Combining the Use of Mini Parasitoid Insectaries and Predator Power to Control Pecan Nut Casebearer

Project Coordinator

J. Joe Ellington
New Mexico State UniversityDepartment of Entomology, Plant Pathology & Weed Science
Box 30003/Dept. 3BE
Las Cruces, New Mexico 88003-8003
505-646-3225
505-646-8087 (fax)

Project Duration: July 1, 1999 - December 31, 2000.

		Matching Funds (if any)
	Request	Non-Federal	Federal	Total
First Year Funding	19,990			19,990
Total Funding Request	39,980			39,980

Abstract

Pecan pest management in New Mexico historically consisted of intense insecticide use that led to pesticide resistance. By 1987 biological control was adopted due to resistance problems and was successful until 1993. By 1993, the pecan nut casebearer (PNC) Acrobasis nuxvorella (Nunzig) began to infest New Mexico pecans and became such a serious problem that producers started spraying their orchards with alarming frequency. Without an integrated pest management approach that successfully controls PNC with little or no pesticide use, the risk that Mesilla Vally pecan growers will return to frequent pesticide application is very high.

Our objective is to develop mini-insectaries for parasitic Hymenoptera, Trichogramma pretiosum (Riley), and complete an action index based on naturally occurring predators. The mini-insectaries are designed to hang from trees in orchards and provide a continuous culture of parasitoids that will sting PNC eggs, and thus prevent destructive PNC larvae from emerging to damage pecan nutlets. The action threshold, or predator power index, was initially developed in 1998 at NMSU. The index predicts percent mortality of PNC eggs given predator density. The index will indicate if predator densities are too low to control PNC or that densities will sufficiently control PNC. Both controlling PNC in the egg stage to decrease the population and predicting the control power of predators will help prevent the unnecessary use of pesticides. Reducing pesticide use will prevent pest resistance, preserve the naturally diverse ecology that is already present in pecan orchards, and will be less expensive than spraying. These techniques, when tested and verified, will be communicated to growers through seminars, cooperative extension, durable brochures, and via the Internet. If the entire valley can be convinced to adopt these measures of control, the risk of heavy pesticide load in the valley will decrease dramatically.

Objectives

Determine the feasibility of field rearing parasitoids in mini-insectaries to control pecan nut casebearer in pecans.
Finalize the development of an action threshold for natural predator power that indicates: a) there is sufficient predator density to control pecan nut casebearer; b) additional control is necessary to boost the predator population.
Communicate the results of objectives 1 & 2 to growers in the form of seminars, printed materials, and a web site devoted to addressing IPM in pecans.

Justification

Objectives 1 and 2

Pecan trees in New Mexico have been attacked typically by three species of aphids (Homoptera:Aphididae) and, as of 1993, one species of Lepidoptera. The aphid complex (AC) includes the yellow pecan aphid Moneliopsis pecanis (Bissel), the black-margined pecan aphid Monellia caryella (Fitch), and the black pecan aphid Melanocallis caryaefoliae (Davis). The lepidopteran pecan nut casebearer Acrobasis nuxvorella (Neunzig) (PNC) attacks nutlets and reduces overall yield.

From the late 1940’s to the late 1970’s, the number and rates of insecticide applications in pecans steadily increased until the pecan aphid complex became resistant to insecticides. By 1986, six different insecticides (Pydrin®, Ammo®, Disyston®, Cymbush®, Thiodan®, and Lorsban®) were used with little effect on aphids but costing a total of $190.00 per acre - excluding the cost of environmental pollution. By 1987, Stahmann Farms (the states largest grower) adopted a biological control program that relied on improved cultural practices, ecological diversification, predator releases, and sampling pest and predator densities to effectively time and reduce insecticide inputs. Until 1993, spray applications and beneficial release rates decreased while yield and quality remained high (LaRock and Ellington 1996).

Since 1993, PNC has begun infesting pecans in the Mesilla Valley. As a result, insecticide applications have increased and now threaten to return pecan insect control to an intense insecticide regime. A system, such as mini-insectaries, to rear parasitoids directly in the orchards could reduce the risk that growers return to an intense PNC insecticide control regime. In-field mini-insectaries could be inexpensive, easily maintained, and could provide parasitoid stock superior to lab reared stock in that they are subject to natural selection in the environment in which they will be used. Estimated season long cost using this technique is approximately $12/acre at a rate of 150,000 T. pretiosum per acre, which is much less than conventional insecticide. Efficient in-field mini insectaries could effectively stabilize T. pretiosum populations and control insect pests in the field at a lower cost than mass releases.

