Tree Fruit Research & Extension Center

Insect Ecology & Behavior Laboratory

Vince Jones' Research


Inexpensive Immuno-marking Systems to Measure Insect Movement Patterns

Over the past 4 years Dr. Jones' Insect Ecology and Behaivior lab has worked to develop immuno-marking systems that can be used to determine the movement patterns of insects that naturally occur in the environment. The markers are common food products that can be sprayed on an area using commercially available spray equipment. Depending on the marker and the dose applied, the insects can be marked directly by the application or indirectly by walking across the dried residues. Insects are then collected and processed with commercially available antibodies to determine whether they have the mark or not. Used correctly, there is no cross-reaction between the different markers, allowing us to detect multiple marks on a single insect. The collapsing panels presented below contain more detailed information about this project and the techniques used to mark insects to measure their movement.
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Overview: Need for Ways to Mark Large Areas to Track Inter-area Movement Patterns

Studies of insect dispersal in different ecosystems have relied on a variety of methods to determine movement patterns. Typically, marked insects are released from a limited number of sites, and traps of some sort are used to map their movement patterns. Marks that have been used include fluorescent powders, rare (trace) elements, radioisotopes, internal or external dyes, and insect mutilation (e.g. punctures of an elytra). The marks are applied to lab-reared or field-collected insects depending on the particular study. While each of the marking and collecting methods has advantages and disadvantages, all of these studies typically use the dispersal of a relatively small portion of the population from a few release points to extrapolate the population level movement patterns. A large increase in our understanding of movement patterns would become possible if a large number of wild insects could be marked directly in the field and, further, if several different marks were available that could be used to track inter-area movement.

The major problem is that few of the above techniques can be used to mark the naturally occurring insects within an area. The barriers are costs, time, and the lack of multiple marks. Immuno marking is a technique developed by James Hagler, who is currently at the USDA-ARS Western Cotton Research Lab in Phoenix, AZ. James developed a marking technique that used commercially available antibodies in an ELISA format to detect highly specific vertebrate proteins (chicken and rabbit IgG) applied to insects. The technique is simple, rapid, and sensitive. Unfortunately, the cost of the specific marker proteins is roughly $500 per liter, which restricts the use of this technique to marking in small enclosed areas or spiking diet of lab reared insects.

The techniques described in this web site and the references within overcome the cost barrier by using inexpensive and easily obtained food proteins that can be diluted in water and sprayed using normal pesticide application methods or applied as a dust. All the markers can be detected in the ppb range using indirect ELSIA techniques.

What is ELISA?

ELISA stands for Enzyme-Linked Immuno-Sorbant Assay. It was developed 50+ years ago, although it has undergone a number of changes since the original protocols. Most commonly, the assays are run in 96 well plates (8 rows x 12 columns) that are made of polystyrene. We are using what is called an indirect ELISA to detect the proteins we are spraying in the field.

