Xavier Foissac, INRA, Bordeaux, France, and Michael Maixner, Julius Kühn Institute, Siebeldingen, Germany.

220 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Phellinus noxius

Scientific Name Phellinus noxius (Corner) G. Cunn. 1965

Synonyms: Corticum spp., Fomes noxius, Hymenochaete noxia, Hymenochaete noxius, Phellinidium noxium, Poria setulalsocrocea

Common Name Brown root rot, brown cocoa root rot, brown root, brown tea root disease, collar rot, stem rot

Type of Pest Fungus

Taxonomic Position Class: Basidiomycetes, Order: Hymenochaetales, Family: Hymenochaetaceae

Reason for Inclusion in Manual CAPS Target: AHP Prioritized Pest List – 2006 through 2009.

Pest Description The basidiocarp, also referred to as a sporocarp or conk, is perennial, solitary or imbricate, sessile with a broad basal attachment, commonly resupinate. Basidiocarps are not always produced in nature but can be induced in the laboratory (Bolland et al., 1984). Pileus 5 to 13 x 6 to 25 x 2 to 4 cm (1.97 to 5.12 x 2.36 to 9.84 x 0.79 to 1.57 in.), applanate, dimidiate or appressed-reflexed; upper surface deep reddish-brown to umbrinous, soon blackening, at first tomentose, glabrescent, sometimes with narrow concentric zonation, developing a thick crust; margin white then concolorous, obtuse. Context up to 1 cm (0.39 in.) thick, golden brown, blackening with KOH, silky-zonate fibrous, woody. Pore surface grayish-brown to umbrinous; pores irregular, polygonal, 6 to 8 mm (0.24 to 0.31 in.), 75 to 175 µm diameter, dissepiments 25 to 100 µm thick, brittle and lacerate; tubes stratified, developing two to five layers, 1 to 4 mm (0.04 to 0.16 in.) to each layer, darker than context, carbonaceous.

Basidiospores approximately 4 x 3 µm, ovoid to broadly ellipsoid, hyaline, with a smooth, slightly thickened wall, and irregular guttulate contents. Basidia 12 to 16 x 4 to 5 µm, short clavate, four-spored. Setae absent. Setal hyphae present both in the context and the dissepiment trama. Context setal hyphae radially arranged, up to 600 x 4 to 13 µm, unbranched or rarely branching, with a thick dark chestnut brown wall and capillary lumen; apex acute to obtuse, occasionally nodulose. Tramal setal hyphae diverging to project into the tube cavity, 55 to 100 x 9 to 18 µm, with a thick dark chestnut-brown wall (2.5 to 7.5 µm thick) and a broad obtuse apex. Hyphal system dimitic with generative and skeletal hyphae, non-agglutinated in the context, but strongly

221 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

agglutinated in the dissepiments. Generative hyphae 1 to 6.5 µm diameter, hyaline or brownish, wall thin to somewhat thickening, freely branching, simple septate. Skeletal hyphae 5 to 9 µm diameter, unbranched, of unlimited growth, with a thick reddish-brown wall (up to 2.5 µm thick) and continuous lumen, non-septate (Pagler and Waterson, 1968).

Biology and Ecology Phellinus noxius is a facultative parasite. It obtains nutrients from living or dead plants. As a tropical plant pathogen, its mycelium grows best at 25 to 30°C (77 to 86°F); it does not grow at temperatures below 4°C (39°F) or above 40°C (104°F). Colonies grown in the laboratory have distinctive raised brown and white plaques (patches) (Fig. 1) and occasionally produce arthrospores, which are asexual spores formed by Figure 1. Mycelia (left) and arthrospores (right) of P. noxius. the division of Photos courtesy of Pao-Jen Ann. special hyphae into one-celled segments. Ma ny fungi produce asexual spores, which allows dissemination of the fungus within the same plant or from plant to plant. Arthrospores of P. noxius have not been observed in nature, however, and their importance remains unknown (Brooks, 2002).

The name brown root rot refers to a brown to black mycelial crust formed by the fungus on the surface of infected roots and stem bases (Chang and Yang, 1998). However, P. noxius is generally considered a white rust fungus because of its ability to degrade lignin, a complex molecule that gives wood much of its strength and brown color (Fig. 2) (Chang and Yang, 1998). The mycelium of P. noxius also forms microhyphae and hyphae with extracellular sheaths. These structures are found where wood is being Figure 2. Dry, honeycombed, white wood rot caused by P. noxius. Photo courtesy of Fred Brooks.

222 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner destroyed but their exact function is not yet known (Brooks, 2002).

P. noxius is spread in two main ways. The first is by windborne spores (basidiospores), which can infect freshly cut tree stumps or fresh wounds. The second and most important is by root-to-root contact. The leading edge of the mycelial sleeve will infect healthy roots of other trees if they touch. Infected root pieces can remain viable for many years in the soil (Bolland, 1984; CABI, 2007). Basidiospores, the sexual spores, of P. noxius contain one nucleus (monokaryotic) and are haploid. When haploid hyphae (n) of different mating types touch each other, they can fuse to form a dikaryotic mycelium with cells containing two haploid nuclei (n + n). This dikaryotic mycelium is the most common condition in nature. The hyphae can be modified (swelling, thickening, or gluing together) to produce a mycelial crust or basidiocarps, which often start as small round patches on stems of dead trees. The patches may continue to grow flat against the wood, grow out into a shelf-like conk, or a combination of both. The upper surface of the basidiocarp is sterile, the lower fertile surface covered with small tubes or pores, These tubes are lined with basidia, whose two nuclei fuse and undergo meiosis, producing four haploid basidiospores. During wet weather, basidiospores are released and spread by the wind. Basidiospores may be responsible for some long distance dispersal of the fungus (Brooks, 2002).

