Research Project Title: Thrips TSWVIINSV Control Systems
Thrips and the tospoviruses they transmit represent a formidable pest problem facing floricultural producers across the US. Here we propose a researchlimplementaflon program which combines recent research with the development of new information to combat this problem in a multi-disciplinary/multi-state research proposal.

Project Areas and Goals:

1. Samphng/ldentification
a Validationlimplementation of a monitoring/identification program for thrips across a variety of floriculture crops. This program, developed in California, will be evaluated in the western US in addition to Vermont and Texas.
b. Economic thresholds for western flower thrips have been developed for roses and research will extend this to other crops including impatiens and potted chrysanthemums.
c. The samplinglthreshold work will include development of a new monitoring system for detecting the presence of viruliferous thrips in samples. This system and a new immunology method for tospoviruses will be used to provide growers with an early warning system for tospoviruses to assess thresholds for viruliferous thrips.

2. Pesticides
a. Develop practical recommendations for more effective use of the fungus


Beauveria



b. Evaluate new ‘reduced risk’ pesticides as they become available
c. Determine compatibility with natural enemies of thrips and other pests

3. Biological control of Thrips
a. Evaluate recommendations for use of commercially available natural enemies

b. Evaluate the entomopathogenic nematode,

Thripinema siddiqui,


and parasitoids in the genus


Ceranisus

c. Develop/evaluate a mechanical delivery system for natural enemies

4. Postharvest Control

a. Evaluate different methods of postharvest disinfestation techniques

b. Develop new methods using novel pesticides and/or controlled atmospheres

Researchers:

Dr. Michael P. Parrella
Dr. Diane Ullman
Dr. Brook Murphy
Dr. Elizabeth Mitcham
Dr. Ken Giles
Dr. Arnold Hara
Dr. Kevin Heinz
Dr. Michael Brownbridge
Dr. Karen Robb
Ms. Julie Newman
Mr. Steve Tjosvold
Department of Entomology, UC Davis
Department of Entomology, UC Davis
Department of Entomology, UC Davis
Department of Pomology, UC Davis
Department of Biol. & Agric. Engineering, UC Davis
Department of Entomology, University of HI, Hilo
Department of Entomology, Texas A&M, College Station
Department of Entomology, University of VT, Burlington
UC Cooperative Extension, San Diego Co.
UC Cooperative Extension, Ventura and Santa Barbara Cos.
UC Cooperative Extension, Monterey and Santa Cruz Cos.

Proposed Project Duration Start: 7/1/98 Completion: 7/1/02

Total Estimated Cost
Year 1: $126,000 Year2: $192,000 Year3: $196,000 Year 4: $200,000

Note: This is a multi-year proposal, but only YEAR 1 will be described in the ensuing pages. This is a necessity because of space limitations. As it is the proposal is too long.

 

1

Executive Summary

 

Correct and timely identification of thrips attacking floriculture crops in the greenhouse and field is the backbone of a complete Integrated Pest Management (IPM) program for this pest. A monitoring system that helps growers locate infective thrips and their sources is also essential to forging effective 1PM strategies. We propose to validate sampling plans and identification keys developed over the past few years across diverse areas of the US and to incorporate technology to assess the capability of thrips to vector TSWV/INSV. Validation work will commence simultaneously with research to develop sampling protocols and economic thresholds for other crops (including impatiens and potted chrysanthemums). As cultivars with ‘resistance’ or ‘tolerance’ to the tospoviruses become available, these will be included in our development of economic thresholds. Continued evaluation of reduced risk pesticides will be an important part of this proposal, as well as developing recommendations for their effective use by growers. We know that Beauveria bassiana

is an effective thrips control material, but we propose additional research to further develop use protocols for growers. We expect there to be many new materials in the next few years, including several other species of entomopathogenic fungi. Many of these reduced risk materials alfozl compatibility with natural enemies, and we will evaluate their compatibility with commercially available natural enemies of thrips and other greenhouse pests. Although natural enemies of thrips are available commercially, there are few studies documenting their effectiveness in floriculture crops. An evaluation of commercially available natural enemies for thrips control in floriculture will be initiated as well as a search for new natural enemies. A more effective delivery system for natural enemies in the greenhouse will also be investigated. Finally we will evaluate the potential of controlled atmospheres in combination with heat and/or reduced risk pesticides to provide 100% mortality of thrips after flowers are harvested.