Indirect ELISA

  1. Insects are collected individually and treated carefully to prevent cross contamination. Individual insects are placed in a 1.5 ml microcentrifuge tube and 1 ml of buffer is placed in the tube for 3 minutes. The insect is then removed and discarded.
  2. An 80 µl aliquot of the buffer is then applied to a single well in the ELISA plate. This is then incubated and the proteins in the sample attach to the sides of the well.
  3. After a suitable incubation period, the wells are washed with a buffer + detergent to remove any loosely bound proteins.
  4. At this point, a non-specific (non-reacting) protein is added at a fairly high concentration to the wells and incubated. This is done so that the portions of the well where no proteins have become bound are “blocked” – this prevents an antibody or reagent later in the assay from binding to the well and potentially giving a false positive.
  5. An antibody specific to the marker (called a primary antibody) is then placed in the well and incubated. During the incubation process, the antibodies bind tightly to the marker proteins. Because the wells have been blocked, the antibodies cannot bind directly to the wells to later affect the ELISA readings. The primary antibodies are generated in a typical lab animal (e.g., goat, rabbit, chicken, etc.). The primary antibody is typically named something like rabbit anti-chicken egg albumin. This would indicate the antibody was developed in a rabbit against chicken egg albumin.
  6. The wells are then washed again to remove any excess and unbound antibodies.
  7. The secondary antibody is added at this point. This antibody recognizes the primary antibody, by looking for the source of the antibody. For example, in our rabbit anti-chicken egg albumin antibody mentioned above, the secondary antibody would attach to a protein of a given conformation that would identify it as coming from a rabbit. The secondary antibodies are also conjugated typically with either a peroxidase or alkaline phosphatase enzyme. A secondary antibody to recognize the primary antibody would then be named by its source animal and what it recognizes. In our case, we use a donkey anti-rabbit IgG (H+L) with peroxidase conjugate. The secondary antibody is incubated in the well and attaches to the primary antibody.
  8. We then wash out the wells, which removes the unbound and excess secondary antibody.
  9. We add an indicator dye (we use TMB) to the wells and incubate for 5-20 minutes. The wells that are positive for the marker proteins will have the secondary antibodies present with the peroxidase enzyme present which causes the indicator to turn a blue color depending on the concentration of the original sample.
  10. We then add a 2 N Sulfuric Acid solution (“stop solution”) to the wells, which inactivates the peroxidase enzyme and turns the wells a yellowish color.
  11. The plate is then read using the plate reader at 450 nm.

Step-by-Step Diagram

For each of the markers, the major differences are simply the primary antibodies used. Our egg, soy, and wheat assays both use the same secondary antibody since the primary antibodies were all developed in rabbits (donkey anti-rabbit IgG (H+L) with peroxidase conjugate). The secondary antibody used in the casein assay is different because the primary antibody was developed in a sheep; thus it is a donkey anti-sheep (whole molecule) peroxidase conjugate.

Although the major differences are the antibodies, the washes, detergents, blockers and antibody diluents are slightly different and need modified as laid out in the protocols. We have tweaked virtually every part of each of the assays for the most consistent and sensitive responses.

Other Types of ELISA

There are a wide variety of different types of ELISA reactions, each with its strengths and weaknesses. The ELISA Guidebook (link) has details of each, their strengths and weaknesses, and good general information on ELISA in general – highly recommended.


There are slightly different protocols for processing a sample for each of the marker proteins. We actually have two separate protocols for the soy protein, one that works well with moths, and one that works well with pear psylla (which is very tough to work with). Each of the protocols is available below as a pdf file and lists each of the steps needed, along with the times for incubation, etc.

Casein Protocol      Egg Albumin Protocol
Soy-Moth Protocol      Soy-Psylla Protocol
Project 1 – Determining the movement of Colpoclypeus florus (Hymenoptera: Eulophidae) between extra-orchard locations to obliquebanded and Pandemis leafrollers inside apple orchards
This is an on-going project performed in collaboration with Dr. Tom Unruh, USDA-ARS, Wapato, WA.


Work by Unruh, Brunner and Pfannenstiel has led to the use of rose gardens adjacent to leafroller-infested apple orchards to provide the parasitoid C. florus with overwintering fifth instar larvae of the strawberry leafroller. PLR and OBLR overwinter primarily as second instars and are unsuitable C. florus overwintering hosts. The idea is that the rose gardens act as a “nursery” for large numbers of C. florus that will emerge early in the spring and begin parasitizing the overwintering generation of either PLR or OBLR. Without suitable overwintering hosts, C. florus parasitism is generally negligible until late in the summer generation, often necessitating pesticide applications to prevent fruit damage. A key point not answered so far in this research is the area of influence of each rose garden (i.e., its “active space”).