P. noxius is present in native tropical forests and plantations on infected roots, stumps, and woody debris. It does not form survival structures such as sclerotia or resting spores but may persist in dead roots and colonized wood for many years (Chang, 1996). Centers of disease radiate outward as roots of healthy trees come in contact with roots of diseased trees. Trees of all ages are susceptible. A mycelial crust forms around infected roots, secreting wood-rotting enzymes as it moves up the roots to the stem. Crusts 0.5 to 1.0 cm (0.2 to 0.39 in.) thick usually extend 1 to 2 m (3 to 6 feet) up the stem, though crusts almost 5 m (15 ft) high have been measured. If the tree does not die from severe root rot, it is killed when the crust surrounds the stem and the underlying mycelium destroys the sapwood. Figure 3. Mycelial crust of P. noxius on multi-trunked tree in the rainforest. Symptoms/Signs Photo courtesy of Fred Brooks. Symptoms of brown root rot are similar www.bugwood.org. to those caused by other root rot

223 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

pathogens: slow plant growth, yellowing and wilting of leaves, defoliation, branch dieback, and plant death (Brooks, 2002). Although dead wood is initially discolored reddish brown, it later becomes white, dry, and crumbly.

Signs of the pathogen, unlike the symptoms, are distinctive for this disease. P. noxius forms a thick, dark brown to black crust of mycelium around infected roots and lower stems (Fig. 3), which gives the disease its name. The leading edge of the crust is creamy white, glistens with drops of clear, brownish exudate, and is usually noticeable even in the dark understory of the rainforest. Patches of white mycelium are present between the bark and sapwood. As colonization progresses, white, soft, crumbly wood becomes laced with reddish Figure 4. With age, reddish hyphae in wood strands of fungus hyphae that turn agglutinate (stick together), turn black and black with age (Fig. 4). brittle. Photo courtesy of Fred Brooks. Basidiocarps, or fruiting bodies, are purplish brown bracts (conks) with yellow-white growing margins and concentric blackish zones towards the edges (CABI, 2007). The basidocarps are gray to brown on the spore-forming surface (Brooks, 2002). Unlike other similar fungi, there are no rhizomorphs. Spread is by physical contact with the root encrustations.

Pest Importance P. noxius is known to cause extensive damage in a wide range of species, including common species used in industrial forest plantations, commercial fruit orchards, commodity crops such as cacao and rubber, and non-commercial components of native forests (Hodges, 2005). The fungus would be expected to cause similar losses if introduced into the tropical and subtropical regions of North America.

Due to the extremely diverse host range and geographical distribution, the economic impact of P. noxius is highly variable (CABI, 2007). The impact can vary from insignificant to losses of 60% in rubber tree plantations after 21 years (Nandris et al., 1987). P. noxius has caused 10% mortality in plantations of Swietenia macrophylla in Fiji and 50% mortality in a three-year-old plantation of Pinus merkusii in Indonesia (Bolland, 1984).

Brown root rot has become one of the most serious problems of fruit and forest trees in central and southern Taiwan at elevations less than 800 m (Chang and Yang, 1998).

224 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Known Hosts Chang and Wang (1998) showed that P. noxius has a broad host range that includes woody hardwoods, conifers, and annual herbaceous plants. It seems, however, to be more saprotrophic than parasitic on annual herbaceous plants, because it does not cause severe wilting of host foliage. More than 200 woody plant species representing 59 families have so far been recorded as host plants for P. noxius. Hosts were not classified as major or minor hosts, because most hosts were reported simply as hosts with no attempt to classify economic loss or susceptibility to P. noxius.

Acacia spp. (wattles), Actinidaphne pedicellata (litsea), Adenanthera pavonina (coralwood), Agathis palmerstonii (kauri gum), Albizia spp. (albizia), Aleurites spp. (Indian walnut), Alstonia scholaris (blackboard tree), Anacardium occidentale (cashew), Annona spp. (sour sop, custard apple), Aralia elata (Japanese aralia), Araucaria cunninghamii (colonial, hoop pine), Araucaria spp., Ardisia spp., Areca spp., Areca triandra (wild areca palm), Argusia argentea, Artemisia capillaris (wormwood), Artemisia spp., Artocarpus spp. (breadfruit), Averrhoa carambola (carambola), Barleria cristata (crested Philippine violet), Barringtonia spp., Bauhinia spp. (bauhinia), Bischofia javanica (autumn maple tree), Blepharocarya involucrigera (bolly gum), Boehmeria nivea (ramie), Bombax ceiba (silk cotton), Bougainvillea spp. (bougainvillea), Breynia nivosa (snowflower), Broussonetia kazinoki (small paper mulberry), Broussonetia papyrifera (paper mulberry), Buxus bodinieri, Cajanus cajan (pigeon pea), Callicarpa japonica var. luxurians, Calocedrus formosana (Taiwan incense cedar), Calophyllum inophyllum (Indian beauty leaf), Calophyllum spp., Camellia japonica (camellia), Camellia sinensis (tea), Canangia odorata (ilang-ilan), Canarium spp., (galip, galipnut), Carica papaya (papaya), Cassia spp., Castenospermum australe, Castanospora alphandii, Casuarina spp., Cedrella mexicana, Ceiba pentandra, Celtis sinensis, Cerbera manghas (Madagascar ordeal bean), Chamaecyparis formosensis (Taiwan red cypress), Chlorophora excelsa (African Teak), Chorisia speciosa (floss silk tree), Chrysalidocarpus lutescens (yellow areca palm), Cinnamomum camphora (camphor), Cinnamomum spp., Cinnamomum zeylanicum (Ceylon cinnamon), Citrus spp., Clutia spp. (clutia), Cocos nucifera (coconut), Codiaeum variegatum (croton), Coffea spp. (coffee), Cola nitida (bitter cola), Cordia spp. (cordia), Crataegus spp. (hawthorne), Crescentia cujete (calabash), Crossostylis biflora, Crotalaria spp. Cryptocarya spp. (crytocarya), Cupressus lusitanica, Cycas taiwaniana (Taiwan cycas), Dalbergia sissoo (sissoo tree), Delonix regia (flame tree), Dimocarpus longan (longan), Diospyros spp., Distylium racemosum, Duranta repens (creeping sky flower), Dysoxylum samoense (maota), Ehretia dichotoma, Elaeis guinennsis (African oil palm), Elaeis spp., Elaeocarpus serratus (Ceylon olive), Elaecarpus sylvestris var. ellipticus, Elattostachys falcata, Eribotrya japonica (loquat), Erythrina variegata (Indian coral tree), Eucalyptus spp., Eugenia uniflora, Euonymus japonicus (Japanese euonymus), Euphoria longana (longan), Ficus spp., Firmiana simplex (Chinese parasol-tree), Flemingia macrophylla, Flindersia brayleyana (Queensland Maple), Flueggea flexuosa (bushweed, poumuli), Fraxinus formosana (island ash), Garcinia mangostana (mangosteen), Garcinia spp., Gardenia spp.(cape jasmine), Glochidion spp., Gmelina arborea (white teak), Grevillea robusta (silver oak), Hedera australiana, Heritiera spp., Hernadia nymphaeifolia, Hevea brasilensis (rubber tree), Hevea spp., Hibiscus spp. (hibiscus), Hydrangea chinensis