To encourage producers to refrain from returning to intense insecticide use to control PNC, an initial year of work has been done to determine the density of predators needed to keep PNC populations at acceptable levels. Another year of data will provide enough information to develop a series of action thresholds where measurement of the predator/pest density in a given tree or orchard block will provide an estimate of pest mortality. Action can then be taken to induce higher mortality when needed.

Objective 3

Because a few other large growers and many medium to small growers in the valley use pesticides to control PNC in their orchards, communicating the results obtained from objective one to these growers could exponentially reduce the risk and quantity of pesticide use. Seminars will be used to train extension specialists and other interested parties in ways to set up, maintain, and evaluate the effectiveness of mini-insectaries. Print media will provide each grower with a durable brochure that explains the use of parasitic wasps and the construction and use of mini-insectaries. A web site with the same information will provide information quickly and easily, and will also provide growers with a means to e-mail the project coordinator with specific questions, concerns, or improvements to the system.

Literature Review

Pecan (Carya illinoinensis Wangenheim C. Koch) was the sixth leading agricultural commodity in New Mexico in 1991. Production was 29,000,000 pounds of nuts making New Mexico the third leading pecan-producing state in the United States following Georgia and Texas.

Parasitoids

The pecan is a native to the U. S. and unlike many introduced horticultural crops, pecan insects consist almost entirely of species that have evolved with the pecan. Twenty-six species of primary native egg and larval parasitoids were reared from PNC in Texas (Nickels et al. 1950). Gunasena & Harris (1988) found PNC larvae and pupae of the first summer generation, in Texas, were parasitized by at least 24 species belonging to 10 families in two orders. Total observed parasitism of first-summer generation PNC by all parasitoids collectively ranged from 13.6% to 47.1% in 1983 and 1984. A characteristic of this system appears to be lack of a dominant parasitoid. The complex is made up largely of non-host specific hymenopterous larval parasitoids. The host range may vary from 1-79. Effective parasitism results from low levels of parasitism by numerous parasitoids.

Because many PNC parasitoids are non-host specific, arboreal species, New Mexico may experience more PNC damage than Georgia. However, if hyperparasites are eliminated and non-sprayed fields chosen for release, native parasitoids from Georgia and Texas may establish and provide good PNC control. Habitat management may prove to be essential to establish exotic parasitoids. In an attempt to biologically control PNC, Calliephialtes graphlithae (Cress.), Phaneratoma fascieata (Prov), Lixophaga mediocris (Aldrich), Macrocentrus instabilis (Musebeck) and Agathis acrobasidis (Cushman) were imported from W.L. Tedders Southwestern, Fruit Tree Nut Laboratory, Agricultural Research Service, Bryan, Georgia in 1995.

Some 500,000,000 Trichogramma pretiosum (Riley) from a commercial source were released in the Stahmann orchard for PNC control in 1997. Approximately 39% of sampled PNC eggs from this release were parasitized by T. pretiosum.

Waterson and Stone (1982) reported that between 6-52% of black margined pecan aphids were parasitized by a native parasitoid, Aphelinus perpallidus (Gahn) in six orchards in the El Paso Valley (30 miles south of the Mesilla Valley) in 1980. Although A. perpallidus is always present it does not provide consistently high levels of aphid mortality.

Trioxys pallidus (Haliday) is an introduced aphid parasitoid from the Near East which has effectively controlled walnut aphids in California and filbert aphids in Oregon. Releases were made against pecan aphids in Georgia resulting in establishment but not control (Tedders, personal communication). Heavy insecticide spraying evidently reduced populations of T. pallidus in Georgia.

Trioxys pallidus was released in New Mexico on 2/4/88, 3/10/89, 3/20/97 and 3/25/98 from collections made at U.C. Berkeley by K. Hagen. T. pallidus has yet to be recovered. Because AC cannot be artificially reared, T. pallidus needs to be cultured in the greenhouse on pecan seedlings to observe behavior and parasitization rates before release.

Host and Parasitoid Rearing

Although no work has been done, that we know of, on rearing parasitoids directly in the field as proposed in this study, there is a great deal of information on mass rearing parasitoids in laboratory environments, from which the mini-insectaries will be inoculated.