The rose/strawberry gardens are extremely difficult to spray because of the multiple layers of the canopy. We tried marking the area using an orchard handgun, but had little success in the first year. Our problems were magnified because we could not get C. florus to come to any sticky trap that we had available. To solve the problems, we first covered the rose garden with thule (a fine mesh screen) and switched to applying soy flour using a common fertilizer spreader ( Fig. 1). The soy flour coated the plants and also was caught by the thule. Parasitoids that emerged from the leaf litter either picked up the soy flour on the leaves or when they crawled through the netting ( Fig. 2).

To overcome the trapping problem, we developed leafroller-baited traps. A shoot was inserted through a card coated with sticky stuff and into a 100 ml floral vial. The shoot was then infested with 4-5th instar OBLR larvae in the lab. The next day, we eliminated shoots that leafrollers did not colonize, and placed the others in the orchard at different distances from the garden (Fig. 3).


We are still collecting data on this project, but our first year’s data showed good marking and recovery of parasitoids at least 171 feet away from the garden. This means the area of effect was at least 2 acres, and likely much larger. We are placing our traps further away from the garden, marking a larger part of the garden, running it at multiple locations, and throughout the entire season.

Project 2 – Movement of pear psylla predators between the ground cover and tree canopy
This is a current project performed in collaboration with Dr. Dave Horton, USDA-ARS, Wapato, WA.


Horton’s work to date has shown that it is easy to prompt build-up of natural enemies in the ground cover. However, the major problem in determining the impact of those natural enemies on psylla populations in the trees is the inability to determine the movement patterns of those natural enemies from the ground cover to the tree. To date, he has not been able to consistently show significant increases in the number of predators in the trees in grass vs. cover crop areas, even though predators are much more common in the cover crop area ground cover. While there are some indirect methods and changes in plot design that he is pursuing that may help determine the importance of these (potentially) added natural enemies, the protein marker should be a very powerful tool in determining their movement patterns.


Both last year and this year we applied the egg marker to the orchard ground cover with a weed sprayer mounted on an ATV ( Fig. 1). Last year we applied the marker as a 20% solution and had 97% marking for insects collected from the ground cover. This year we reduced the rate applied to 10% egg whites and marking remained nearly the same. Insect samples were collected from both the ground cover and the canopy over the course of the experiment. Ground insects were tested for presence of the mark to determine marking success, and tree-collected insects were tested for the marker to determine movement between the two areas.

This year we concentrated on expanding our identification of the specimens collected. In particular, we identified the adult ladybird beetles and lacewings to species so that we could determine habitat specificity and the tendency of each species to move between the ground cover and the canopy. We also collected and tested immatures of ladybird beetles, green lacewings, Anthocoris, and Deraeocoris. For the immature ladybird beetles and lacewings, identification to species was not possible.

We also classified each predator species collected in the tree canopy as to its habitat preference. To do this, for each species we calculated the percentage of the total captures (ground + tree) that occurred in the tree. If greater than 70% were collected in the tree, we tentatively considered this a species that preferred the tree; if between 30 and 70% were collected in the tree, the species was considered a “generalist”; and if less than 30% were collected in tree it would be considered a species that preferred the ground cover. This classification should be viewed as a very rough guide, in part because the sampling methods were different between the tree and ground collections, so that we may have some differences in efficiencies between the two sampling methods that would distort the percentages.

The proportion of individuals in each species collected in the tree canopies that were positive for the ground cover mark was calculated. This value, along with the habitat preferences classification was used to determine how important the ground cover was in population dynamics occurring in the canopy.


Using the classification system described above, four of the 14 species collected in the tree canopy could be described as “tree species” ( Table 1). Of the four tree species, two were commonly collected (Anthocoris tomentosus and Deraeocoris brevis), with 31% of the Anthocoris and 15% of the Deraeocoris collected in the tree canopy testing positive for the ground cover marker. Immature Anthocoris and Deraeocoris were also found to move between the ground cover and the canopy, but at roughly 1/3 the rates of the adults. A species of lacewing, Chrysoperla plorabunda and a ladybird beetle, C. septempunctata, also moved between the ground and tree with 23.5 and 12.5% of those collected in the trees testing positive for the ground cover marker, respectively. The ladybird beetle Harmonia axyridis was only found in the tree canopy, but 8.6% of that species also tested positive for the ground cover mark.