225 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

(Chinese hydrangea), Ilex rotunda (kurogane-mochi), Inocarpus fagifer (Tahitian chestnut), Intsia bijuga (Borneo-teak), Ipomoea pescaprae (beach morning glory), Jatropha integerrima (peregrina), Jatropha pandurfolia, Keteleeria davidiana var. formosana (Taiwan keteleeria), Khaya ivorensis (African Mahogony), Kigelia pinnata (sausage tree), Kissodendron australianum, Kleinhovia hospita (guest tree), Koelreuteria spp., Lactuca indica (wild lettuce), Lagerstroemia spp. (crape myrtle), Lantana camara (lantana), Leucaena leudocephala (white popinac), Leucaena spp., Ligustrum japonicum, Liquidambar formosana (sweetgum), Litchi chinensis (litchi), Litsea spp., Macaranga spp. (lau pata), Machilus spp. (tamagusu), Maesa tenera, Mallotus paniculatus (turn-in-the-wind), Mangifera indica (mango), Manihot utilissima (cassava), Maytenus diversifolia, Melaleuca leucadendron (cajuput tree), Melia azedarach (China berry), Melicope merrilli (melicope), Melodinus augustifolius (narrow leafed melodinus), Michelia spp. (michelia), Morinda citrifolia (Indian mulberry), Morus australis, Muntingia calabura (Indian cherry), Murraya paniculata (orange jasmine), Myristica fatua (nandai), Nandina domestica (sacred bamboo), Neolitsea parvigemma (small bud Neolitsea), Neonauclea forsteri, Nephelium lappaceum (rambutan), Nerium oleander (oleander), Ochroma spp. (balsa), Osmanthus fragrans (sweet osmanthus), Pachira macrocarpa (Malabar chestnut), Palaquium formosanum (Taiwan guayule, Formosan nato tree), Persea americana (avocado), Persea spp., Pinus spp. (pine), Piper nigrum (black pepper), Pipturus argenteus (native mulberry), Pistacia chinesensis (Chinese pistachio), Pittosporum tobira, Planchonella spp., Podocarpus macrophyllus (yew), Pometia pinnata (Fijian longan, taun), Pongamia pinnata (pongamia), Populus spp. (poplar), Prunus spp. (Japanese plum, peach), Pterocarpus indicus (rose wood), Pyrus spp. (pear), Rhaphiolepis indica var. umbellata, Rhodendron obtusum (rhododendron), Rhus spp.(Japanese waxtree), Rosa spp. (rose), Roystonea regia (royal palm), Salix babylonica (willow), Samanea saman (crow bean tree), Sauramja oldhami, Schefflera spp. (scheffera), Spondias dulcis (Jewish plum),Stenocarpus sinuatus (fire wheel tree), Sterculia spp. (sterculia), Swietenia spp. (mahogany), Syzygium spp. (wax apple), Tabebuia chrysantha (yellow golden bell tree), Taiwania cryptomerioides (Taiwania), Tectonia grandis (teak), Tephrosia spp., Terminalia spp. (tropical almond), Thea spp., Theobroma cacao (cocoa), Thespesia populnea (Pacific rosewood, portia tree), Thevetia peruviana (luckynut), Trema orientalis (charcoal tree), Ulmus parviolia (Chinese elm), Urena lobata (bur mallow, cadillo), Vitis spp. (grape), Vitis vinifera (grape), and Zelkova serrata var. serrat (Bolland, 1984; Neil, 1986; Almonicar, 1992; Ann et al., 2002; Supriadi et al., 2004; CABI, 2007; Sahashi et al., 2007; Sashashi et al., 2012).

Known Vectors (or associated insects) P. noxius does not have a known vector or associated organisms.

Known Distribution The fungus is confined mainly to tropical areas of the world.