Currently at NMSU, T. pretiosum is being mass reared following the techniques described by Morrison (1985). Using S. cerealella eggs and 9 special rearing boxes, we can produce approximately 9,000,000 parasitized eggs at a rate of about 1,000,000 per box per day. Using cold storage manipulation techniques describe by Stinner, Ridgway and Kinzer (1974) allows parasitized eggs to be stored for up to 8 days allowing accumulation for mass release on a weekly basis.

Predators

The most formidable PNC predators in New Mexico are the lacewings Chrysoperla carneae (Stevens), Chrysoperla rufilabris (Burmeister), Chrysopa nigricornis (Burmeister), the lady beetles Hippodamia convergens (Guerin-Meneville) and the arboreal Olla v-nigrum (Mulsant), and a mirid Deraeocoris nebulosus (Uhler). Natural pecan nut casebearer egg mortality currently runs from 40-90%. (Ellington et. al. 1998).

The predacious mirid Deraeocoris nebulosus (Uhler) resides around developing nutlets and in residual husks. D. nebulosus occurs in high numbers (up to 12,000/tree) and they have been observed preying on PNC and AC. D. nebulosus may be one of the best predators in pecan orchards in New Mexico. (Carrillo and Ellington 1997).

Enhancing and Estimating Predator Power

Several researchers (Ehler and van den Bosch 1974, Hagler et al. 1992, Sherratt and Harvey 1993, van Alphen and Jervis 1996) considered predators to be polyphagous because they feed on many food items and show little or no taxonomic affinity regarding prey species attacked. However, certain predators may prefer one species to another. Fye (1979), in laboratory studies, showed that food preference by cotton predators varied with development stage and prey choice. Food choices may tend to be site specific, depending on environmental factors and species availability. However, it is unlikely that predators differentiate between primary consumers and other predators as food. According to Rosenheim et al. (1995), predator-predator interactions may limit control of a primary consumer through intraguild predation, which consists of competition for a food resource and a mutual trophic interaction between two predators. These authors cite many examples of disrupted biological control programs due, in large measure, to intraguild predation. Ellington et al. (1997) found predators in cotton feed widely on both primary consumers and other predators. They found predators were associated with primary consumers 46% of the time and with other predators 53% of the time. Since quantifying specific predator food preferences under different site-specific environmental conditions and arthropod host densities is a very expensive and labor intensive process, it may be possible to sum predator density and use it as an aid in predicting pest mortality to establish economic thresholds. Generalists or polyphagous predators could exert a degree of control consistent and more complete than that achieved by a specialist; and mortality caused by generalists could be density independent (Huffaker et al. 1969).

Multiple and logistic regression has been used to predict mortality as a function of environmental and pathogenic factors. Armstrong and Peairs in 1996 used multiple regression to examine the relationship between the winter environment and mortality of the Russian wheat aphid. Logistic regression was used in 1991 by Woods et al. to examine viral transmission dynamics and predict mortality in gypsy moths. In Woods’ study, it was determined that logistic models might not accurately predict mortality above 90%. Ellington et al. (1998 a and b) in a one year pre-study used logistic regression to predict PNC egg mortality (simulated by gluing bollworm eggs on pecan leaves) as a function of predator density. Models were fit based on geographic blocks, time, length of time eggs were exposed to predators, and predator density. Few instances of mortality greater than 90% occurred, so logistic regression, in this case, is appropriate for predicting mortality.

Approach and Methods

In-field Mini Parasitoid Insectaries

Before starting the Sitatroga cerealella/T. pretiosum mini-insectaries, wheat will be bioassayed to detect insecticide contamination by exposing the wheat to adult S. cerealella in pint jars for seven days (Morrison 1985). All wheat and production equipment will be exposed to heat treating at 66° C for 24 hours before use. Then the wheat will be soaked in water to restore moisture content to ca. 15%.

S. cerealella eggs will be decontaminated in a 10% formalin solution for 10 minutes and added to the soaked grain in bulk ,on a weekly basis, for 4 weeks to reduce generation cycling. One month later S. cerealella eggs, parasitized by T. pretiosum, will be added for 4 weeks, on a weekly basis, to reduce generation cycling and to establish the T. pretiosum culture.

Mini-insectary containers will be new friction topped tin cans (i.e. paint cans) with 20, 1/16” in diameter holes drilled into the sides to allow T. pretiosum to exit the containers. Tin will be used so that the inside of the mini-insectary is completely dark for maximum contrast with the exit holes to attain maximum parasitoid attraction. The mini-insectaries will have wire handles for easy hanging in trees.