Spiders (a mix of species) were classified as habitat generalists and were found in nearly equal abundance in the ground and tree collections. In terms of overall abundance, spiders were the second most common predator found in the tree samples (first was Deraeocoris). About 16% of the spiders collected in the tree samples tested positive for the ground cover marker.


The ground cover is visited by a number of the different predators, even ones that abundance data would suggest are “tree” species. The Anthocoris found in the ground cover likely originated in the tree and only visit the ground cover for short periods (suggested by the high percentage collected in the canopy that tested positive for the ground cover mark and the low numbers found in the ground cover). Surprisingly, even the less mobile immatures move down to the ground cover and back. The importance of the ground cover cannot be further defined with the experiments we have performed to date, because we did not collect prey abundance data (pea aphid in the cover crop and pear psylla in the canopy); therefore, the tendency of predators to move between the habitats and switch prey fed upon cannot be determined.

Project 3 – Movement patterns of codling moth and the effect of mating disruption on movement patterns.


Codling moth (CM), Cydia pomonella L., is the most important pest of apples in the western US apple producing areas. While some research on codling moth movement has been done, to date only colony-reared moths have been used. Research in this area needs to be done because moths in colony experience conditions totally unlike those in the field, commonly resulting in behavior and size anomalies. In terms of behavior, the confined conditions, high density and relatively minimal flight requirements to insure mating may allow weaker males to mate and females may become less likely to discriminate among males. This can lead to a shift in the lab colony toward less fit males and females that do not react the same as the wild moths.

We need to understand how females and males respond to an area under MD. For example, in CM, females can detect the pheromone levels in the orchard. Does this cause them to move out of the orchard or out of the area where pheromone concentrations are high? If so, is damage heaviest in those areas of a block where the pheromone concentrations are lowest throughout the majority of the flight curve? In addition, are males attracted from a non-MD area toward a MD area and thus increase the probability of mating within the MD area because of higher male density? Our studies in field cages and lab wind tunnels suggest that an increase in male density results in an increased probability of mating, particularly in MD situations, even though the time required for mate location is higher in these situations.


We set up two experiments, each at a different orchard and each consisting of seven one-acre plots. At the first orchard, mating disruption was not used for the first generation, but before the second-generation flight, Isomate C++ dispensers (400/acre) were applied to the center plot only. Also at the first orchard, we trapped in an adjacent orchard block that had MD present throughout both CM generations. In the second orchard, mating disruption dispensers were uniformly present throughout the plots and during both generations. We used high density trapping to follow CM movement in both situations. In each of the orchards, we used nine traps per acre (63 total) to trap moths. Combo pheromone/DA lures were used in both experiments, regardless of MD presence. The traps in each plot were numbered and their GPS coordinates were mapped. The GPS coordinates were used to determine the distance of each marked moth from the center of the plot of origin.


We calculated the distance from the origin for each marked moth captured. To summarize the distance flown, we sorted the data from shortest to longest distance and calculated the distance flown by 1, 5, 10, 25, 50, 75, 90, 95, and 99% of the marked moths as well as the average distance flown. The different percentiles were then graphed on the y-axis and the distance from the origin was graphed on the x-axis. This graph allowed us to compare the distances flown in the different MD situations and determine if differences existed.


First Orchard, No MD present, first generation: The trap captures were highest at the south end of the plot and in the adjacent orchard (treated with MD). Marked moth captures were highest in these same areas. Fifty percent of the marked moths were captured within 392 feet of origin, but the converse of this is that 50% of the population traveled more than 392 feet (Fig. 1A). The cumulative percentage catch was linearly related to distance.

