Managing “Pests” with Fewer Insecticides: Introducing Biocontrols at Smith College Botanical Gardens Smith Botanical Garden, 1899. Photo taken by Katherine E. McClellan, Courtesy of Smith College Archives. EVS 300 Semester Project Maribeth Kniffin Smith College Spring 2007 “...biological and chemical control are considered as supplementary to one another as the two edges of the same sword... nature’s own balance provides the major part of the protection that is required for the successful pursuit of agriculture... insecticides should be used so as to interfere with natural control of pests as little as possible... .” -Hokins et al., Proc. 6th Pac. Sci. Congr. 1 This paper introduces the concept of biocontrols, their history, and their potential for use at the Smith College Botanical Garden. By performing an insecticide inventory, examining biocontrol efficiency through an experiment, assessing the risks of biocontrols, analyzing a cost-benefit analysis, and speaking with other community members involved using biocontrols, I formulated a biocontrol/insecticide application plan that will hopefully be put to use. Introducing Biocontrols Botanical gardens throughout the world use insecticides to control unwanted insects populations in order to maximize visitor comfort and appeal. Any insecticides applied to a greenhouse could potentially impact the health of the person performing the application, the greenhouse visitors, and the surrounding ecosystem (Material Safely Data Sheets). Insecticides have varying levels of toxicity and re-entry intervals, thus, proper application techniques, such as evening insecticide application, is necessary to allow for sufficient breakdown of the chemicals. Unfortunately, as history has demonstrated, seldom do we learn the true toxicity of an insecticide (and any pesticide) until after the harm has been done (Carson, 1962). For this reason, limiting the use of insecticides is to humanity’s advantage. In numerous greenhouses worldwide, insecticide application has been reduced by adopting an Integrated Pest Management Program (IPM). IPM is based on a preventative rather than a reactive ideology that integrates the use of chemical, biocontrols, physical means, observation, and education to control pest populations (Green Methods Catalog, 2004). Pests, in this sense, can be insects, mites, weeds, pathogens, and even nonarthropod animals as well (Kogan, 1998). This study focuses on insect pests and their management through the use of biocontrols. Biocontrols can reduce insecticide applications by using natural enemies to control pest populations. There are two main types of biocontrols insects: predators and 2 parasitoids (Green Methods, 2007). Predators act by consuming the pests, while parasitoids spend the bulk of their life attached to or within a target host organism, which will ultimately kill the host, in this case a pest. Both organisms are commercially available by growers worldwide (IPM Labs). For example, in the northeast, IPM Labs and The Green Spot are two well known commercial growers. Since biocontrols are based on a preventative rather than reactive ideology, using biocontrols not only calls for a change in technique, but also in the way we think about pest control in general. As stated in the Green Methods catalogue, using biocontrols necessitates four Ps: planning, prevention, patience, and perseverance (Green Methods Catalogue, 2004). Unlike insecticides, biocontrols will not kill pest insects instantly; conditions must be favorable for the biocontrol insect and adequate numbers must be released, etc. Biocontrol introduction must be planned and monitored. And when it is, it has been very successful (Fred, personal interview, Hoddle et al., 1998). History of Biocontrols The use of biocontrols, especially predation, has been readily seen through the course of history, as early as 4,000 years ago in Egypt where domestic cats were depicted as useful in rodent control (Green Methods, 2007). The 16th century Chinese released predaceous ants, Oncophylla smaradina, to control pests in citrus orchards since the ants fed on foliage-feeding insects. The Chinese attached bamboo bridges between trees to aid in ant motility (Green Methods, 2007). Similarly in 1762, in Mauritius, Mynan birds were relocated from India for locust control, which was the first successful importation of an organism from one country to another for biological control (Green Methods, 2007). 