This project is multifaceted in nature-it provides elements growers can use today to help solve their thrips/virus problems such as implementation of monitoring/sampling programs and more effective use of reckiced risk pesticides and commercially available biological control agents. The proposed research will 1) expand sampling/monitoring programs onto other major crops, 2) address thrips as vectors of tospoviruses, 3) search for new effective natural enemies and develop delivery systems for their use, 4) expand the search for ~ risk pesticides and, 5) develop protocols for postharvest control of thrips in cut flowers. Many of the components of this proposal go far beyond thrips control and have broad applicability to control of many other key floricuitiuc pests.

 

2

Description and Objectives

 

Progress to Date & Future Anticipated Benefits

 

Materials and Methods

 

Objective 1. Sampling/identification

 

a. Validation I implementadon.

Previous work has established that sticky trap captures of thrips are indicative of the number of thrips feeding on roses and chrysanthemum. The results demonstrated that pest management decisions for thrips can be based on the use of traps alone. From these data we estimated the number of traps needed to monitor thrips with a constant level of precision for commercial operations. This portion of the objective will validate that the required number of traps is both accurate and economical for determining thrips abundance in the crop. Furthermore, under this objective we will validate identification keys for identifying thrips species caught on traps or within the crop.

 

In cooperation with commercial growers and local farm advisors we will validate trap sampling prccedures in at least two commercial operations in three distinct geographical regions of the US At each location, sampling procedures will be applied in at least two greenhouses. Traps will be deployed according to procedure guidelines (8 to 9 traps per greenhouse, hung in the center of benches or beds 4 to 6 inches above the crop). Traps will be removed and replaced weekly with new traps. Traps will be returned to the laboratory and the identification and number of thrips recorded. Each week chrysanthemum or rose samples will be removed from the greenhouse and returned to the laboratory where the numbers of thrips in the crop will be recorded. Trap capture data will be correlated with thrips numbers in the crop to determine if trap sample numbers accurately reflect thrips numbers in the crop.

 

Upon successful validation of sticky traps as a monitoring tool, we will begin implementation of the sampling program using sticky traps to monitor thrips and economic thresholds to guide pest management decisions. Using the same cooperators as above, we will compare thrips control using the monitoring procedure with conventional pest control practices. Success or failure will be determined by comparing the number of pesticide applications reqtered with and without monitoring, the quality of the harvested crop, and the cost of controlling thrips.

 

3



I

b. Economic thresholds.


The relationship between thrips density and crop damage have been determined for roses and garden chrysanthemums. Under this objective we will expand this work to include impatiens and potted chrysanthemums. Relationships will be established by monitoring the cumulative number of thrips on the plant and the cumulative degree of damage occurring on the plant. The results will establish the thrips density that can be tolerated before significant economic loss is encountered.

 

In cooperation with commercial growers we will monitor tlrrips densities in impatiens and potted chrysanthemums. Trials will be conducted in at least two greenhouses in two geographical areas. At least three different commonly grown cultivars of impatiens or chrysanthemums will be evaluated. For each cultivar in each greenhouse, thrips numbers will be monitored by removing whole plants and returning them to the laboratory where thrips will be washed from foliage, identified and t~eir numbers recorded. The plant material will then be examined and the amount of feeding damage quantified (using a rating system developed for roses this past year). The procedures will be repeated for spring and summer thrips populations. Results will be statistically correlated to determine the relationship between thrips density and damage.

 

C. Viruliferous thrips monitoring.


The monitoring program for thrips discussed above can be modified to include the threat of TSWV or INSV transmission by thrips. This research will benefit the floricultural industry by providing growers with a tool for early detection of infective thrips and locating their sources. The information gained about how numbers of tospovirus vectors relate to virus incidence and spread will lead to more precise systems for making decisions and may reveal new control methods. During our fffst year of funding we have made significant progress toward developing and field testing a monitoring system that combines directional yellow sticky traps and petunia indicator plants (Robb et al. 1998). We have shown that ELISA is useful for quantifying thrips vectors, but that adequately reliable and rapid information can be acquired by growers using directional traps and indicator plants, a technique requiring less technology and cost (i). We plan to use ELISA as a resech tool to address the relationship between infective thrips numbers and disease incidence (ii) -~ in essence this similar to the economic threshold work described above, but will consider threshold levels in light of viruliferous thrips being in the greenhouse. Our most intensive efforts will focus on refining the directional trap/indicator plant monitoring system as an integrated pest management tool for field grown flowers (i.e. determining the minimum number of

 

4

 

I

monitoring stations needed to provide adequate information) and adapting the monitoring system to other production environments (i.e. greenhouses).