First Orchard, MD present in the center plot only, second generation: As with the first generation, trap catches were still highest in the southern part of the plot and in the adjacent orchard. The distance moved and the shape of the cumulative percentage catch was virtually identical, indicating the MD plot was unimportant in the degree of movement (Fig 1B).

Second Orchard, MD present uniformly throughout the area, first generation: As with the other orchard, we found trap catches were clustered, with most of the moths caught in three of the seven acres monitored. However, a plot of the cumulative percentage of moths caught versus distance was not a straight line, but a curve (Fig. 1C). In addition, in comparison to the first orchard, the distances flown were markedly shorter. For example, 50% of the moths were caught within 128 feet of origin versus 392 feet in the first orchard.

Second Orchard, MD present uniformly throughout the area, Second generation: The second generation showed a pattern of movement similar to the results in the first orchard; the cumulative percentage moth catch was a linear function of distance (Fig. 1D). In this plot, 50% of the moths were caught within 308 feet of origin, compared to 128 feet in the first generation.

Difference in movement patterns between first and second generations in the second orchard (uniform MD): In reviewing the experimental procedures, we changed nothing in the protocols that would have affected the shape of the dispersal curve. However, a key difference is that the first orchard received Guthion® cover sprays for both generations. Guthion® was also used in the second generation in the second experimental orchard. For the first generation in the second experimental orchard, Assail® 70WP was applied for two cover sprays. Data from Jay Brunner’s lab shows that Assail® causes high codling moth adult mortality and may have been the reason for our different shaped curves. A material with a high activity against adults would tend to kill them as they move throughout the orchard, resulting in an overall shorter distance flown.

The data obtained clearly show that codling moth is highly mobile. Although the combo pheromone/DA lures catch primarily males, we do have some data on the females and it tends to be similar (at least in the average distance flown) to the male data. Our data suggest that 50% of the individuals travel more than 400’, but it also means the scale of our experiments need to be increased. In all cases, we obtained multiple marked individuals at the traps furthest from their origins (>800’). Our data do not indicate that MD plots are a source of moths or have the tendency to concentrate moths; however, more information on this point is required.

Available Markers & Costs

Currently, we have three marker proteins developed and have begun working on the fourth. The three markers are soy protein (as soymilk or soy flour), egg albumin (as egg whites), and casein (as cow's milk). All of these can be applied as powders or as liquid. We currently have the most experience in using the markers as liquids, but in one of our research projects are using the soy flour as a marking technique. The fourth marker we are currently developing is wheat flour.

The soymilk and cow's milk markers are typically applied at 20% solution and the egg whites at 10%. The costs per liter of each marker solution as of May 2006 from our sources are $0.26, $0.14, and $0.12 for soymilk, cow's milk, and egg whites, respectively.

We have tested a wide-variety of soymilk products and found a wide range of potency in our ELSIA reactions. Our studies have lead us to settled on Silk™ Organic Soy Milk Plain (White Wave, Inc. Boulder, CO 80301). We can buy this in a variety of sizes from a quart up to 5 gallon bags.

Cow's milk can be used as either non-fat or whole milk. We have not seen consistent differences between the different forms of cow's milk and just buy the cheapest at the time.

Egg whites can be bought in a variety of forms and all of them work well. For small-scale studies, we use All Whites™ (Papetti Foods, Elizabeth, NJ 07206), which is available in the local supermarkets. We use All Whites™, because it is 100% egg whites with no additives and the cost is similar to Egg Beaters™ or the generic brands found in the stores. For large-scale marking, we buy MGW Brand frozen pasteurized egg whites (M. G. Waldbaum Co., Wakefield NE 68784). We can order this directly from a distributor or at our local supermarkets. If ordering from the supermarket, there is typically a week or more lead-time required to get the product. The other tip is that the frozen egg whites take 2-3 days to thaw at room temperature, so order early!

