Thrips–Tospovirus Control Systems Progress Reports - June 2001
Team Leader: Michael P. Parrella
Department of Entomology, University of California at
Davis
Ms. Julie Newman
UC Cooperative Fxtension,
Ventura and Santa Barbara Cos.
Mr. Steve Tjosvold
UC Cooperative Extension,
Monterey and Santa Cruz Cos.
Proposed Project Duration
Start: 7/1/98
Completion: 7/1/02
Total Estimated Cost:
Year 1: $126,000
Year 2: $192,00
Year 3: $196,000
Year 4: $200,000
Project Research & Anticipated Industry Benefits
The main thrust of this project is to combine researchers
from across the country to help solve the thrips/tospovirus problem. Each of
these individuals brings a unique set of skills to the
table and is responsible for developing a piece of the
Thrips/Tospovirus 1PM puzzle. Michael Parrella (California) is concentrating on
developing and validating sampling plans, thresholds, and identification keys
for thrips attacking floriculture crops. In addition, he is examining
the performance of reduced risk pesticides and their compatibility
with natural enemies. Michael Brownbridge (Vermont) is looking at ways to
improve the performance of the fungus, Beauveria bassiana, for thrips control. Kevin
Heinz is evaluating the potential of a new biological control agent of
thrips, the nematode Thripinema. In addition, Dr.
Heinz is determining the utility of commercially available natural
enemies such as predatory mites in the genus Amblyseius. Diane Uliman
(California) is trying to understand the population dynamics of viruliferous
thrips in floriculture crops in the field and greenhouse and is examining the
potential of ‘induced’ resistance for control of the thrips and virus. Dr.
Uliman is also developing a rapid bioassay to tell if an individual thrips is a
virus vector. Beth Mitcham (California) and Arnold Hara
(Hawaii) are examining various postharvest treatment methods to control western
flower thrips. The Farm Advisors in this project (Karen
Robb in San Diego, Steve Tjosvold in Santa Cruz and Monterey,
and Julie Newman in Santa Barbara and Ventura) will be heavily involved
this coming year as we try to implement and validate this
holistic 1PM program with growers throughout California. The Farm Advisors are
receiving funds from the USDA-ARS National Floriculture
Initiative to cooperate in this project.
Each of us is making good progress and
we have an agreement to submit five
articles (one from each of us) to a trade magazine in the
coining year. This has been delayed until the final year of the project to allow
for a more complete overview of accomplishments. Publishing of the thrips
identification key is taking more time than expected, and we are considering
other outlets for publication. Many of us will travel to Italy (July 2-7)
and we will be participating in the 7th International Symposium
on Thysanoptera http://www.keele.ac.uk/depts/aep/thrips2001).
Summary of Professional/Published Information
Arthurs, S. and K. M. Heinz. 2001. Effect of temperature
and moisture on the development, transmission and reproduction of T.
nicklewoodi in F. occidentalis Journal of NematologyArthurs, S. and K. M. Heinz. 2001. In
vivo rearing of Thripinema nicklewoodi (Tylenchida:
Allantonematidae), an obligate parasite of Frankliniella
occidentalis (Pergande) (Thysanoptera: Thripidae). Journal
of Economic Entomology.Arhturs, S. and K. M. Heinz. 2001. Prospects for
entomopathogenic and entomophilic nematodes in foliar environments. Environmental
EntomologyCasey, C. A. L. Bolkan, J. Newman, K. Robb, S. Tjosvold, J.
MacDonald and M. P. Parrella. 2001. Glasshouses. Integrated pest management
works for rose growers. GrowerTalks, July 2001 (in press).Heinz, K.M. 2001. Pests: What arc they, where did they come
from, and what can be done about them? Greenhouse Business 7(1):
19-20.Heinz, K.M. 2000. Biological control in ornamental crops.
I.D. Greene [ed.], Proceedings of the 16th Conference on Insect and Disease
Management, The Society of American Florists; Alexandria, Virginia. San Jose,
CA, January, 2000.Krauter, P.C., S.P. Thompson, and K.M. Heinz. 2000.
Greenhouse IPM: Case Study. Ornamental Outlook. 9(3):
37-38, 42.Parrella, M., C. Casey, J. Newman, K. Robb, and S. Tjosvold.
2000 Integrating natural enemy release and biological pesticides for insect
management. Pp. 57-58 in Proceedings of the California Conference on
Biological Control, Riverside, Ca.Parrella, M.P. 2000. 1PM on the Flower Farm. Roses as the
Model System. Proceedings of Flowers 2000, The Australia Flower Conference.
Crop Production Program, pp. 1- 3Parrella, M. P. 2001. Redefining Integrated Pest Management
for the greenhouse. GM Pro. (submitted).