3 Insect parasitoidism was not formally recognized until the turn of the 17th Century with the first record from the Italian, Aldrovandi (1602) (Green Methods, 2007). Then in 1700, Antoni van Leeuwenhoek described the phenomenon of parasitoidism in insects. Even Erasmus Darwin in 1800 mentioned the useful role of parasitoids and predators in regulating insect pests (Desmond, 2002). In the United States, biocontrols were introduced when cottony-cushion scales decimated the citrus industry in California in 1889 (Britannica, 2007). Here, the vedalia lady beetle was introduced as a control and eventually the citrus industry recovered and thrived. Ironically, pesticide control replaced the lady beetle, which killed off the lady beetle population (Britannica, 2007). Once again, the cottony-cushion scale is in need of control. The 1940s to mid-1960s are known as the “Dark Ages of Pest control” since biocontrols rapidly and drastically decreased with the introduction of pesticides (van Lenteren and Woets, 1988). Despite the trend, biocontrols retained a few steady supporters, notably Rachael Carson and Everett J. “Deke” Dietrick (Carson, 1962). With the increasing pesticide awareness in the late-1960s and 70s, biocontrols one again began to take hold and in 1972 Nixon introduced IPM to U.S. Congress (Council on Environmental Quality, 1972). By 1979, 75% of 51 entomology departments that were surveyed in the United States offered a course in IPM (Pieters, 1979) and in 1970, whitefly parasitoid was used on 100 hectares greenhouse crops, while in 1993, its use had increased to 4800 hectares greenhouse crops (Hoddle et. al, 1998). 4 Introduction: Biocontrol Project at Smith College Botanical Garden The Smith Botanical Garden has six main pests: whitefly, mealy bug, scales, aphids, spider mites, and thrips. Two of these pests, whiteflies and mealy bugs, tend to be the most problematic, especially in the Palm House and Show House. Whiteflies cause leaf chlorosis, leaf withering, premature dehiscence, and defoliation, and can result in a plant yield reduction of over 50% or even plant death (Byrne and Bellows, 1991). Mealy bugs cause leaf chlorosis, premature dehiscence, sticky residue, and powdery mildew (personal observation). Currently, the greenhouse pests are controlled using numerous insecticides, although there are many disadvantages in doing so (Nate Saxe, personal interview). With the constant flow of people visiting the greenhouses and necessary re-entry intervals (REI) after insecticide applications, applying insecticides is a nuisance and difficult to arrange (Steve Sojkowski, personal interview). Any insecticide applications must be carried out after hours, typically done on Fridays, and due to health hazards, applying insecticides requires state training (Steve Sojkowski, personal interview). Although trained, health risks remain for those handling insecticides as well as school groups with children who place countless objects in their mouths, including leaves from sprayed plants. Insects also build resistance to insecticides so that more chemical or a change in chemicals is needed for the desired effect. Finally, insecticide prices continually rise. For these reasons, the conservatory manager, Rob Nicholson, was interested in introducing biocontrols to the greenhouses, particularly the parasitoid wasp Encarsia formosa to control the whitefly population and the predator lady beetle Cryptolaemous montouriezi to control the mealy bug population. Rob appointed me to design and carry 5 out this project. Through the use of the two biocontrols, we hoped to reduce the number of pesticide applications. Materials and Methods My project included three main parts: an insecticide inventory, examining biocontrol efficacy, and developing an insecticide/biocontrol insect application plan. The insecticide inventory included researching the insecticides used at the Botanical Garden of Smith College within the last ten years, examining Material Data Safety Sheets for six chemical used in 2006, and comparing number of applications and amount of insecticides used in the last ten years. To examine biocontrol efficacy, I conducted an experiment that investigated whether E. formosa is an effective alternative to insecticides for controlling the whitefly population in the Smith Botanical Garden. This involved researching the biology of E. formosa, as well as assessing the risks of introducing a non-native species into the gardens. Finally, I devised a potential insecticide/beneficial