Reagents Needed, Sources, Disposable items, and Cost Break Down

The spreadsheets below have the latest cost breakdowns, amounts needed, and sources for all the reagents commonly used for each assay. To run a plate (96 samples) a single sample costs 19.3 cents for egg, 14.7 cents for soy, and 17.7 cents for the casein assays. This cost includes the plate cost, antibodies, all solutions, pipette tips, tubes used to mix solutions, buffers, positive and negative standards, detergents, everything we could think about, except labor. Some of the prices could be cheaper by using different plates, buying different reagents (e.g. bulk instead or ready made packets or ready mixed solutions), but labor costs would increase and sources of error increase.

ELISA Supply Costs    Protocol Cost Worksheet
Required Equipment

The equipment required to run the ELISA’s vary depending upon how many samples you are going to run. If your needs are modest, you can forego some of the equipment, which will be noted below.

Essential Equipment:

Microplate Reader – A microplate reader is just a spectrophotometer that measures the color intensity in each well of a 96 well ELISA plate. The newer plate readers come with dual wavelength capability. These examine the samples in two wavelengths of light, one that measures the specific wavelength of the indicator dye (chromophore) that indicates the presence of the marker and one that is an off-wavelength. The off-wavelength is used to correct for turbidity of the sample or scratches or imperfections in the ELISA plate. The cost of the microplate readers is highly variable and ranges from around $4-18 thousand dollars and up. For the type of ELISA that we use, all the bells and whistles are not needed. I would recommend the following features:

  • Computer control – this allows you to read the plate, store the results in a program, and run some analysis in that program. At least some of the systems are available for MacOS X, Mac Classic, and Windows.
  • Dual-Wavelength capability.
  • Filters for 405, 450 and 650 nm

Our microplate reader is a Molecular devices Emax and cost around $7,000 dollars in 2001. You should also check out some of the used lab equipment web sites, which have greatly reduced costs and a guarantee that the machine will work.

Pipettes – You will need a wide range of pipettes for the assays. We use digital pipettes exclusively. I would recommend that in addition to a single channel pipette, that you purchase a 8 channel pipette and if you’re going to run a large number of plates, a 12 channel pipette. The 8 and 12 channel pipettes are roughly $600 new, but they can be bought rebuilt from several reputable sources on the web for roughly $200. The other pipettes (single channel) can be bought new as a set, singly, or rebuilt. We strongly recommend that you stick with one brand. In our lab we use Finnpipette, but there are a number of good brands around. Most of the pipettors can use universal tips, which cuts the costs considerably.
Sizes needed:

  • 8 or 12 channel pipettors: 50 to 300 µl
  • Single channel:
    • 1 to 10 µl
    • 10 to 100 µl
    • 100 to 1000 µl
    • 0.5 to 5 ml
    • 1 to 10 ml

High purity water source – If you are running only a few plates total, then you could buy high quality water from some of the typical suppliers. However, it is not very cost effective and frankly, you use a lot of water in the ELISA process. You can either buy a complete water system or make one on your own from various components. We have two systems:

  • Elga Purelab UHQ ultra pure water system (Elga LabWater, US Filter, Lowell, MA 01851) – This system hooks directly to a normal tap water line, has a reverse-osmosis membrane and a deionizer and a UV light to kill any bacteria. It also has various filters to remove particulates and bacteria. While it is a complete system solution, it is also relatively expensive to operate. On the plus side, it also has a built-in water quality (conductance) meter that constantly monitors water quality. If you have de-ionized water in the lab, you can by-pass the deionizer cycle and speed up the system. They also offer many different versions. Other companies also have similar systems (nano pure, etc.). Our system cost roughly $2200 in 2003.

  • Combination of de-ionizer system, filters, and a glass still – We filter the incoming water, run it through a commercial deionizer system, then put the water into a Barnstead Fi-streem III glass still (Barstea International, Dubuque, IA 52001). The still is auto controlled by the water level in a storage carboy. This system is cheap to operate, but more expensive in the start-up cost. The glass still is the major cost, somewhere around $4-5,000.