Parrella: This past year we concentrated on validating
our sampling plan for western flower thrips in greenhouse roses. Previous work
has indicated that sticky traps can be a cost effective, accurate tool for
thrips monitoring in greenhouse roses. Current recommendations from that work
suggest using 7 to 9 blue or yellow cards per pest management unit (the same or
similar rose cultivar). It has been determined that 25 to 50 adult western
flower thrips per card per week corresponds to 1 to 2 thrips per flower, which
is the action threshold. Yellow sticky cards were placed in each pest management
unit of the test greenhouses. Each week the number of western flower thrips on
each card were counted and the card was replaced. Ten harvestable flowers were
also collected weekly from four of the pest management units in each greenhouse.
Thrips per flower were counted to confirm that trap count is a predictor of the
number of thrips per flower. Validation took place for 7 weeks at the test
greenhouse in Watsonville and for 20 weeks at the test greenhouse in Nipomo.
Data was collected for 7 weeks in red, yellow, white, pink and orange flowers in
Watsonville. Data was collected for 4 weeks (red flowers) and 20 weeks (pink
flowers) in Nipomo. The data collected to date supports the sampling plan that
we have developed. In Watsonville, we found that the regression equation
generated from this data predicted that 25 to 50 thrips per yellow trap would
correspond to 0.65 to 1.19 thrips per flower. (Table 1). In Nipomo, we
found that the regression equation generated from this data predicted that 25 to
50 thrips per yellow trap would correspond to 0.28 to 0.75 thrips per flower. In
both locations, the range varies with flower color. However, both are acceptable
ranges, as noticeable injury generally does not occur until there arc 2 thrips
per flower. Similar work on validation is ongoing in Vermont and Texas.
| Flower color |
r2 |
p |
Equation of the regression line |
Predicted thrips per flower at 25 thrips per card |
Predicted thrips per flower at 50 thrips per card |
|
All |
0.36 |
<.0001 |
y = 0.06119 + 0.02265x |
0.65 |
1.19 |
| Yellow/orange | 0.26 | 0.1053 | y = 0.6013 1 + 0.01962x | 1.09 | 1.58 |
| White/pink | 0.45 | 0.0002 | y -0.0754 + 0.01936x | 0.41 | 0.89 |
Table 1
. Predicted number of western flower thrips per
flower at 25 and 50 thrips per yellow sticky trap (Watsonville).
Brownbridge: In previous phases of this project we
identified both the solid full cone (SFC) and flat fan (FF) type of nozzles as
those most efficiently depositing spores on plants and providing some coverage
on lower leaf surfaces. Using the train system, however, we decided to
reevaluate both the hollow cone (HC) and fog (FOG) type nozzles along with the
SFC and FF types to provide a complete evaluation of their relative efficacy in
two different spray systems. This was particularly important for the hollow cone
nozzle where movement of the plants in the spray stream could enhance coverage.
In a replicated experiment, 4 nozzle types were tested at a spray pressure of 40
psi and at angles of 10 or 30¬?. Two speeds were used for the plant train (20
and 42 fl/mm) and a standard spray concentration of i07 spores/ml was
used. Sprays were applied to single row of plants traveling perpendicular to the
direction of the spray. Leaves were sampled from the lower and upper canopy and
spore deposition was quantified on both the upper and lower leaf surfaces.
The type of nozzle used and the speed of the train
significantly affected the total number of spores deposited and the rate of
deposition on the upper or lower leaf surfaces. Only the flat fan and solid full
cone nozzles deposited spores on the lower leaf surface at a rate equivalent to
10% or more of the of the spores deposited on the upper leaf surface. As
indicated in our previous research, the hollow cone and fog nozzles were
relatively inefficient and were thus excluded from any subsequent evaluations.
Consistent with our previous findings, spray angle did not influence the rate of
spore deposition onto lower leaf surfaces, with 6205 and 6354 spores/cm2 for
10 and 30 degree spray angles, respectively. As might be expected, more spores
were deposited on the side of the plant closest to the spray nozzle.
In the second experiment flat fan and solid cone nozzles of
two different sizes were included, delivering 0.10 and 0.40 gal/mm (flat fan),
and 0.06 and 0.19 gal/mm (solid full cone). A single row of plants was sprayed
on both sides at a spray angle of 10¬?, 40 psi of pressure and a single train
speed of 20 ft/mm.
As expected, by increasing the size of the spray aperture, we dramatically
increased the number of spores deposited on the lower leaf surface. The
influence of larger nozzle sizes is consistent with our findings from last year,
even though the nozzles used this year were of a relatively smaller aperture to
reduce delivery volumes. Spraying on two sides of a plant and using a slightly
larger nozzle size significantly increased the total number of spores deposited.