insect application schedule for the houses that have the introduced beneficial insects by determining more exact numbers of E. formosa needed for future releases rates, gathering information from neighboring gardens that have used biocontrols, performing a cost-benefit analysis for biocontrols versus insecticides, and ultimately learning from my experiment. Part I. Pesticide Inventory To make an inventory of the pesticides within the last five years, I talked to Rob Nicholson, the conservatory manager, and Steve Sojkowski and Nate Saxe, the greenhouse assistants. They directed me to the Pesticide Application Records and the pesticide Material Safety Data Sheets. I also researched the possible effects the pesticides have on humans and the environment through primary and secondary literature. 6 Part II. Biocontrol Efficacy Whitefly and Parasitoid Wasp Biology and Assessing the Risks of E. Formosa To determine the whitefly and parasitoid wasp biology and assess the risks of introducing E. formosa into Smith College Botanical Garden, I conducted research using primary literature along with commercial and governmental websites. Experiment To conduct the beneficial insect experiment, on March 16th I released 10,000 black parasitized scales of E. formosa ordered from IPM Labs into two houses within the Smith Botanical Garden: 7,000 scales in the palm house and 3,000 scales in the show house. The warm temperate house was the control and received no biocontrol insects. Scales were attached to cards (100 scales/card), which I dispersed evenly throughout the greenhouses, hanging them on branches of whitefly infested plants. Each scale on the release card contained a developing parasite that emerged within 8-10 days to fly throughout the greenhouse searching for whiteflies. Throughout the duration of the 5 week experiment, I assessed the whitefly population in two ways: 1) by counting the number of larvae on leaves and; 2) by counting the number of whiteflies on yellow sticky cards. I selected four plants per house that contained whitefly larvae on the undersides of their leaves. The four plants I chose to observe in the Show House were Centranthenum punctam subspecies camporum in the Asteraceae family, Eranthemann pulchellum in the Acanthaceae family, Cananga fruticosa in the Bignoniaceae family, and Mitrephora wangii in the Annonaceae family. The four plants observed in the Palm House were Hibiscus rosa-sinensis in the Malvaceae family, Acalypha hispida in the Euphorbiaceae family, Tibouchina 7 grandifolia in the Melastomataceae family, and Brunfelsia lactea in the Solancaceae family. Finally, the four plants observed in the Warm Temperature House were Senecio petatis in the Asteraceae family, Malvaviscus arboreus in the Malvaceae family, Monolena primuliflora in the Melastomaceae family, and Gossypium barbadense in the Malvaceae family. From March 16th through April 25th, I counted the number of larvae on 3 leaves per plant once a week, scouting for black parasitized scales and wasps. Near two selected plants in each room, I also placed a yellow sticky card (2x3”). Whitefly counts were taken each week before the traps were changed. Part III. Biocontrol/Insecticide Application Plan Future E. formosa Release Rates To assist in formulating a biocontrol/insecticide application plan, I measured dimensions and counted number of plants by hand in each room to help determine exact numbers needed for future E. formosa release rates. Community Interactions I gathered information from neighboring gardens by discussing biocontrols with members in the community, namely Fred from the Deerfield Magic Wings Butterfly Conservatory, the new director of the UMass Botanical Garden, and Carol, the entomologist, at IPM Labs, a well-known commercial biocontrol company in the Locke, New York. Cost-benefit analysis To perform the cost-benefit analysis, I spoke with Rob Nicholson, the conservatory manager concerning the Griffin Greenhouse and Nursery Supplies Catalog, which is the company from which Smith College Botanical Garden orders their pesticides 8 and our annual costs. I researched the cost of biocontrols on the IPM Labs website along with reading primary literature on cost effectiveness of biocontols. Results Part I. Pesticide Inventory Over the last ten years, the Smith College Botanical Garden has used 17 different insecticides in a wide range of chemical families or types. The insecticides fall into one of the following eight categories: insecticidal soap, insect growth regulator, pyrethroid, orthophosphate, plant systemic, oil, cholinesterase inhibitor, or other synthetic chemical. Specifically within the last year (2006), seven different insecticides have been used. Information from the Material Data Safety Sheets on type of chemical, its targets, active ingredients, health/safety alert, effects on wildlife, oral and dermal LD50, signs and symptoms of acute exposure, and overexposure is listed below for six of the seven insecticides (the MSDS for Synergy was could not be located) (Tables 1-6). All insecticides had potential effects on humans and aquatic species. The number of pesticide applications has increased over the past five years from an average of 3-4 applications per month from 1997-2001 to 4-5 applications per month from 2002-2006 (Figure 1). Despite the increase in number of applications, the total amount of insecticides used throughout the past 10 years has roughly stayed the same. The total amount of insecticides used in 2006 can be found in Table 7. Part II. Biocontrol Efficacy Whitefly Biology Whiteflies (order Homopteran) have caused damage globally, including the United States, by affecting vegetable, herbal, and floral crops via extraction of large 9 quantities of phloem sap (Hoddle et al, 1998). Such extractions during high whitefly infestations cause symptoms of leaf chlorosis, leaf withering, premature dehiscence, and defoliation and can result in a plant yield reduction of over 50% or even plant death (Byrne and Bellows, 1991). Whiteflies also negatively impact plants by excreting honeydew, a medium that encourages sooty mold fungi growth, which makes leaves unavailable for food and fiber use. Finally, a few species of whiteflies are vectors of viral plant pathogens, namely, the lettuce infectious yellow virus (Byrne and Bellow, 1991). Whiteflies are the tropical equivalent of aphids, not normally found in temperature climates outside of greenhouses. Their exact origin is in speculation, although, Mound and Halsey noted that most species of Trialeurodes (described below) are found in the New World. They mainly attack species in the families Cruciferae, Leguminosae, Malvaceae, and Solanaceae, but may attack more plant species in greenhouse environments (Hoddle et al., 1998). Whitefly studies in natural settings are lacking. In the United States, two whitefly species are well-known pests in greenhouses: Trialeurodes vaporariorum, the greenhouse whitefly, and Bemisia tabici or B. argentifolii, the sweet potato or silver leaf whitefly, respectively. We initially identified T. vaporariorum in the Botanical Garden of Smith College, and thus, will focus on this species. T. vaporariorum is a small white fly (as the common name describes), approximately 1 mm in length, with four membranous wings that span to roughly 2.5 mm (Figure 2). All life stages, except the egg, produce a colorless or white extracuticular wax primarily made of triacylglycerols (Byrne and Bellows, 1991). T. vaporariorum 10 feeds on plants with its sticky sucking mouth parts, is opisthognathus, and undergoes incomplete metamorphosis (Van Lenteren and Woet, 1988). The life cycle of T. vaporariorum undergoes four main stages: the egg, the larvae stage (1st instar, 2nd instar, and 3rd instar), the pupa (4th instar), and adult whitefly (Beyrn and bellows, 1991) (Figure 3). The larvae and pupa stage are similar in appearance; both look like small, white scales on the underside of leaves (Figure 2). A typical T. vaporariorum life cycle is 15 days under the developmental threshold temperature for T. vaporariorum of ~10ºC, although it will multiply faster during warmer weather (Byrne and Bellows, 1991). T. vaporariorum adult females lay white eggs on the underside of new plant leaves. If the leaf is glabrous, the eggs will appear in a circular pattern, while no pattern appears on leaves with pubescence. The white eggs will turn into larvae and darken slightly when they are close to hatching. Since T. vaporariorum adult females lay their eggs on new growth towards the top of a plant, older stages or instars are found on lower leaves as a plant grows. When looking at a plant infested with whiteflies, the top leaves will have eggs and adults, the middle leaves will have larvae, and the bottom leaves will have pupae (Fred, personal communication). The progression of instars from the top of the plant to the bottom can help determine the age of infestation. Parasitoid Wasp Biology