Asia: India, Indonesia, Japan, Malaysia, Myanmar, Pakistan, Philippines, Singapore, Sri Lanka, Taiwan, and Vietnam. Africa: Angola, Benin, Burkina Faso, Cameroon, Central African Republic, Congo, Cote d`Ivoire (Ivory Coast), Gabon, Ghana, Kenya, Nigeria,

226 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Sierra Leone, Tanzania, Togo, and Uganda. Central America: Costa Rica, Cuba, and Puerto Rico. South America: Brazil, French Guiana. Oceania: American Samoa, Australia, Federated states of Micronesia, Fiji, Guam, New Caledonia, Niue, Northern Mariana Islands, Papua New Guinea, New Zealand, Samoa, Solomon Islands, and Vanuatu.

Potential Distribution within the United States P. noxius has a broad host range and would undoubtedly find numerous suitable hosts in North America, but would most likely be restricted to tropical or near tropical regions (Hodges, 2005).

A recent risk analysis by USDA-APHIS-PPQ-CPHST shows that portions of the Great Lake states, the northeastern United States, and Colorado, Florida, Idaho, and Utah have the greatest risk of P. noxius establishment based on host availability. Most of the remaining states have counties that are considered at low to moderate risk.

Survey CAPS-Approved Method*: Collect symptomatic plant material via visual survey.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Visual survey: Visual survey is the most common method used to survey for P. noxius. A dark brown mycelial mat or sleeve on the surface of the roots and up to the base of the stem is used reliably for field identification of P. noxius. Soil is scraped away around the collar and the main roots and the distinctive mycelial sleeve is often present (Nandris et al., 1987). Particular attention should be paid to trees that appear wilted or dead. P. noxius tends to be a problem in cleared forests converted to agricultural land (tree farms) or in disturbed areas and surveys should be conducted in these areas.

Baiting: Early detection of the pathogen before typical wilt symptoms are visible is very difficult and time consuming. Baiting out the pathogen by placing sticks of a susceptible host in the soil and retrieving for laboratory examination after three weeks is also conducted, particularly in virgin forests to detect parasites on the root system of wild trees (Nandris et al., 1987; CABI, 2007). According to Nandris et al. (1987), the area around the root collar can be mulched for three weeks to provide a damp zone that allows the superficial mycelium to progress from the roots onto the trunk of rubber trees. When the mulch is removed, the mycelial filaments of the pathogen can be observed.

Key Diagnostics/Identification CAPS-Approved Method*: Confirmation of P. noxius is via morphological identification.

A. Surface sterilized diseased root tissues are plated on potato dextrose agar amended with ampicillin and benomyl or Chang (1995) medium.

227 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

B. The cultural characteristics of the fungus are examined and compared to photos in Ann et al. (2002) and Brooks et al. (2002).

C. The Key of the Polyporaceae described by Cunningham (1965) is then used for identification of the fungus.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: According to Ann et al (1999), after plating surface sterilized diseased root tissue on potato dextrose agar amended with ampicillin and benomyl, the cultural and morphological characteristics are the fungus are examined and compared. The Key of the Polyporaceae described by Cunningham (1965) is then used for identification of the fungus. In culture, mycelia are initially white and then brown with irregular dark brown lines or patches. In addition, staghorn-like hyphae and arthrospores, but no clamp connections are commonly observed (Sahasi et al., 2007).

Chang (1995) developed a selective medium for P. noxius using malt extract agar as a basal medium amended with benomyl, dicloran, ampicillin, and gallic acid. Tergitol NP-7 was added for isolation from soil.

Bolland et al. (1984) developed a method to induce sporulation in basidiocarps of P. noxius to obtain single spore isolates.

Easily Confused Pests P. noxius basidiocarps are sometimes confused with P. lamaensis, another tropical Phellinus species. P. lamaensis sporocarps have short, reddish-brown, cone-shaped cells called hymenial setae growing into their pores, however, P. noxius does not.

References Almonicar, R.S. 1992. Two types of root rot diseases affecting Acacia mangium. Nitrogen Fixing Tree Research Reports 10: 94-95.

Ann, P.J., Lee, H.L., and Huang, T.C. 1999. Brown root rot of 10 species of fruit trees caused by Phellinus noxius in Taiwan. Plant Disease 83: 746-750.

Ann, P.J., Chang, T.T., and Ko, W.H. 2002. Phellinus noxius brown root rot of fruit and ornamental trees in Taiwan. Plant Disease 86(8): 820-826.

Bolland, L. 1984. Phellinus noxius: cause of a significant root-rot in Queensland hoop pine plantations. Australian Forestry 47(1): 2-10.

Bolland, L., Griffin, D.M., and Heather, W.A. 1984. Induction of sporulation in basidiomes of Phellinus noxius and preparation of single spore isolates. Bulletin of the British Mycological Society 18(2): 131-133.

Brooks, F.E. 2002. Brown root rot. The Plant Health Instructor. DOI: 10.1094/PHI-I-2002-0923-01.

Chang, T.-T. 1995. A selective medium for Phellinus noxius. European Journal of Forest Pathology 25: 185-190.

228 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Chang, T.T. 1996. Survival of Phellinus noxius in soil and in the roots of dead host plants. Phytopathology 86: 272-276.

Chang, T.T., and Yang, W.W. 1998. Phellinus noxius in Taiwan: distribution, host plants and the pH and texture of rhizosphere soils of infected hosts. Mycological Research 102(9): 1085-1088.

Cunningham, G.H. 1965. Phellinus. In: Polyporaceae in New Zealand. N.Z. Dep. Sc. Ind. Res. Bull. 164: 217-240.

Hodges, C.S. 2005. Phellinus noxius. Pest Reports – EXFOR Database. http://spfnic.fs.fed.us/exfor/data/pestreports.cfm?pestidval=50&langdisplay=english.

Nandris, D., Nicole, M., and Geiger, J.P. 1987. Root rot disease of rubber trees. Plant Disease 71(4): 298-306.