Determine and compare the efficacy of detecting infective thrips with ELISA and petunia indicator plants for use as tools for monitoring tospovirus vectors in floricultural crops. This subobjective will be expanded to test the monitoring system in different production environments. A large flower production area in southern California (San Diego county) that has been alflicted with serious TSWV epidemics has been the site of our experiments and we have expanded to a greenhouse production area of approximately 10,000 sq. ft. In the field, six trapping stations will be placed at the edges of the production area and within the blocks of flowers, a split plot design will be used to compare two regimes. One with five trapping stations (5/trapping stations/replication x 6 replicates) and one with two trapping stations (2/trapping stations/replication x 6 replicates). In the greenhouse production area, a similar strategy will used except special stations will be placed at doors to the house. Each trapping station will have a directional trap kept at crop canopy (post with adjustable height clips and two, tw~sided sticky cards placed such that thrips moving from the north, south, east or west will be trapped) and a petunia indicator plant on a blue background (an attractive color to thrips), kept at crop canopy on an adjustable height platform. Sticky traps and plants will be collected and replaced weekly. Thrips will be counted, identified, removed from sticky cards and tested with ELISA for the TSWV NSs proteins (as in Bandla et al. 1994). Local lesions will be counted on the petunia and thrips feeding damage noted. The monitoring program will be continuous (12 months/year) to look for trends in the presence and location of tospovirus vector sources. Normal grower practice will continue throughout the project. Comparison of data obtained with each method will be made using ANOVA. Results from the first year of study show that monitoring with directional traps and indicator plants provide rapid information for grower use without the technical challenges of conducting ELISA. The ELISA test is still important, but will be used most intensively to address subobjective ii. Funds from another a source are currenfly being used to develop a different serological method called a direct tissue blotting assay for detecting infected plants and insects (Robb et al. 1998). If successful, this method may prove more useful to growers and will be incorporated into the ongoing research proposed here.

 

Determine the relationship between numbers of infective thrips, sources of infective thrips and disease incidence. The same location and procedures desccibed in subobjective (i) will be used.

 

5

Crop plants will simultaneously be collected for TSWV ELISA testing and estimation of TSWV incidence. Our method involves identifying 100 plants/block in a uniform distribution. Visual symptoms are recorded and plants sampled monthly for testing in groups of 10 by TSWV ELISA. When TSWV is detected in a group, an additional test of the individuals in the group is done to determine which plant(s) was infected. Once a plant is infected (two positive ELISA tests), it is not tested again. This methodology has been used successfully by others (Kumar et al. 1995a; Kumar et al. 1995b) and has worked well in our trials last year. We have expanded testing at some locations to include TSWV and INSV. Multiple regression analysis will be used to correlate the thrips density, the proportion that are infective and lesions on petunia with the incidence of virus infection in the crop.

 

Objective 2. Pesticides

 

a. Microbials:

Beauveria bassiana.


This pathogenic fungi has been found to be quite effective against aphids, thrips and whiteflies in commercial greenhouses. However, guidelines for the effective use of this product is critical for proper performance in commercial operations. In this objective we will optimize field recommendations that maximize effectiveness and reliability at the least cost.

Develop practical recommendations for the use of Beauveria bassiana for thrips management.

 

Less than 1% of a pesticide that is applied actually reaches the target: 99% becomes environmental pollution and an expense without benefit (Pimental & Levitan 1986). There is a huge opportunity to increase efficacy and reduce pesticide use by improving and optimizing application protocols; spray droplet size, pressure and volume directly affect penetration and deposition on foliage, and are direcdy influenced by the type of sprayer and spray protocols used (Lindquist & Powell

1995).


Very few studies document how fungal deposition, and efficacy, is affected by sprayer type, and how protocols can be altered to enhance coverage and host contact of insect-killing fungi such as


Beauveija ba:ssia~

Hydraulic sprayers are commonly used in greenhouses. They are inexpensive and well-suited to “spot~’ applications. They have been used to successfully apply fungi to control whiteflies and thrips (van der Schaaf et al. 1991, Butt & Brownbridge 1997). Air-assisted electrostatic sprayers can provide better penetration of the crop canopy and more uniform coverage than hydraulic sprayers (Palumbo & Coates 1996). In addition, problems of drift and run-off are reduced and lower spray volumes are used. However, although deposition may be improved, better insect control does not always follow (Cayley et al. 1984). Fungi are living insecticides, and other factors ate

 

6

critical to successful infection. Electrostatic sprayers cannot be assumed to work equally well for all products. It is, therefore, essential to evaluate different spray techniques and devise practical guidelines for

their effective use.