Plant coverage was relatively consistent between the upper and lower leaf canopy
but dependent on the type of nozzle and the spray volume applied.
In the final experiment we tested the effectiveness of spray applications
made to 4 rows of plants using only the larger sizes for both the flat fan and
solid full cone nozzles (FFTP8004 and SFC-l). We also moved the plants through
the spray stream at higher speeds (42 and 59 fl/mm) to more accurately replicate
actual spray conditions. Although all of our data to date indicates that spray
angle has only minimal influence on spore deposition, we assessed two angles of
spray - 10 and 30¬? ¬ó to confirm the validity of this observation. Spore
suspensions were applied from both sides of the 4 rows of plants and a plant in
the first and second row was sampled. Four leaves approximately at right angles
to each other were sampled from the mid-canopy to determine spore deposition
rates on the upper and lower leaf surfaces.
At higher speeds, a greater proportion of spores were deposited on the lower
leaf surface than occurred at lower speeds. Both the nozzle used and spray speed
affected the number of spores deposited but they did not interact significantly.
Spray angle had no significant influence on the rate of deposition. Overall the
flat fan nozzle appeared to be most effective in delivering spores into the
interior section of the second row of plants.
Conclusions From 2000-2001 Spray Trials
Our findings provide some clear indications relevant to the use of low
pressure sprayers for targeted delivery of fungal spores. While slower movement
of a spray across a row of plants will increase the total number of spores
deposited, higher speeds tend to facilitate deposition onto the lower leaf
surfaces. Flat fan nozzles and solid full cone nozzles have consistently been
shown to be the most effective in delivering spores to the undersides of leaves
and onto plants in general. The results from the current trails, combined with
those previously reported, reinforce earlier conclusions that nozzle size is
important for spore delivery and economic efficiency. Use of nozzles that
deliver high spray volumes, a SFC 10 for instance, are not efficient and are
wasteful. Too fine a spray limits spore deposition on lower leaf surfaces. Use
of a nozzle that produces slightly larger droplets appears to move the leaf
canopy allowing greater deposition on lower leaf surfaces.
To obtain thorough plant coverage using a low pressure sprayer may require
that applications are made down both sides of a row (or rows) of plants.
However, this system requires more material and time, and may not be feasible
given the close spacing of plants in most production houses. Our previous
research on hydraulic and electrostatic sprayers demonstrated that good plant
coverage could be achieved at distances of up to 3 meters. However, this work
was done using single, stationary plants and now needs to be evaluated using
multiple rows of moving plants.
Heinz: In our lab in Texas we have been working with
two new approaches to thrips biological control. One involves the use of a
parasitic nematode of western flower thrips (WFT) about which little is known, Thripinema
nicklewoodi. Nernatodes, also called roundworms, are microscopic worms that
are found in diverse habitats and many species are parasites of plants or
animals. Research conducted last year demonstrated promising results in terms of
efficacy (albeit on a small scale) and in our abilities to mass produce the
parasite. We have been able to maintain a healthy culture for approximately 2
years, and have developed techniques for scaling up production. This year, we
focused improving our rearing procedure and on several biological aspects,
information necessary for conducting full efficacy trials.
Quantifying rearing procedure: Obtaining regular supplies of
nematode-infected thrips is essential for subsequent experiments and
implementation of a control strategy. The nematode, an obligate parasite of
western flower thrips (WFT) must be cultured on WFT. We have developed a simple
rearing procedure whereby colonies are reared in discrete cohorts on excised
bean leaves in plastic boxes. To maintain the nematode in culture, WFT from the
colony together with thrips from which nematodes are actively emerging are
aspirated into 1 .5rnl Eppendorf vials. Under suitable conditions of temperature
and moisture, transmission of the nematode is achieved. Following a temperature
dependent incubation period in plastic colony boxes, the infection status of
surviving thrips is checked by isolating thrips individually in Eppendorf vials
and assessing nematode emergence. Alternatively, thrips may be dissected out
directly under a dissecting microscope. In a recent bioassay, using this
technique we were able to achieve an average increased output to input ratio of
infected thrips of 2.27 ± 0.28 (n=12 vials) between subsequent generations
(based on an inoculation of 50 propupae/pupae + 4 infected adult thrips and a
48-hr exposure period at 25¬?C; mean survival rate = 0.3 x mean transmission
rate = 0.64). Other parameters that have recently or are currently being
estimated for their effect on both transmission and host survival during
nematode incubation are: the effect of host stage exposed (2nd instar larvae /
pupae / adults); exposure period (48 vs. 24 hour) and container (Eppendorf vial
vs. larger shell vial) (n=6 vials).