Neil, P.E. 1986. A preliminary note on Phellinus noxius root-rot of Cordia alliodora plantings in Vanuatu. European Journal of Forest Pathology 16(5/6): 274-280.

Pegler, D.N., and Waterson, J.M. 1968. Phellinus noxius, C.M.I. descriptions of pathogenic fungi and bacteria. Commonwealth Mycological Institute 195. 2 pgs.

Sahasi, N., Akiba, M., Ishihara, M., Abe, Y., and Morita, S. 2007. First report of the brown root rot disease caused by Phellinus noxius, its distribution and newly recorded host plants in Amami Islands, southern Japan. Forest Pathology 37: 167-173.

Sahasi, N., Akiba, M., Ishihara, M., Ota, Y., and Kanzaki, N. 2012. Brown root rot of trees caused by Phellinus noxius in the Ryjkyu Islands, subtropical areas of Japan. Forest Pathology. Doi: 10.1111/j.1439- 0329.2012.00767.

Supriadi, E., Adhi, M., Wahyuno, D., Rahayuningsih, S., Karyani, N., and Dahsyat, M. 2004. Brown root rot disease of cashew in West Nusa Tenggara: distribution and its causal organism. Indonesian Journal of Agricultural Science 5: 32-36.

229 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Pseudopezicula tracheiphila

Scientific Name Pseudopezicula tracheiphila, Müller – Thurgau 1913 (Korf et. al, 1986)

Synonyms: Pseudopeziza tracheiphila, Phialophora tracheiphila (anamorph)

Common Name(s) Rotbrenner, red fire

Type of Pest Fungal pathogen

Taxonomic Position Phylum: Ascomycota, Class: Ascomycetes, Order: Helotiales, Family: Helotiaceae

Reason for Inclusion in Manual CAPS target: AHP Prioritized Pest List - 2014

Background Pseudopezicula tracheiphila is an Ascomycete fungus. The primary morphological character that distinguishes members of the Ascomycota is the ascus (plural asci), a sac-like cell containing the ascospores cleaved from within by free cell formation after karyogamy and meiosis. Eight ascospores typically are formed within the ascus but this number may vary from one to over a thousand according to the species. Asci are typically formed in an ascocarp (i.e., a perithecium, pseudothecium, Figure 1. Eight-spored asci and branch paraphyses apothecium, or of Pseudopezicula tracheiphila. Photo courtesy W. cleistothecium). Gärtel; Reprinted, by permission, from Pearson, Ascomycetes may have two R.C., and Goheen, A.C. 1988. Compendium of distinct reproductive phases, Grape Diseases. American Phytopathological one sexual (teleomorph) Society, St. Paul, MN.

Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

involving the formation of the asci and ascospores, and the other asexual (anamorph), with spore/conidia production occurring at different times on the same mycelium.

Pest Description Teleomorph (sexual stage): Pseudopezicula tracheiphila Anamorph (asexual stage): Phialophora tracheiphila

Pseudopezicula tracheiphila: The fungus produces apothecia (an open, cuplike or saucerlike, ascus-bearing fungal fruiting body (ascocarp), often supported on a stalk) as the perfect/sexual stage. Apothecia are erumpent from the leaf tissue and are often associated with the veins. They are minute, up to 0.6 mm in diameter, sessile, gelatinous, whitish to faintly colored and gregarious. Apothecia contain inoperculate, broadly clavate, eight- spored asci (115 to 145 x 18 to 28 μm) (Fig. 1.), Figure 2. Hyphae of Phialophora which show a blue pore in iodine after treatment tracheiphila growing in a sine-wave with aqueous potassium hydroxide. Ascospores pattern inside a vessel element of a are hyaline, ellipsoid (19 to 27 x 9 to 14 μm) and leaf. Photo courtesy of H. Schüepp; are flattened on one side. During germination, Reprinted, by permission, from ascospores produce a spore-like vesicle on one Pearson, R.C., and Goheen, A.C. side, which is characteristic of the fungus. 1988. Compendium of Grape Paraphyses are branched and curved or slightly Diseases. American deformed at the apex, filiform, septate and Phytopathological Society, St. Paul, hyaline (Plant Health Australia, 2004). MN.

The formal/technical description is available in Korf et al. (1986).

Phialophora tracheiphila: The fungus produces short, hyaline and septate conidiophores that carry unicellular, ellipsoid, aseptate, and hyaline conidia (2 to 3 x 1.5 to 2 μm) (Konig et al., 2009). Generally the conidiophores are coarser than the vegetative hyphae (Plant Health Australia, 2004).

Cultural characteristics: The diagnostic feature of the fungus is the presence of sine-wave pattern (Fig. 2) of hyphae observed in the xylem vessel elements when affected plant tissues are cleared by boiling in 2% aqueous potassium hydroxide for 2 to 3 minutes (Plant Health Australia, 2004).

231 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Biology and Ecology P. tracheiphila is an erumpent discomycete forming tiny, densely aggregated apothecia, 95 to 125 apothecia per cm², mostly on the lower surface of leaves. On the upper surface, less than 1 to 20 apothecia per cm² are typically observed (Korf et. al, 1986). The fungus overwinters in infected vine leaves on the ground. In Germany, P. tracheiphila usually occurs after frequent rains in April and May and affects the shoots and leaves. Conditions required for the fungus to invade the vascular system are not well understood; however, soil conditions and water supply that place the vine under temporary stress appear to be important factors (Konig et. al, 2009).