 

Materials and Methods. An electrostatic and a standard hydraulic sprayer will be evaluated. Water-and oil-based formulations of B. bas~iana will be obtained from Mycotech Corp., Butte, MT and tested using a series of standard protocols. All sprays will be tested for viability before application. Fungi will be sprayed as aqueous suspensions using 0.02% Silwet as a wetting agent. Recommended rates of application will be used and each sprayer will be calibrated to deliver the same number of conidia per unit are~

 

Greenhouse trials will be done on chrysanthemums to identify factors affecting coverage, then repeated on impatiens. Results will show if spray parameters suitable for one crop apply to the other, despite distinctly different leaf structures. Even-aged plants will be arranged in blocks of 32 in a greenhouse and replicate sprays made on teree consecutive days for each variable. After each spray, 10 plants will be randomly sampled from each block and coverage assessed using a unique method recenfly developed in our laboratory (Gouli et al. in preparation).

 

The following sprayer-specific adjustments will be made to assess their effect on spray coverage and spore

 

deposition 1) Electrostatic sprayer: pressure - 5 and 10 psi; orifice discs - #20 and #40; distance above the crop -

 

15, 30 and 45 cm but maintain a distance of 1.5

m from each row. Application time will be held constant and

determined on a per plant basis prior to testing; 2) Hydraulic sprayer using the spray gun and 5-nozzle lance

 

attachinents - pressure 150 and 250 psi; nozzle sizes - Teejet no 6004 and 6008; distance above the crop - 15, 30 and

 

45 cm using the spray gun and directed vertically at bench height using the 5-nozzle lance.

 

The mean number of spores/unit area of leaf surface will be compared for differences using ANOVA. Means will be separated using Tukey’s test to reveal differences between formulations, method of application, deposition on upper and lower leaf surfaces of mums and impatiens. Data will be interpreted with assistance from the Experiment Station Statistician and incorporated into plans.for pilot testing in larger scale greenhouse trials for efficacy against thrips.

 

Laboratory evaluation of compatibility of Beauveria bassiana with reduced-risk pesticides.

 

Biological control and reduced-risk pesticides will play an increasingly important role in future floral 1PM programs. Such initiatives will rely on the concurrent use of several control strategies to effectively regulate a range

 

7

of insect pests. It is critical that we understand the interactions between the different components of an 1PM program to develop management strategies that are efficient and cost-effective. For example, if the materials are fungicidal, tank-mix spray applications should not be made and spray applications should be scheduled aceordingly (see Majchrowicz & Poprawski 1993, Li & Holdam 1994). Fungal efficacy may also be improved if used in conjunction with selected materials that can promote the infection process, either as a result of their effects on the insect cuticle, through promotion of fungal germination, or simply stressing the insect and pee-disposing it to infection (Ar~derson et al. 1989, Hassan & Chamley 1989, Joshi & Chamley 1992, Lindquist 1993, Quintela & McCoy 1996).

Here we will assess the compatibility of insect-killing fungi with reduced-risk insecticides being evaluated in other sections of this project. Initially, laboratory tests will be done to determine effects of these materials on spore germination to predict any potentially negative effects on insect infection.

 

Materials and Methods: i. Fungal strains and culture. B. bassiana

726 (OHA strain) and


M. anisopliae ESC- 1 will be used throughout. This strain of B.


bassiana has recendy been registered as BotaniGard® (Mycotech Corp., Butte, ~ and has been brought to market this year.


M. arnsoplzae


ESC 1 is currently registered by EcoScience Corp. ~ast Brunswick, NJ) for greenhouse use, but is not yet available. The fungi have different host spectra, so their inclusion in the trials will reveal differences in sensitivity of two species to the different test materials, and whether combined applications enhances infection.

Tests will be carried out on non-formulated fungi only. For all trials, fungi will first be grown on quater strength Saboraud dextrose agar supplemented with

0.25%


wlv yeast extract (SDAY) and incubated at


220


C for 1O~ 14 d. Conidia will be harvested in


2.5%


Tween 80, centrifuged, resuspended in sterile distilled water and held at 40 C until use. ii. Selection of insecticides. Reduced-risk insecticides will be provided by UC Davis, along with information on recommended dose rates and preparation. Experiments on compatibility will be carried out using four

insecticide doses: X (recommended rate), 0.5X, 0.25X and 0.

lx. iii.


Effect on spore germination. Conidia will be suspended in the pesticides at 106/ml. 25 micro-liter samples will be transferred to tenth-strength SDAY and overlaid with a sterile glass cover slip. Spore germination rate will be assessed after 24, 48 and 72 h using phase contrast microscopy. Controls will be set up using fungi suspended in sterile distilled water only. This test represents ‘worst-case’ test conditions for the fungi, but will reveal potentially positive or negative interactions between the insecticide and the spores that could influence the infection process. iv. Data analysis. Data ge:atel

 

8

r

in each evaluation will be separately analyzed using ANOVA and the means compared using Duncan’s multiple range test to allow comparisons to be made between germination rate in insecticide and control treatrnents, and between pesticides.