E. formosa (Hymenoptera: Aphelinidae) is a parasitoid wasp used worldwide as a biocontrol of greenhouse whitefly populations on indoor vegetables and ornamental plants (van Lenteren et al., 1996). Use of E. formosa as a biocontrol was first introduced 11 in Europe in the late 1920’s (Hoddle et al., 1998). Shortly after, interest in biocontrols, such as E. formosa, declined with the widespread availability of pesticides. By the 1970s, interest resumed and in 1993, E. formosa was used on 4,800 hectares of greenhouse crops (Hoddle et al., 1998, van Lenteren and Woets, 1988). The native habitat of E. formosa is unknown, although the parasitoid is speculated to be of tropical and Western hemisphere origin. E. formosa is widely distributed in greenhouses, especially in Europe and Russia, and has successfully been used to control the whitefly Trialeurodes vaporariorum on tomato (Lycopersicon esculentum), cucumber (Cucumis sativus), eggplant (Solanum melongena var. esculenta), gerbera (Gerbera jamesonii), poinsettia (Euphorbia pulcherrima), marigolds (Tagetes erecta), and strawberry (Fragaria X ananassa) (Hoddle et al., 1998). Identifying the correct whitefly species is critical prior to parasitoid use since E. formosa is quite effective for the prevention and low-infestation management of T. vaporariorum, the greenhouse whitefly, but much less effective on B. tabici and B. argentifolii. E. formosa cannot control sweet potato whitefly unless high numbers are released. The best way to distinguish between the species is to examine the whitefly’s pupae using a powerful microscope. Pupae of T. vaporariorum contain a distinctive ridge with sides at a 90º angle to the leaf’s surface and are fringed with long hairs, while, pupae of B. tabici and B. argentifolii have a dome profile and lack long hairs. E. Formosa are the size of a pencil point, approximately ~0.6 mm in length, which is much smaller than T. vaporariorum (Green Methods, 2007) (Figure 4). Females are abundant and have a black head and thorax with a yellow abdomen, while males are rare and dark in color (van Lenteren and Woets, 1988; personal observation). 12 The E. formosa parasitoids control the T. vaporariorum population by parasitizing or consuming whitefly larvae. Adult females lay eggs inside the immature stages of whiteflies, preferring two-week old scales, i. e. the fourth instar (van Lenteren et al., 1996). As the young parasitic wasp develops inside the scale, the scale darkens until it is black, a process which lasts approximately 10 days (van Lenteren et al., 1996). Another 10 days after the scale turns black, the adult wasp chews a round exit hole on the dorsal surface of the fourth instar pupa and emerges (Green Methods, 2007). Since adult females lay an average of 5 eggs per day and have a twelve day life expectancy, an adult female will oviposit approximately 60 eggs. Daily egg maturation and ovipotition rates decline as wasps age (Hoddle et al., 1998). The complete life cycle lasts about 28 days in commercial greenhouses (Hoddle et al., 1998). Optimal temperatures are near 23ºC. Although E. formosa mainly feeds on the honeydew secreted by the whiteflies, a small number of whitefly scales are also directly eaten by adult wasps (van Lenteren et al., 1996). Adult females host-feed on immature stages of whiteflies by puncturing the body with their ovipositors and consuming hemolymph (Green Methods, 2007). Wasps may enlarge and feed from the wounds for up to six minutes (Green Methods, 2007); wasps will feed on three nymphs per day; thus, including both parasiziting and feeding on the host larvae, an average of 96 whitefly nymph per wasp are killed (Hoddle et al., 1998). The rate at which hosts are encountered is dependent on the parasitoids' walking speed, whitefly size, and number of hosts on a leaf (Hoddle et al., 1998). Walking speed is decreased by leaf venation, high trichome densities, high amounts of honeydew, the number of encounters with nymphs suitable for host feeding and parasitism, decreasing temperature, low barometric pressure, and smaller egg loads (Hoddle et al. 1998). 