Experimental studies: Understanding the influence of
temperature and moisture on hostnematode biology is central to predicting
conditions under which T nicklewoodi may be an effective biological control
agent. Despite this need, there are no published data on the effect of
temperature for any life-history parameter of T nicklewoodi (apart from
the survival of free-living nematodes in water) or any other Allantonematid spp.
In response, a range of bioassays are being conducted to evaluate the
importance of temperature on in vivo incubation (pre-nematode release) period,
number of release days, daily release rate and host survival and the effect of
temperature and moisture on transmission (see table 1 for an examples of early
results).
Table 1. Nematode development at 2 different temperatures
| Incubation temp. (¬?C) |
In vivo incubation period (days) |
No. release days (in Eppendorf vials) |
No. nematodes| released/day |
Total no. nematodes released/thrips |
| 20 | 15.8±1.5 | 7.2 ±0.9 | 22± 1.2 | 149.8±16.2 |
| 25 | 12.4 ¬± 0.64 | 6.7 ¬± 04 | 18.5 ¬± 0.43 | ———- |
Ullman/Robb: This year our program has been focused on technology
transfer of the petunia indicator program to professional scouts and on
exploring elicitors of induced resistance in the laboratory. In the field, we
have placed an emphasis on evaluating the use of petunia indicator plants for
detecting the presence of tospovirus-transmitting thrips in ornamental
production for its user-friendliness. Professional scouts were trained to
distinguish thrips feeding scars from the local lesions, which indicate the
presence of viruliferous thrips. Scouts monitored greenhouse and field-grown
flower crops weekly and used the information gained from the indicator plants to
choose which, if any, pesticide should be used that week to control thrips. As a
result of this information, applications of highly effective pesticides, such as
Conserve, were minimized and tospovirus incidence was maintained at very low
levels. In the laboratory, we have conducted trials with jasmonic acid (JA) as
an elicitor of thrips resistance and BHT as an elicitor of tospovirus
resistance. Elicitation of resistance with jasmonic acid alone and/or in
combination with BHT resulted in a dramatic reduction in thrips feeding on
tomato. We tested JA alone and found no reduction in tospovirus transmission,
however, we are hopeful that JA and BHT combined may be effective against both
the thrips and the virus. We also began studying JA induced resistance in
chrysanthemum. We have tested potted chrysanthemum varieties and found that
induction occurs and it results in reduced feeding by the thrips, however not as
dramatic as was observed in tomato. We plan to explore several chrysanthemum
varieties, as well as lisianthus in the laboratory and we will conduct
greenhouse trials in San Diego with a grower cooperator.
Mitcham/Hara: The tolerance of chrysanthemum cuttings and western
flower thrips to forced hot air treatments was tested. Cuttings and insects were
exposed to 45, 48, 50, and 52¬?C for up to 120 minutes. Results indicate that
plant tolerance is good up to 120 minutes at 45¬?C and 48¬?C. Plant tolerance to
the higher temperatures, 50¬?C and 52¬?C, was good for up to 80 and 40 minutes,
respectively. Treatments within the tolerance limits of the chrysanthemum
cuttings were effective against melon aphid and western flower thrips.
Preliminary results indicate that exposure to 48¬?C for 80 minutes or 50¬?C for
40 minutes would be effective against western flower thrips. Exposure to 45¬?C
for 90 minutes or 48¬?C for 80 minutes would be effective against melon aphid.
In addition to western flower thrips and melon aphid, research on the control of
two-spotted spider mites, an agromyzid Ieafminer and greenhouse whitefly is
currently underway.
A postharvest treatment to control western flower thrips and
melon thrips is being sought for Hawaiian grown Dendrobium orchids. The effect
of high temperature 35 C in combination with 55% carbon dioxide or 1 .5% oxygen
was tested. In addition, several chemical fumigants (acetaldehyde, methyl
formate and ethyl forniate) were tested as alternative postharvest control
measures. Complete insect mortality occurred at 48 hours in 1.5% O 2 (35¬?C) and
at 12 hours in 55% CO2 (35¬?C). Although the 55% CO2 treatment
resulted in a shorter exposure time for insect control, these treatments also
resulted in considerable damage to the orchids. Of the fumigants, acetaldehydc
proved to be the most promising. Pretreatment with 0.5-1.0% acetaldehyde for 1
hour followed by exposure to air or 1.5% 02 (8-24 hours) at 35¬?C resulted in
high insect mortality and minor phytotoxicity. Acetaldehyde is a volatile
compound naturally produced and metabolized by plant materials, and may be
considered as Generally Recognized as Safe (GRAS) by regulatory authorities.
Additional work is needed to determine the effects of concentration and exposure
time on thrips mortality and orchid tolerance.