 

Objective 3: Biological Control of the Western Flower Thrips

 

Approach to Development of a Biological Control Component to Thrips Management

 

Candidate Selection. Successful biological pest control is frequently hampered by the use of poorly adapted natural enemies or the inappropriate release of well adapted ones. Thus, one goal will be to (1) identify those natural enemies with a suite of life history characteristics identifying them as potential candidates for subsequent field testing (this method has been used successfully with other pest-natural enemy systems by Heinz and Parrella), and (2) to develop mass-rearing practices of new natural enemies that will keep producer costs low but will maximize natural enemy quality.

 

Release Strate~es. As demonstrated in Heinz’s previous AFE-funded project on efficient release strategies for aphid biological control, proper release strategies must be developed to achieve the best possible level of control for a specific number of natural enemies released. Development of efficient release strategies will be the second phase of the program.

 

Evaluation Procedures. For biological control of WFT to be effective, it must (1) reduce the physical damage caused by WFT feeding, (2) reduce the potential or actual levels of TSWVIINSV occurring within greenhouses, and (3) be economically cost effective. Each of these parameters will be tested within the context of commercial production schemes.

 

Material and Methods

 

Several species of natural enemies with life histories conducive to thrips biological control have not been studied within the context of greenhouse and nursery floriculture production. We will study these natural enemies and they include the parasitic wasp Ceranisus menes, the predaceous mite Iphisejus (Amblyseius) degenerans, and the entomopathogenic nematode Thripinema nicklewoodi.

 

Clari~ hos: and natural enemy relationships. To determine the relationship between thrips life stage and

 

natural enemy attack rates, the five life stages of WIPI’ will be uniformly distributed among the apical meristem,

J

leaves, and soil of potted plants. A known number of natural enemies (1 wasp per plant, 10 mites per plant, and

j

9

 

J

10


nematodes per


Ctn2


of leaf surface area) will be released onto the potted plants. WFr will be recollected after 24

hours to assess thrips survivorship by life stage, and all thrips will be dissected under a microscope to determine the

numbers of infected (1,y nematodes) or parasitized (by the wasps) WFF by life stage and location within the plant.

There will be 10 replicates of each life stage and an untreated control, and tests will be performed at 22~C and

60% rh.

f

 

r

f

Each

WFT


life stage successfully parasitized by


T. nicklewoodi


will be studied in detail to determine the effect of


T nicklewoodi


parasitism on


WET


life history characteristics. Because the mites and parasitic wasps kill their hosts outright, similar studies will not be performed with these natural enemies. Groups of 10 thrips each will be housed in a modified munger cell clamped to a chrysanthemum leaf. The thrips will then be treated with a single application of


T. nicklewoodi


or biological saline (as a control). Each treatrnent will be replicated 30 times with each munger cell acting as a replicate. The thrips within each munger cell will be censused daily for thrips development and mortality, and the munger cells will be transferred to new uninfected leaves every other day. Six days after each transfer, the previously exposed foliage will be assessed for the numbers of feeding scars, thrips larvae, and uneclosed thrips eggs.

For all life stages attacked by each of the natural enemies, a dose response curve will be established by exposing 30 groups of 20 WET to each of five different concentrations of infective

T nicklewoodi


or release rates of mites or wasps. Natural enemies will be introduced into small cages containing WET infested plants, held there for 24 hours, and WET numbers censused immediately after. Thrips potentially infected by


T. nicklewoodi


or parasitized by


C. menes


will be transferred individually to small vials for three days. Upon completion of this incubation phase, the WET will be dissected to assess parasitism by


T nicklewoodi or C. menes.

Environmental Constraints Natural Enemy Attack

To assess the importance of plant phenology on the effectiveness of nematede applications, plants in three & stages of development (vegetative, flower bud formation, and open flower) will be infested with WET life stages

 

susceptible to attack by each of the three natural enemies. Three individuals per leaf, bud, or flower will be placed on individual plants. One hour after infesting the plants with thrips, a standard nematode dosage or mite or wasp

 

release rate will be applied to test plants to observe the efficacy of control relative to the plant part. Thirty pots per plant and thrips development stage will be tested per natural enemy under conditions of 22~C and 60% di. Every

 

10

leaf, bud, and flower of every plant will be sampled 24 his after application of each natural enemy and the status of each thrips aesessed.