13 Establishing the E. formosa population early in the season before whiteflies get out of control is important for parasitoid success (Green Methods Catalogue, 2007). When populations are established, the use of E. formosa can lead to a safer means of pest management, be economically advantageous, and involve less labor. Assessing the Risks of E. formosa After investigation, minimal risks were associated with introducing E. formosa into the Smith College Botanical Garden. As van Lenteren and Loomans (1988) state biocontrol with E. Formosa “has operated effectively over decades without any establishment outside of greenhouses… or any deleterious affects on native fauna.” E. formosa is one of the forty-seven beneficial insects that have been certified for importation by USDA-APHIS standards. Experiment Throughout the 5 week period, I observed no significant change in the whitefly scale population (Figure 5), nor did I find any black scale upon the leaves. Part III. Biocontrol/Insecticide Application Plan Future E. formosa Release Rates The room dimensions and plant counts per room are as follows: the Show House is 541.5 ft2 and has approximately 200 plants, the Palm house is about 940 ft2 and has about 1000 plants, and the Warm Temperate House is 788.5 ft2 with roughly 435 plants. Community Interactions Fred from the Deerfield Magic Wings Butterfly Conservatory was very helpful and showed me all of the biocontrols that they use, while discussing appropriate techniques. He has had success using E. formosa on his greenhouse whitefly population, 14 especially on Lantana. When the whitefly population increases too much, he alternates applications of biocontrols and Enstar, an insect growth regulator. For him, whitefly populations increase in fall and spring (similar to Smith College Botanical Gardens). Magic Wings spends approximately $1,000 on biocontrols per year. The UMass Botanical Garden recently released biocontrols without extensive planning; the biocontrols were a “gift” from an entomology class that was finished with using them. As of yet, they have no results. I also spoke with Carol, the entomologist from a well-known commercial biocontrol company in the Locke, New York called IPM Labs. She explained that most IPM customers are in agriculture (monocropping) or floriculture. Many IPM customers have had success with biocontrols, specifically E. formosa, including Karen McNaughton at Mahoneys garden center in the Boston, although biocontrols are much less straight forward than using insecticides. Since biocontrol use is different in botanical gardens than in other settings because there is never a time when you can sterile your greenhouse and plants from pests to start anew with biocontrols and low numbers of pests. She recommended emailing Jody Fetzer at the New York Botanical Garden to discuss biocontrol use in botanical gardens, as Fetzer heavily uses biocontrols. She also articulated how important it is to reduce the numbers of pests before introducing the beneficials and recommended releasing low populations (1/ft2) multiple times, possibly alternating with an insect growth regulators. Cost Benefit Analysis The pesticide yearly expenditure at Smith College Botanical Gardens is roughly $600 (Rob Nicholson, personal interview). This sum does not include the uncalculated 15 cost of safety for workers, school groups, and aquatic and other species in the surrounding ecosystem. E. formosa costs $16.50 for 2000 parasitized scales, which would be sufficient for 1 release in the both the Palm and Show House. If we average 10 releases per year, plus a shipping and handing cost of $50; the total cost is $210/year for E. formosa. Past studies indicate that he cost of E. formosa is often more costly, but can be comparable to foliar pesticides (Hoddle, 1998; Van Driesche et al., 2002). The calculated cost difference between single application of systemic insecticide imidaploprid was six cents (Van Driesche et al., 2002). Discussion Part I. Insecticide Inventory As listed in the insecticide inventory, there are potential health risks involved in using the types of insecticides currently available at the Smith College Botanical Garden. For humans, many of acute risks involve eye and skin irritation; all of them are of danger to aquatic species. When looking at figure 4, there appears to be an increase in number of pesticide applications within the last five years. After consulting the greenhouse assistants, they explained that this is a result of frequent (but very low volume) Scythe applications under benches to control weeds. Thus, it did not involve a significant increase in total volume of yearly pesticides applied. Part II. Beneficial Insect Experiment We did not see any parasitism after the first introduction of E. formosa in the two greenhouses (remember four Ps). As Carol from IPM Labs explained, this