Download Maribeth Kniffin

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.
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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
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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).
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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).
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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
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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.
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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 ½ 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
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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
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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
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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
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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
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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).
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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).
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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 ½ 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,
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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
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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
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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.
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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.
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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.
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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
days
10
days
Figure 3. Diagram of Whitefly (T. vaporariorum)and Parasitoid Wasp (E. formosa)
lifecycle. http://www.buglogical.com/encarcia_control_whiteFly/encarsia.asp
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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
250
Enranthemann
pulchemllum
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 ½ 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.
Total Amount
of Insecticides Used 2006
Marathon (oz.)
34
M-pede (fl. oz.)
619
Avid (fl. oz.)
5.75
Conserve
10
Synergy (oz.)
90.5
Talstar (oz.)
19.75
Mavrik (oz.)
14
25
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27