Mini-insectaries will be inoculated from the lab culture. S. cerealella and T. pretiosum cultures will be removed from the bulk container and placed into cheesecloth to fill one half of each of the mini-insectary containers. The wheat will have been infested at a rate of 1 larva per two wheat kernels. Soaked hard winter wheat with a protein content of 13 - 15% will top off the containers to provide a food supply over the season for the moth larvae while a 2 in space will be left at the top for adult moths to breed. Sample containers will be observed weekly to check for contamination by mites and/or mold. Containers will be checked monthly to track grain depletion and for refilling if necessary. In the case of mite contamination, each mini-insectary will be provided with a strip of Apistan to control mites.

Mini-insectaries will be hung from pecan trees before first PNC oviposition, around May 15th, at the rates of 0, 1, 20, and 50 per tree per acre. Average tree planting rate is approximately 50 per acre. There will be 5, 1 acre replications per treatment in a randomized complete block design. Ten PNC eggs from each replication will be removed weekly to the laboratory and checked for parasitization during each PNC generation. Percent larval infestation will be documented in early, mid, and late season by counting 30 nutlets in 10 trees per replication. One PNC pheromone trap will be placed in one replication of each treatment and checked weekly.

Ten mini-insectaries will be set up in the laboratory in clear plastic containers. Emergence of T. pretiosum will be counted weekly through the summer.

Predator Power Sampling and Estimation

Four trees per replication, for a total of 20 trees, will be randomly selected twice per month and 20 PNC eggs flagged per replication and observed for a two week period to determine predation rates. Trees will then be fogged with a broad-spectrum insecticide (Kicker) (Ellington et al. 1999b). Insects that fall from the trees will be collected on plastic sheets placed beneath the tree. Temperature and humidity loggers will be placed on each randomly selected tree at the beginning of the observation period.

A database of 20 measures of PNC predator density taken 14 times over the season (twice per month) will provide 20 x 7 = 140 observations per year. Time and year will be included in model estimation and blocked on at least a monthly basis. Time may be blocked in even larger units if the lifecycles of PNC or predators indicates that larger blocks can be used, which will allow for more degrees of freedom in model estimation. PNC egg mortality/fecundity will be measured by counting the number of PNC eggs in the beginning of the two week period and counting them daily to see if they are eaten by a predator, parasitized, or emerged. Logistic regression will be used to fit a model such that mortality/fecundity is a function of predator density, parasite density, alternate food source density (aphids and other species’ eggs) time (seasonal), time (annual), temperature, humidity, and geographic blocking factors. An index will be created based on PNC mortality/fecundity and the predators present at a given period in the PNC lifecycle.

Impact Assessment

Impact assessment will be carried out by comparing experimental plots to non-experimental plots within the Stahmann Farms orchard and to the average situation encountered by other growers in the valley. Yields/acre, pesticide costs/acre, amount of pesticide applied per acre, and management costs per acre will be collected over the life of the project. Non-experimental costs/acre will be compared with experimental costs/acre as well as experimental and non-experimental yields/acre. Beneficial insects will be sampled in experimental and non-experimental plots and compared in terms of predator diversity and density to quantify how well the program preserves the natural pecan orchard ecology. Multivariate analysis will be used to generalize characteristics between experimental plots and non-experimental plots on the same farm as well as with yields and costs taken from other farms.

Appendix A. Literature Cited

Armstrong, J. S., F. B. Peairs, 1996, Environmental parameters related to winter mortality of the Russian wheat aphid (Homoptera: Aphididae): basis forpredicting mortality. Jour. Econ. Entomol. 89: 5, 1281-1287.

Carrillo, T., and J. Ellington. 1997. Recent developments in biological control of pecan insects. Fourteenth western pecan conference proceedings.

Ehler, L. E., and R. van den Bosch. 1974. An analysis of the natural biological control of Trichoplusia in (Lepidoptera: Noctuidae) in cotton and alfalfa. Can. Entomol. 106: 1067 - 1073.

Ellington, J., T. Carillo and M. Southward. 1997. Association among cotton arthropods. Env. Entomol. 26: 1004 - 1008.

Ellington, J., D. Richman, S. Meeks, T. Carillo, S. Liesner, and S. Ball. Sept. 1998a. Biological Control of Insect Pests in Pecans. Presented at the Arizona Pecan Growers Association Meeting. Tucson, Az.

Ellington, J., D. Richman, S. Meeks, T. Carillo, S. Liesner, and S. Ball. Sept. 1998b. Biological Control of Insect Pests in Pecans. Annual Research Report. New Mexico State University. Department of Entomology, Plant Pathology & Weed Science.