Vortexer – All of the solutions used in the ELISA process need to be thoroughly mixed. We use a mini-vortexer to mix everything in microcentrifuge tubes, 15 ml and 50 ml tubes.

Magnetic Stirring Plate – Used for making buffers or washing solutions where large volume is needed, we use a magnetic stirring plate, large enough to hold and mix 2-3 liters of solution.

Balance – A balance with the ability to measure 1 mg or better is needed for making various solutions.

Large freezer for sample storage – These can be standard freezers (-20°C).

Recommended Equipment (high volume):

Microplate Washer – There are a number of microplate washers on the market, some great and some absolute junk. The whole plate washers are quicker, but less flexible in terms of the ability to change solutions (i.e., they need to be purged and require about 1⁄2 liter or more solution). The strip washers are a lot more flexible, but 4-5 fold slower. We use a full plate washer (Biotek EL- 404) because of the speed issue.

Repeating Pipette – These are used primarily for filling the large number of microcentrifuge tubes with sample buffer – they really make it quick and simple. They cost roughly $400 new, can also be purchased rebuilt.

Dry Block Heat Plates – These are used to provide uniform heat across all the different wells of the ELISA plate. You can run your reactions at room temperature or overnight in the refrigerator, but it greatly increases the time required to run the ELISA. Dry block heaters are roughly $1200 to hold 3 ELISA plates.

Rotators – These are useful in mixing slowly the reagents in the ELISA plates to insure the optimal reactions. Some ELISA manuals recommend this over heat blocks to speed the ELISA reactions. Having a dry block heater on a rotator is even better, but is probably overkill in most situations.

Multiple Carboys – For holding different buffers, washing solutions, and water storage – Use whatever volume and shape is best for your application.

pH Meter – Useful in checking buffer solutions.

Miscellaneous Items:

Microcentrifuge Tube Holders – Laid out as 96 well format. These are useful in organizing samples for transfer to the microplates used in ELISA. These can also be used with tops to freeze the samples as a group for later re-testing if needed.

Universal Test Tube Holders – Used to hold 15 ml, 50 ml, and 1.5 ml tubes. These are used to hold various solutions (e.g., antibody solutions, blocking solutions) until they are needed.

Pipette Stand – Keeps them organized and out of the way.

Hand Washer – These are used to apply a wash solution if only a few plates are being processed. They are also useful if a particular solution is used only once during the ELISA protocol.

Frequently Asked Questions:

1. How sensitive are the assays?

A. The soy, egg, and milk assays all can detect the marker proteins at ≤ 30 ppb.

2. Is contamination a problem?

A. You bet! The experiments must be set up so that each step along the way is optimized to reduce the probability of contamination. For example, we have people collect the traps, place wax paper on them at the collection point, and placed in a cooler. Those people would not handle the traps again until they have showered to prevent contamination. Each insect is collected separately using a toothpick; the tooth picks are arranged so that they don’t touch, and the person needs to frequently wash their hands to prevent contamination. If the insect is a moth and large numbers are caught on the trap, the scales of a marked individual can contaminate other individuals, so you must collect the moths frequently enough, use a trap inefficient enough to prevent capture of large numbers, or use enough sticky material on the trap to minimize scale shedding. Insects that touch each other in the trap will probably both register marked.

3. Does the sticky material on the trap interfere with the ELISA reaction?

A. The different sticky materials we’ve tested do not interfere with the ELSIA reaction, except if they physically coat the specimen and actually seal the surface. We either scrape traps with a paint scraper to get a uniform thickness or use a special sticky material (“Sticky Stuff” from Tanglefoot Corp.) that is tacky to the touch, but strong enough to retain the insects. It works especially well with parasitoids and small moths because it doesn’t wick up on the insect when its hot.

4. Do the insects dry out in the traps?

A. Yes, our humidity in Washington is low and during the heat of the summer, you need to collect samples every 2-3 days to prevent them from becoming brittle. This is especially a problem when using the sticky stuff that doesn’t wick up on the insects.