Using the best dosage or release rate, the most susceptible life stage to each natural enemy, and plant growth stages yielding significantly different levels of predation or parasitism, trials will be run in growth chambers to determine the influence of temperature and humidity on parasitism or predation rates of WFT. Four treatments will be tested: (1) warm, wet (30cC, 90% rh), (2) warm, dry (30~C, 30% rh), (3) cool, wet (20⁰C, 90% rh), and (4) cool, dry (20⁵C, 30% rh). Plants will be pre-infested (one hour prior to the test) with 100 WET of the desired thrips life stage, subsequently treated with a standard density of natural enemies, and immediately returned to the chambers. At 48 his, the foliage and flowers of the test plants will be examined for WET, and in the cases of C. menes and T. nicklewoodii

dissected to detect parasitism. There will be five plants for each treatment, replicated four times.

Results obtained from Objective I will

be


compared to data existing for other commercially available natural enemies. Natural enemies will be assigned ranks aceording to their performance for each variable used in making comparisons among species. Measurements of agreement among the rankings of the natural enemies will be obtained through calculation of Kendall’s coefficient of concordance


(W) (Statsoft 1994). Values of W


can range from 0 (no concordance) to 1 (perfect concordance). Based upon this analysis, natural enemies ranking high in each of the performance variables will be considered ideal candidates for flirther testing. Conversely, natural enemies ranking low each of the performance variables will be discarded from further consideration.

A reliable supply of new natural enemies (from Subobjective 1) to be considered for in-depth testing (from Objective 2) will be needed for development of an effective biological control program. Each of the natural enemies will be reared on its natural host, WET. Modificctions of the simple rearing method of Doane

et al. (1995)


will be made to develcp a low~ost rearing procedire to enable resenich on WET biological control. Using locally obtai red and inexpensive materials ~oly~pylene feezr containers, bean leaves grown from food grade red kidney beans, bee pollen non~dorant feminine napkins, and purified water) cohorts of unifoim age will be produ~d Such cohorts are necessary for relia~e production of high c~iality naturil enemies. Dr. Heinz, has environmentally controlled isolation rooms as part of his laboratory at Texas A&M that can be devoted to this mass-rearing. Cost inputs and natural enemy yields will be continuously monitored to generate an economic assessment of the mass rearing programs.

Objective 4: Postharvest Control

 

11

Shipment of cuttings and other ornamental products nationally and internationally necessitates reliable control of insect and mite pests which may be present ~ara 1994). Current methods often utilize methyl bromide fumigation which can cause some product damage. In addition, with the planned phase-out of methyl bromide use in the US in 2001, alternative means of insect control are neede~ The major insect pests of concern on chrysanthemum cuttings are melon aphid (Aphis gossypii),


sweetpotato whitefly


(Bemisia tabacii) and serpentine lealminer (Liriomyza tnfolii), and western flower thrips (Frankliniella occidentalis).


This project will focus on control of these three pests. Initial studies will be done on rooted and unrooted chrysanthemum cuttings will expand to cut flowers.

Controlled atmospheres employing high carbon dioxide and/or low oxygen atmospheres have shown promise for control of various insect and mite pests. Treatment efficacy is determined by the concentrations of carbon dioxide and oxygen, temperature, and length of exposure. Treatments at warmer temperatures are shorter in duration, however treatment at low temperatures may be better tolerated by the host plant. Feasible treatment parameters are determined by tolerance of the host plant to insecticidal controlled atmosphere treatment, the duration of typical post-cutting life of the host plant and compatibility with commercial practices.

Previous research has demonstrated several potential treatments for insect control utilizing controlled atmospheres. Mortality of 5th instars of codling moth, one of the most important of quarantine species, 100% after a 48-hour exposure to 95% carbon dioxide at 68 degrees F. Complete mortality of western flower thrips on strawberries was obtained by applying 90% carbon dioxide for two days at 36 degrees F. A five day treatment with 60% carbon dioxide at 32 degrees F gave complete control of New Zealand flower thrips and green peach aphid on asparagus.

In our laboratory, we have developed an eight day treatment with 45% carbon dioxide at 35 degrees F to control three pests on table grapes: omnivorous leafroller, western flower thrips and Pacific spider mite. Additional research with two spotted spider mite indicates that exposure to very low oxygen concentration at room temperature results in significant mortality.

In addition to the promise of controlled atmospheres for insect and mite control, research with cinaamic aldehyde indicates tremendous promise as a postharvest insect control treatment. Field experiments have shown that sprays containing as little as 0.3% cinnamic aldehyde resulted in nearly instant kill of a wide variety of inseets.

12

I

determined using gas chromatography. After various exposure periods, the container will be vented with fresh air and the cuttings with insects transferred to room temperature air for recovery.