Fye, R. E. 1979. Cotton insect populations. U. S. Dept. Agric. Tech. Bull. 1592.

Gunasena, G. H. And M. K. Harris. 1988. Parasites of hickory shuckworm and pecan nut casebearer with five new host-parasite records. S.W. Entomol. 28:291-338.

Hagler, J. R., A. C. Cohen, D. Bradley Dunlop, and F. J. Enriquez. 1992. Field evaluation of predation on Lygus hesperus (Hemiptera: Miridae) using a species and stage specific monoclonal antibody. Environ. Entomol. 21: 896 - 900.

Huffaker, C. B., M. van de Vrie, and J. A. McMurtry. 1969. The ecololgy of tetranychid mites and their natural control. Annu. Rev. Entomol. 14: 125 - 174.

LaRock, D.R. and J.J. Ellington. 1996. An integrated pest management approach, emphasizing biological control, for pecan aphids. Southwest. Entomol. 21: 153-166.

Morrison, R. 1985. Mass Production of Trichogramma pretiosum (Riley). The Southwestern Entomologist. Suppl. No. 8.

Nickles, C.B., W.C. Pierce and C.C. Pinkney, 1950. Parasites of the pecan nut casebearer in Texas. U.S. Dept. Agric. Tech. Bull. 1011, 21pp.

Rosenheim J. A., H. K. Kaya, L. E. Ehler, J. J. Marois, and B. A. Jaffe. 1995. Intraguild predation among biological control agents theory and evidence. Bio. Control. 5: 303 - 335.

Sherratt, T. N., and I. F. Harvey. 1993. Frequency-dependent food selection by arthropods: a review. Bio. J. Linn. Soc. 48: 167 - 86.

Stinner, R. E., R. L. Ridgeway, and R. E. Kinzer. 1974. Storage, manipulation of emergence, and estimation of numbers of Trichogramma pretiosum for field release. Southwest. Entomol. 3:62-8.

Tedders, W.L. 1983. Insect management in deciduous orchard ecosystems: habitat manipulation. Environ. Manag. 7:29-34.

Tedders, W. L. 1995. Personal Communication.

Van Alphen and M. S. Jevis. 1996. Foraging behavior, pp. 1 - 62. In M. J. Jervis and N. Kidd [eds.], Insect natural enemies. Chapman & Hall, London.

Waterson, G.P. and J.D. Stone. 1982. Parasites of blackmargined aphids and their effect on aphid populations in far-West Texas. Enviro. Entomol. 11:667-674.

Woods, S. A., J. S. Elkinton, K. D. Murray, A. M. Liebhold, J. R. Gould, J. D. Podgwaite. 1991. Transmission dynamics of a nuclear polyhedrosis virus and predicting mortality in gypsy moth (Lepidoptera: Lymantriidae) populations. Jour. Econ. Entomol. 84:2, 423 - 430.

Appendix B. Timetable

Could not be reproduced for inclusion in this posting.

Appendix C. Major Participants

J. Joe Ellington, PhD with New Mexico State University
Dr. Ellington will contribute 2% of his time to the project as well as laboratory resources, computing resources, and the resources of a new 3000 sq. ft. insectary at NMSU.

Stahmann Farms
Stahmann Farms, the valley’s largest grower with orchards here and in Australia, will contribute sufficient acreage to carry out this experiment. They have also provided monetary and material support for the mass rearing of T. pretiosum and Deraeocoris nebulosus.

Project Budget

Project Period: July 1, 1999 - December 31, 2000

Budget Category	Grant Funding	Other Funding	Total Funding
Personnel	27,774	29,780	57,554
Fringe Benefits	556	7,742	8,298
Travel	1,980		1,980
Equipment
Supplies	5,070		5,070
Contractual
Other (indirect costs @ 13%)	4,600		4,600
Total	39,980	37,522	77,502

Budget Justification

Personnel & Fringes - Funding is requested for one graduate student and seven work-study students with a fringe rate of 2%.

Matching funds come from 2% of Dr. Joe Ellington’s salary and fringes calculated at 26% of salary.

Travel - Travel is requested to and from the field site at $0.33/mile.

Supplies - Supply funds requested are for 20 HOBO temperature and humidity loggers, shuttle, and windows 95 software. Supply funds are also requested for S. cerealalla eggs, grain, rearing supplies and tin buckets.

Other (Indirect Cost) - NMSU has a standard negotiated rate of 43.6%. We are waiving 30.6% of this and requesting IDC at 13%.