The source of inoculum of the disease is ascospores, which are formed sexually in asci within apothecia. P. tracheiphila is composed of two different mating types and exhibits a bipolar heterothallic mating system. Apothecia are formed primarily on fallen leaves in the spring. Apothecia may also develop on current-season infected leaves in late summer or fall. Apothecia development requires sufficient wetness on fallen leaves. Depending on weather conditions, apothecia with mature ascospores may be present throughout the season. Under wet and warm conditions, ascospores are released before bud burst. Disease incidence and severity depend on the abundance of apothecia on fallen leaves on the ground of the vineyard and on the number of released ascospores and leaf wetness (Konig et. al, 2009). Heavy rainfall and prolonged leaf wetness favor infection and lead to severe Figure 3. Rotbrenner on grape leaves. Photos disease. The fungus has an courtesy of Dr. Michael Maxiner, Julius Kühn incubation period of two-to-four Institute (JKI), Federal Research Centre for weeks from infection to symptom Cultivated Plants Institute for Plant Protection in development (Konig et al., 2009). Fruit Crops and Viticulture

The fungus is primarily spread by water and airborne spores (Plant Health Australia, 2009). Unlike other members of Peziculoideae that attack woody stems

232 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner and branches, these species occur on leaves, mostly in association with the vascular elements.

Symptoms/Signs Lesions on leaves (Fig. 3, 4) are initially yellow on white cultivars and bright red to reddish brown on red cultivars. A reddish brown necrosis develops in the center of the lesion (Fig. 4), leaving only a thin margin of yellow or red tissue between the necrotic and green areas of the leaf. The lesions are typically confined to the major veins and the edge of the leaf and are several cm wide. Early infections occur on the first to the sixth leaf position of young shoots, resulting in minor losses. Later infections attack leaves up to the 10th or 12th position on the shoot, Figure 4. Closeup of rotbrenner on grape which results in severe defoliation. The leaves. Photo courtesy of Dr. Michael fungus also attacks the inflorescences Maxiner, Julius Kühn Institute (JKI), Federal and berries causing them to rot, dry out Research Centre for Cultivated Plants and fall. Institute for Plant Protection in Fruit Crops and Viticulture Young leaves are susceptible after they reach a width of about 5 cm but the probability of infections increases from the 6- leaf stage. After an incubation period of two to four weeks, the fungus invades the vascular elements of infected leaves, causing symptom development (Konig et. al, 2009).

Pest Importance Rotbrenner, caused by Pseudopezicula tracheiphila, is an important grape disease in the cool viticulture regions of Europe (e.g., Austria, France, Germany, Hungary, and Switzerland) (Pearson et al., 1991). Rotbrenner causes economic yield loss in all common European cultivars of Vitis vinifera (grape), especially on cultivars Elbling, Müller-Thurgau, and the red cv. Domina, where it causes strong early leaf fall and depression of the quality of wine by a lower sugar content of the must.

Attack of flower clusters results in a depression of yield from approximately 25 to 100% (Holz, 2000). Early infections generally result in less loss than late infections. Inflorescences can be attacked before or during flowering, causing them to rot and dry out. High levels of infection during flowering can lead to severe yield losses of up to 90%.

233 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Pseudopezicula tracheiphila is on the harmful organism list for Brazil, Chile, and China. This pest could have trade implications with these countries if the pest was found in the United States.

Known Hosts Major Hosts: Parthenocissus quinquefolia (Virginia creeper), Parthenocissus tricuspidata (Boston ivy), Vitis labrusca (fox grape),Vitis riparia (riverbank grape), and Vitis vinifera (grapevine) (Korf et al., 1986).

Known Vectors (or associated insects) Pseudopezicula tracheiphila is not vectored by any insect and is not known to be a vector.

Known Distribution Asia: Jordan, Turkey. Europe: Austria, France, Germany, Hungary, Switzerland, Romania, Montenegro (former Yugoslavia), Serbia, Moldova, Ukraine, and Russia. Africa: Tunisia. South America: Brazil (Korf et al., 1986; USDA ARS, 2013).

The North American report from New York is now known to be a related species Pseudopezicula tetraspora (Korf et al., 1986; Pearson et al., 1986; Pearson et al., 1988). The South America isolate may also be P. tetraspora, a related North American species (USDA ARS, 2013).

Potential Distribution within the United States Rotbrenner is most likely to occur in the cool viticulture regions of the United States. Based on interception and commodity destination data, the most likely places for the introduction of this pest would be in California, New York, New Jersey, Illinois, Kansas, and Georgia.

Pathway Vitis sp. are regulated via 7CFR319.37–5(b)(1) for Pseudopezicula tracheiphila. Vitis spp. (all propagules except seeds) are prohibited from all countries except for Canada when meeting the required import conditions (Plants for Planting Manual, 2012). This fungal pathogen is not known to be seedborne in grape.

Parthenocissus propagative material, however, doesn’t appear to be subject to regulations and there appears to be an open pathway for Parthenocissus PM. Although shipments appear to be mostly seed, there were 16 shipments of Parthenocissus sp. PM since 2001 from countries where this pest is known to occur. There were an additional 46 shipments of Vitis sp. PM from countries where this pest is known to occur. These shipments also appear to be mostly seed.

234 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

There have been one hundred interceptions of host material that could harbor P. tracheiphila from countries known to have the disease. Many (~90) of these interceptions were destined for propagation.

Survey CAPS-Approved Method*: The CAPS-approved survey method is to collect symptomatic plant tissue by visual survey.

*For the most up-to-date methods for survey and identification, see Approved Methods on the CAPS Resource and Collaboration Site, at http://caps.ceris.purdue.edu/.