 

For combination carbon dioxide and cinnamic aldehyde tests, the container with infested chrysanthemum cuttings will be flushed with known concentrations of carbon dioxide (6 to 20%) for several minutes. Liquid cinnamic aldehyde will then be injected into the container and allowed to vaporize. After specific periods of time, the container will be vented with fresh air and cuttings with insects transferred to room temperature air for recovery.

 

After a recovery period of 24 to 36 hours in room temperature air, insect mortality will be assessed by lack of movement when insects are prodded. In addition, the appearance of the mum cuttings will be noted to obtain preliminary information about plant tolerance to the treatments applied.

 

When promising insect control treatments are identified which also show promise for mum tolerance, the effect of these treatments on the mum cuttings will be determined using non-infested mum cuttings, both rooted and unrooted. Visual evaluation of the cuttings will be made immediately after treatment and after storage for approximately three and 10 days at 41⁰F. In addition, with assistance from Yoder Brothers, cuttings from the most promising treatments will be thoroughly tested for ability to root and grow normally.

 

Following our initial work with mum cuttings, the effective treatments with controlled atmospheres and/or cinnamic aldehyde will be expanded to other propagative materials and cut flowers. Flowers and plants will be tested for their tolerance to controlled atmospheres, cinnamic aldehyde fumigation, and heat treatment, and various combinations of the three treatments. Arnold Hara in Hawaii will contribute to the testing of tropical and subtropical flower tolerance and treatment efficacy. We have preliminary work on ‘Dendrobium’ orchids which indicated promising tolerance to high carbon dioxide atmospheres. The efficacy of these atmospheres for control of thrips must be determined. For controlled atmosphere treatments, flowers will be shipped from Hawaii to UC Davis for treatment in the Dept. of Pomology Postharvest Laboratory.

 

14

Literature Cited:

Anderson, T.E., A.E. Hajek, D.W. Roberts, H.K. Preistler, & J.L.Robertson. 1989. Colorado Potato Beede (Coleoptera: Chrysomelidae): effects of combinations of Beauveria bassiana with insecticides. 3. Econ. Entomol. 82:83-89.

Bandla, M.D., D.M. Westeot, D.E. Uliman, T.L. German, & J.L, Sherwood. 1994. Use of monoclonal antibody to the nonstructural protein encoed by the small RNA of the tomato spotted wilt tospovirus to identify viruliferous thrips. Phytopathology 84:1427-1431.

Butt, T. & M. Brownbridge. 1997. Fungal pathogens of thrips, pp. 399-434. In T. Lewis [ed.], Thrips as Crop Pests. CAB International, Wallingford, Oxon, UK.

Cayley, G.R., P. Etheridge, D.C. Griffiths, F.T. Phillips, B.J. Pye & G.C. Scott. 1984. A review of the performance of electrostatically charged rotary atomizers on different crops. Ann. AppI. Biol. 105:379-386.

Doane, E.N., B.L. Parker & Y. Pivot. 1995. Method for mass rearing even-aged western flower thrips on beans, pp.587-5⁹³. In B.L. Parker, 3. Skinner & T. Lewis, [eds.], Thrips Biology and Management. NATO ASI Series. Series A: Life Sciences Vol.276.

Hara, A. H. 1994. Ornamentals and Flowers, pp. 32⁹-³⁴⁷. In R.E. Paul & 3. W. Armstrong [eds.]. Insect Pests and Fresh Horticultural Products: Treatmens and Responses. CAB International, Wallingford, UK.

Hassan, A.E.M. & A.K. Chamley. 1989. Ultrastructural study of the penetration by Metarhizium anisopliae through Dimilin-affected cuticle of Manduca sexta. 3. Invertebr. Pathol. 54:117-124.

Joshi, L. & A.K. Chamley. 1992. Synergism between entomopathogenic fungi, Metarhizium spp., and the benzoylphenyl urea insecticide, teflubenzuron, against the desert locust, Schistocerca gregaria. BCPC Pests and Diseases Monograph 4C-5: 369-374.

Kumar, N.K.K., D.E. Ullman, & 3.3. Cho. 1995a. Frankliniella occidentalis (Thysanoptera: Thripidae) landing and resistance to tomato spotted wilt tospovirus among Lycopersicon accessions with additional comments on Thrips tabaci (Thy sanoptera: Thripidae). Environmental Entomology 24:513-520.

Kumar, N.K.K., D.E. Ullman, & 3.3. Cho. 1995b. Resistance among Lycopersicon species to Franklinielia occidentalis (Thysanoptera: Tlrripidae). Journal of Economic Entomology 88(4): 1057-1065.