is a common 16 occurrence because there are many factors involved. First of all, the wasps must be synchronized with whitefly population. Thus, it is important to perform one weekly introduction of E. formosa over the course of 4-5 weeks. Another factor is insecticide application history. Some chemicals may deposit residuals on the leaves that can last from weeks to months. Also, honeydew excretions from whiteflies can inhibit motility of E. formosa; rinsing the leaves before parasitoid releases may be helpful. Most importantly, amidst the experiment, I identified the whitefly species on Hibiscus in the Palm House to be a species other than T. vaporariorum. Unfortunately, E. formosa is not as effective on other whitefly species. I sent a Hibiscus leaf to Carol, the entomologist at IPM labs, and she identified the whitefly species as Bemisia tabici or B. argentifolii, the sweet potato or silver leaf whitefly. Also, the same Hibiscus plant has too many whiteflies for E. formosa control. Decreasing the whitefly population before biocontrol release would aide in successful management. Using insecticides that are least detrimental to the biocontrols is important. Possible insecticide combinations with E. formosa include insecticidal soap, buprofezin, azadirachtin, abamectin, and resmethrin (Hoddle et al., 1998). Part III. Pesticide/Insect Application Schedule Cost-Benefit Analysis The cost benefit analysis is understandably incomplete. There are too many factors to weigh including other biocontrols we would need to introduce, which rooms we would introduce them to, etc. This report could have solely examined the economic implications of biocontrols. 17 Integration of Research In both houses it is first important to reduce population on target plants; thus, we should apply an insecticide that we already have to hot spots. Enstar, the insect growth regulator, or Marathon, the systemic are known to be least detrimental to the biocontrols. After a few days, we should rinse off the leaves to reduce the amount honeydue excretions from whiteflies and release a parasitoid mixture: Eretmocerus eremicus (which targets B. tabici and B. argentifolii) and E. formosa. Parasitoids should be released at 1 order of 2,000 scales weekly, for 4 weeks and plants should be monitored regularly. Hopefully, after this routine, we could establish E. eremicus and E. formosa populations. Throughout the year, bimonthly applications of Enstar or Marathon could be performed to sustain low level populations of whitefly, while affecting as few a number of biocontrols as possible. In general, reducing the stresses on plants will help decrease whitefly populations. Cutting the roots of some plants or putting them in large pots may reduce stress. There is also a large population of spider mites; using a biocontrol for spider mites is possible and has been researched to be effective, which will also reduce the stress. Conclusion Biocontrols have been known to be effective throughout history. They are a possible alternative to insecticides; although, they are not straight forward. Biocontrols require patients, observation, education, and perseverance. Past pest management at Smith College Botanical Garden has been solely through use of insecticides. Currently, we are in the midst of trying to incorporate biocontrols as a viable alternative since it has no known ecosystem threats and is economically feasible. 18 Establishing biocontrols will require planning, patience, and a change in thinking, but will help us cultivate awareness of the affects we have on our community, environment, and Earth. Ultimately, if the Smith College Botanical Garden and other botanical gardens incorporated beneficial insects into the pest control scheme, the demand for pesticides could decrease on a larger scale and fewer pesticides would need to be transported and made. The Smith College Botanical Garden could be a model institution. 19 Number of Applications Tables and Figures 100 90 80 70 60 50 40 30 20 10 0 1997 1998 1999 2000 2001 2002 2003 2004 2005 2006 Figure 1. Total Number of Pesticide Applications, 1997-2006. Figure 2. Left: adult whitefly; center: adult whiteflies on leaf; right: Small white scales on the underside of Hibiscus leaves. 