5. What are the rates of the markers used?

A. We currently use 10% solution of egg whites, and 20% solutions of soy and milk.

6. Does the water used to dilute the markers matter?

A. Yes. We have shown that the water used can have a big effect on the detectability of the markers. Part of this is overcome by the high concentration of marker in the water, part can be reduced by adding certain additives (0.3 g/L EDTA for egg and milk, nothing for soy) improved assay sensitivity. We are not sure what the problem is with the water, except it doesn’t appear to be pH – it may be water hardness, organic content, etc.

7. How stable are the markers?

A. We have shown that they are stable at least 19 days for all three markers. We also have some data from an early test where we could detect the markers on apple leaves 45 days after application (during heat of summer with no rainfall). This does not mean that the residue would be picked up and detectable over that full period. We did get good marking with egg and milk over the 19-day period from insects walking across the dried residues.

8. Are all the markers equal in their ability to mark insects walking across the dried residue?

A. No – egg is clearly best, milk is roughly 2-3 fold poorer, soy rarely marks in this fashion.

9. Can you add a spreader to the markers to improve marking?

A. You can add a spreader, but it will reduce the marking to some extent. We have tested a wide variety of materials and have determined the lowest dose that actually allows wetting of the insect cuticle so that effects are minimized. However, in our residual studies we did not see markedly better marking. See the section on spreaders.

10. Are the markers rain fast?

A. Depends on how much rainfall occurs. In our lab wash-off studies, the milk marker appeared to be the most rain fast. The egg and soy were less so. If the amount of rain is low, it may improve marking by re-suspending the markers. Dew may cause the same effect.

11. If my sample has a large amount of other proteins present, can I still use the indirect ELISA?

A. The sample procedure we recommend is to leave larger insects (moths, etc.) in the extraction buffer only 3 minutes. This prevents extensive extraction of alternate proteins. You can do shorter intervals, all the markers are rapidly extracted from the insect. If you still have too much protein extracted, you can dilute the samples, especially for the egg and milk assays. We found that diluting the sample so that it was from 12.5–25% of the full strength solution was best for milk, and 6.25–12.5% of full strength was best for egg. Those dilutions still gave 100% marking fidelity. The soy marker is not improved to a significant degree by diluting the sample.

12. Can you reduce the dosage applied to the field to reduce marking by walking over a dried residue?

A. Yes. We found no marking when pear psylla walked across a dried spray solution of milk at 1%, and very little marking from soy when applied at 2%. Soy always has a low percentage marked by the residue, even when applied at 20%. Egg is very difficult to use in this fashion, even at 0.0625% solution resulted in 37.5% marking of psylla 1 day after application and when the leaves were field aged for 7 days, the marking rate was still 16.7%, much better than we get with 20% soy.

13. Have you evaluated which types of microplates work best?

A. Yes –we have standardized on Nunc Polysorp plates. The signal to noise ratio is consistently good with all three markers. However, if absolute cost is a consideration, then the untreated plates would be fine for egg and soy assays. For the milk assay, the Polysorp plates are best with Nunc Maxisorp very close; use which ever is cheaper.

14. Have you tried using powdered milk, powdered eggs, or soy flour to mark insects?

A. Yes – all the dry formulations are highly detectable and are useful in certain circumstances. The powdered milk is a bit of a problem because it has a larger grain size and might need to be run through a blender to reduce the grain size. Powdered eggs work great, but are fairly expensive in small quantities. Soy flour works great and we’re using it in one of our current projects – see Project 1 above.




Vincent P. Jones

Professor & Entomologist

Department of Entomology, Washington State University Tree Fruit Research & Extension Center, Wenatchee, WA 98801

(509) 663-8181 ext. 273 (phone) (509) 662-8714 (fax)



Links of Interest

Codling moth mating videos

Immuno-marking studies

Decision Aid System user statistics


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