Li., D.P. & D.G. Holdam. 1994. Effects of pesticides on growth and sporulation of Metarhizium anisopliae (Deuteromycotina: Hyphomycetes). 3. Invertebr. Pathol. 63:209-211.

Lindquist, R.K. 1993. Integrated insect, mite and disease management programs on greenhouse crops: pesticides and application methods, pp. 38-⁴⁴. In K. Robb & 3. Hall [eds.], Proc. 9th Conf. on Insect and Disease Management on Ornamentals. Del Mar, CA. Soc. American Florists, Alexandria, VA.

Lindquist, R.K. & C.C. Powell. 1995. Pesticide application methods and equipment for disease and insect management, pp. 94-⁹⁷. In M.L. Gaston, S.L. Carver & C.A. Irwin [eds.], Tips on the Use and Safety of Chemicals, Biologicals and the Environment on Floriculture Crops. Ohio Florists’ Association, Columbus, OH.

Majchrowicz, I. & T.3. Poprawski. 1993. Effects in vitro of nine fungicides on growth of entomopathogenic fungi. Biocontrol Sci. Technol. 3:321-336.

Palumbo, J.C. & W.E. Coates. 1996. Air-assisted electrostatic application of pyrethroid and endosulfan mixtures for sweet potato whitefly (Homoptera: Aleyrodidae)control and spray deposition in cauliflower. 3. Econ. Entomol. 89: 970-980.

Pimental, D. & L. Levitan. 1986. Pesticides: amounts applied and amounts reaching pests. BioScience 36:86-91.

Quintela, E.D. & C.W. McCoy. 1996. Enhancing germination and pathogenicity of entomopathogenic fungi to larvae of Diaprepes abbreviatus (Coleoptera: Curculionidae) using sublethal doses of imidacloprid, pp. ⁶⁶. In Abstracts, Society for Invertebrate Pathology 29th Annual Meeting, 1-6 Sept.1996. Cordoba, Spain.

Robb, K., C. Casey, A. Whitfield, L. Campbell, & D. Ullman. 1998. Fight impatiens necrotic spot virus and tomato spotted wilt virus using a new monitoring system to make informed decisions. GrowerTalks, (In Press, February Issue).

Van der Schaaf, D.A., W.A. Ravensberg & M. Malais. 1991. Verticillium lecanji as a microbial insecticide against whitefly, pp. 120-123. In Proc., 3rd European Meeting on Microbial Control of Pests, IOBC Working Group on Insect Pathogens and Insect Parasitic Nematodes, 24-27 Feb.1991. Wageningen, Netherlands.

 

15

Budget For 1998/99
Expenses Category UCD UCD Texas A&M UVT
(Parrella) (UllmanlRobb) (Heinz)’ (Brownbridge)
Personnel² 45,000 30,000 25,000 14,000
Lab Supplies 4,000 5,000 8,000 8,000
Travel 1,000 2,000 2,000 2,000

 

Total Project Cost = $146,000

 

Note: The section on mechanical releases of natural enemies (UCD.Giles) will not be intiated this year and is not included. In addition, the section on Postharvest (Mitcham & Hara) will be funded by Yoder Brothers this year. Next year’s request will include budgets for


these two subprojects.

¹Dr. Heinz has a continuing project on aphids funded by the AFE. As a result, he will only require $15,000 in ‘new’ support to get this project off the ground. Full funding (as in the Table above) will be required next year.

²Detailed summaries of individual budgets have been completed. However, the majority of funding goes to support technical help in the form of students and research techmcians.

 

Leader Qualifications

 

All the PIs in this team proposal are well known to the Endow:nent-most have gotten financial support previously and many have continuing grants from the AFE. The PIs make a concerted effort to publish research in scientific journals and all are regular contributors of grower-oriented articles in national and statewide trade publications. In addtition, all the Pis give presentations to growers on a regular basis. Beth Mitcham and Arnold Hara are new to the AFE. Dr. Mitcham is Extension Specialist in the Department of Pomology at UC Davis where she specializes in alternatives to postharvest chemicals for control of insects, decay and physiological disorders and fruit responses to postharvest handling systems. She is currently chair of the University of California Postharvest Working Group. Arnold Hara is a Professor in the Department of Entomology at the Univ. of Hawaii (Hilo) where he concentrates on postharvest insect control and regulatory Entomology. Dr. Hara is the leading authority on pcstharvest insect control in the US. Drs. Mitcham and Hara have worked together previously on postharvest insect control on flowers.

 

16

This entry was posted on Sunday, June 21st, 1998 at 9:29 am and is filed under Floriculture Pest Management, Research Proposals. You can follow any responses to this entry through the RSS 2.0 feed. You can leave a response, or trackback from your own site.