10 10 Figure 3. Diagram of Whitefly (T. vaporariorum)and Parasitoid Wasp (E. formosa) lifecycle. http://www.buglogical.com/encarcia_control_whiteFly/encarsia.asp 20 Figure 4. E. Left: E. Formosa parasitizing whitefly scale; right; whitefly scales: black is parasitized, white is not parasitized. http://www.buglogical.com/encarcia_control_whiteFly/encarsia.asp http://www.ipm.ucdavis.edu/PMG/T/I-HO-TVAP-EF.011.html Average # of Scales/Leaf Average Number of Whitefly Scales/Leaf Show House Centranthenum punctam 300 Enranthemann pulchemllum 250 200 Cananga fruticosa 150 100 Mitrephora wangii 50 0 1 2 3 4 5 Figure 5. Weekly average number of whitefly scales per leaf in the Show House throughout 5 weeks of the experiment. 21 Table 1. Insecticide: Mavrick Aquaflow, MSDS Information Type Targets Active Ingredients Health Safety Alert Effects on Wildlife Synthetic pyrethoid Aphids, Earwigs, Rose Leaf Hoppers, Tent Caterpillar, Orange Tortrix Caterpillar, Omnivorous Looper, Io Moth, Oblique Banded Leafroller, Fruit Tree Leafroller, Omnivorous Leafroller, Thrips, Whiteflies & Mites. Tau-fluvalinate Caution Oral LD50 (mg/kg) Dermal LD50 (mg/kg) Signs and Symptoms Overexposure 5150 (Rat) >2100 (Rabbits) Causes eye and skin irritations. Aggravates asthma and bronchitis. Ingestions cause strong emetic response in animals. Typical symptoms are salivation, nausea, vomiting, and initial excitation followed by sedation. Uteral toxicity causes anorexia, depression and decreased body weight. Highly toxic to fish. Table 2. Insecticide: Conserve, MSDS Information Type Targets Active Ingredients Health Safety Alert Effects on Wildlife Fermentation derived insecticide Tent Caterpillar, Sawflies, Midges, Mites, and Thrips Spinosyn A & D Warning Oral LD50 (mg/kg) Dermal LD50 (mg/kg) Signs and Symptoms Overexposure >5000 (Rat) >5000 (Rabbits) May cause slight temporary eye irritation. Prolonged exposure is not expected to cause adverse affects. Highly toxic to aquatic invertebrates, toxic to bees, moderately toxic to fish. 22 Table 3. Insecticide: Talsar WP, MSDS Information Type Targets Active Ingredients Health Safety Alert Effects on Wildlife Synthetic Pyrethroid Cone Worms, Seed Bugs, Seed Worms Bifenthrin Warning Oral LD50 (mg/kg) Dermal LD50 (mg/kg) Signs and Symptoms Overexposure 632 (Rat) >2000 (Rabbits) Contact with bifenthrin may occasionally produce skin sensations such as rashes, numbing, burning or tingling, which usually subside within 12 hours. Contact with bifenthrin may occasionally produce skin sensations such as rashes, numbing, burning or tingling, which usually subside within 12 hours. Bifenthrin is classified as "restricted use" due to high toxicity to fish and aquatic invertebrates. It is slightly toxic to birds, moderately toxic to mammals, and highly toxic to bees. Table 4. Insecticide: Avid, MSDS Information Type Targets Active Ingredients Health Safety Alert Effects on Wildlife Glycoside Spider Mites, Thrips, Leafminers, Whitefly, Aphids Abamectin Oral LD50 (mg/kg) Dermal LD50 (mg/kg) Signs and Symptoms Overexposure 648 (Rat) >2000 (Rabbits) Causes substantial, but temporary eye injury; skin irritation on some individuals. Contains ingredient known to cause birth defects. May be fatal if swallowed. Warning Toxic to fish, wildlife, and bees. 23 Table 5. Insecticide: Marathon, MSDS Information Type Targets Active Ingredients Health Safety Alert Effects on Wildlife Systemic (Chloronicotinyl) Adelgids, Aphids, Armored Scale, Fungus Gnat, Japanese Beetle, Lacebugs, Leaf Beetles, Leafhoppers, Leafminers, Mealy Bugs, Psylids, Root Weevil Complex, Soft Scale, Thrips, Whiteflies, White Grub Larvae Imidacloprid Caution Highly toxic to aquatic invertebrates. Oral LD50 (mg/kg) Dermal LD50 (mg/kg) Signs and Symptoms Overexposure >4820 (Rat) >2000 (Rabbits) Mildly irritating to the conjunctive of the eye. Excessive long-term exposure to repirable crystaline silica may cause silicosis, a form of disabling, progressive and sometimes fatal fibrotic lung disease. Severe and permanent lung damage may result. Tabe 6. Insecticide: M-pede, MSDS Information Type Targets Active Ingredients Health Safety Alert Effects on Wildlife Insecticidal soap Aphids, Bristly Rose Slug, Earwigs, Rose Leaf Hoppers, Tent Caterpillars, Scale, Thrips, Mites & Whiteflies. Potassium salts of fatty acids Warning Oral LD50 (mg/kg) Dermal LD50 (mg/kg) Signs and Symptoms Overexposure >5000 (Rat) >2000 (Rabbits) Mildly irritating to the eyes and skin. Highly toxic to aquatic invertebrates. Methanol can be fatal or cause blindness if swallow in quantity; however, levels of methanol in this product are not expected to pose an ingestion hazard. 24 Table 7. 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