Literature-Based Methods: Apothecia production from leaf material: Apothecia have been produced under laboratory conditions from outdoor overwintered symptomatic leaves. The leaf residues were thoroughly washed in distilled water and soaked for one hour. The smoothed leaf tissue pieces were placed, partly upper side up; partly lower side up, into Petri dishes, on a two-fold moistened filter paper. The leaf tissue pieces and the filter paper layer were saturated and kept at room temperature (18 to 20°C; 64.4 to 68°F) under scattered light. On slight magnification, development of apothecia was visible on the leaf residue after three or four days. After six to eight days, the apothecia obtained full development (Lehoczky, 1966).

Apothecia production on agar media: Apothecia form on some agar media (malt agar, Vitis-stem extract medium) but not on other media (potato dextrose agar) (Korf et al., 1986). Apothecia can also be produced by placed surface sterilized symptomatic grapevine leaves overlaid on malt extract agar (Plant Health Australia, 2004).

Spore trapping: In Europe, monitoring of the ascospore release by means of spore traps enables disease forecast and timed fungicide application (Pearson et al., 1991; Konig et al., 2009).

Key Diagnostics/Identification CAPS-Approved Method*: Morphological. The diagnostic feature of the fungus is the presence of sine-wave pattern of hyphae observed in the xylem vessel elements when affected plant tissues are cleared by boiling in 2% aqueous potassium hydroxide for 2-3 min (Plant Health Australia, 2004) and the production of eight ascospores within asci/apothecia.

Literature-Based Methods: Culture: The anamorph of the fungus can be grown plating surface sterilized plant material on 2.5% malt extract agar (Plant Health Australia, 2004).

235 Pseudopezicula tracheiphila Primary Pest of Grape Fungus Rotbrenner

Easily Confused Pests Pseudopezicula tetraspora, a four-spored American species, causing angular leaf scorch has similar symptoms to P. tracheiphila (Pearson et al., 1988). Diagnostic features for these two fungi are similar, but P. tetraspora only produces four ascospores in contrast to P. tracheiphila that produces eight. Unlike other members of Peziculoideae, which attack woody stems and branches, these two species occur on leaves, mostly in association with the vascular elements.

References Holz, B. 2000. Integrated control of rotbrenner (Pseudopezicula tracheiphila (Müll. –Thurg.) Korf & Zhuang (Discomycetes Helothiales) in the wine growing region Moselle-Saar-Ruwer. Integrated Control in Viticulture IOBC/wprs Bulletin Vol. 23(4) pp. 59-62

Konig, H., Unden, G., and Frohlich, J. 2009. Biology of Microoganisms on Grapes, in Must and in Wine: 4.3.3 Pseudopezicula tracheiphila.

Korf, R.P., Pearson, R.C., Zhuang, W., and Dubos, B. 1986. Pseudopezicula (Helotiales, Peziculoideae), A new discomycete genus for pathogens causing an angular leaf scorch disease of grapes (“rotbrenner”). Mycotaxon 26: 457-471.

Lehoczky, J. 1966. Sexual spore formation of Pseudopeziza tracheiphila and its role in causing epidemics. Szoloes Gyumolcstermesztes (Grape and Fruit Production) 2: 57-68. English Abstr.

Müller-Thurgau, H. 1913. Der rote Brenner des Weinstockes. Centralblatt fur Bakteriologie Abt. II 38:1-36, pl. 1.

Pearson, R.C., Dubos, B., and Korf, R.P. 1986. Occurrence of Pseudopeziza tracheiphila, causal agent of rotbrenner, on grape in New York (Abstr.). Phytopathology 76: 656-657.

Pearson, R.C. Smith, F.D., and Dubos, B. 1988. Angular leaf scorch, a new disease of grapevines in North America by Pseudopezicula tetraspora.

Pearson, R.C., Siegfried, W., Bodmer, M., and Schuep, H. 1991. Ascospore discharge and Survival in Pseudopezicula tracheilphila, causal agent of rotbrenner of grape. Journal of Phytopathology 132:3 pp. 177-185.

Plant Health Australia. 2009. Pest Risk Review: Rotbrenner. Viticulture Industry Biosecurity Plan 2009.

Plants for Planting Manual. 2012. United States Department of Agriculture, Animal and Plant Health Inspection Service, Plant Protection and Quarantine. Last accessed June 24, 2013. http://www.aphis.usda.gov/import_export/plants/manuals/ports/downloads/plants_for_planting.pdf.

USDA-ARS. 2013. Fungal Database. Pseudopezicula tracheiphila. Queried July 30, 2013.

Reviewer(s): Michael Maixner, Julius Kühn Institute, Siebeldingen, Germany.

236 Appendix A: Diagnostic Resource Contacts

Appendix A: 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 Metz 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 Entomologist - Domestic Identifier USDA, APHIS, PPQ CAPS Office 1911 SW 34th Street Gainesville, FL 32614-7100 Office: (352) 372-3505 ext. 438 Appendix A: Diagnostic Resource Contacts

Fax: (352) 494-5841 [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]

238 Appendix B: Glossary of Terms

Appendix B: 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.

Agglomerate: To form or collect into a rounded mass.

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): are modified wing scales on butterflies and moths that release pheromones. Only males have these scent scales.

Apophysis: A natural swelling or enlargement found at the base of the stalk or seta in certain mosses or on the cone scale of certain conifers.

Areolated: Divided into small spaces or areolations; usually pertains to the cuticle of a nematode.

Arthrospore: A fungal spore resulting from the fragmentation of a hypha.

Bolls: The spherical shaped fruits of cotton and flax.

Calyx: The outer-most group of leaves surrounding the flower; the external-most part of the flower.

Cerarii: These are characteristic of mealybugs and consist of groups of large setae, usually conical, on the lateral margins of the body.

Chorion: The outer shell or covering of the insect egg.

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.

Cisanal setae: In coccids, the shorter and further two of the four setae (commonly known as hairs) near the caudal ring.

Clamp connection: A bridge- or buckle-hyphal protrusion in basidiomycetous fungi, formed at cell division and connecting the newly divided cells.

239 Appendix B: Glossary of Terms

Cockchafer: Any of various large European beetles destructive to vegetation as both larvae and adult.

Coleoptile: A protective sheath enclosing the shoot tip and embryonic leaves.

Cornuti: Literally, the horned ones, which was the name given to a Byzantine auxilium palatinum.

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.

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.

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.

Digitus: Having appendages of the feet (as found in member of the family Coccidae), which may be either broadly dilated or knobbed hairs; tenent hairs, empodial hairs.

Discal: On or relating to the disc of any surface or structure.

Ecdysis: Molting; the process of shedding the exoskeleton.

Endocarp: The hard inner layer of the pericarp of some fruits that contains the seed.

Epiphytotic: Epidemic among plants of a single kind, especially over a wide area.

Exuvia: The cast skin of an arthropod.

Fecundity: The number of offspring per number of potential offspring (e.g., eggs).

Filiform: Thread-like or hair-like.

Flaccid: lacking in strength or firmness or resilience.

240 Appendix B: Glossary of Terms

Frass: Plant fragments made by a wood-boring insect usually mixed with excrement; solid larval insect excrement.

Gubernaculum: Nematodes: Spicule guide; sclerotized accessory piece.

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.

Hyaline: Like glass, transparent and colorless.

Incipient: Beginning to exist; coming into existence.

Inflorescence: A characteristic arrangement of flowers on a stem; a flower cluster.

Leeward: On the side away from the wind.

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).

Lychees: The fruit of a tree native to China. It is nutlike, having a rough but tender shell and containing an aromatic pulp and a single large seed.

Metanotum: The upper surface of the third (posterior) thoracic segment (metathorax).

Micropile: A very small opening in the outer coat of an ovule, through which the pollen tube penetrates; the corresponding opening in the developed seed; one of the minute openings in the insect egg, through which spermatozoa enter in fertilization.

Microtrichium (pl. microtrichia): Small, sclerotized, and non-innervated cuticular projects on the body and wings of insects; which are also found on the tracheae.

Monandrous: Having only one male mate at a time.

Multivoltine: Pertaining to organisms with many generations in a year or season.

Neonate: A recently born larva.

Ocelli: A simple eye of an insect or other arthropod.

Orbicular: Circular in outline.

241 Appendix B: Glossary of Terms

Ostioles: In Heteroptera, one of the lateral metasternal external openings of the stink gland, placed near the coxa in the adult, but paired and dorsal on the abdomen of the nymph. For plant pathology, pore; opening in the papilla or neck of a perithecium, pseudothecium, or pycnidium through which spores are released

Oviparous: Reproduction in which young hatch from eggs.

Palearctic: The biogeographic region including Europe, Asia north of the Himalayas, and Africa north of the Sahara.

Parthenogenetic: Capable of reproduction without mating or male fertilization.

Parthenogenesis: Process of reproduction by the development of an unfertilized egg.

Pedicel: Small slender stalk; stalk bearing an individual flower, inflorescence, or spore.

Phasmids: Any of various stick-like or leaf-like insects, including walking sticks and leaf insects.

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.

Phytophagous: Plant-eating.

Pinaculum: In caterpillars, an enlarged seta-bearing papilla forming a flat plate.

Pleural membrane: A thin serous membrane in mammals that envelopes each lung and folds back to make a lining for the chest cavity.

Polygynic: Phenotypic trait whose expression is controlled by, or associated with, more than one gene.

Polyphagous: Eating many kinds of food.

Pome fruits: A fleshy fruit, such as an apple, pear, or quince, having several seed chambers and an outer fleshy part largely derived from the hypanthium. Also called false fruit.

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.

242 Appendix B: Glossary of Terms

Protonymph: Second instar of a mite.

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.

Reniform: Kidney-shaped.

Reticulate: Descriptive of surface sculpture, usually the insect’s integument, which is covered with net-like lines.

Saccus: A sac.

Sclerotized: Hardened.

Senescence: The last stage in the post-embryonic development of multicellular organisms, during which loss of functions and degradation of biological components occur. A physiological ageing process in which cells and tissues deteriorate and finally die.

Sessile: Not supported on a stem or footstalk; immobile.

Setae: Bristles; commonly known as hairs.

Sign: Indication of disease from direct observation of a pathogen or its parts (see symptom).

Spatulate: Rounded and broad at the top, attenuate at base. Shaped like a spoon, with a narrow end at the base.

Spiracles: Breathing pores; in the plural the lateral openings on the segments of the insect body through which air enters the trachea.

Sternites: The ventral shield or plate of each segment of the body of an insect or other arthropod.

Striated: Numerous parallel, fine, and impressed lines.

Stylet: A stiff, slender, hollow feeding organ of plant-parasitic nematodes or sap- sucking insects, such as aphids or leafhoppers.

243 Appendix B: Glossary of Terms

Symptom: Indication of disease by reaction of the host, e.g. canker, leaf spot, wilt (see sign).

Tarsus: The leg segment immediately beyond the tibia, consisting of one or more segments or subdivisions.

Tegument: Lepidoptera: the tergum in male genitalia. A structure shaped as a hood or inverted trough, positioned dorsal of the anus; the uncus articulates with its caudal margin, derived from the ninth abdominal tergum.

Tergite: A dorsal sclerite or part of a segment, especially when such part consists of a single sclerite.

Termen: The outer margin of a wing, between the apex and the posterior or anal angle.

Tillers (subterranean): A lateral shoot, culm, or stalk arising from a crown bud; common in grasses.

Univoltine: Having only one generation per year.

Variegated: Varied in color, of several colors in indefinite patterns.

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 copulatation.

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