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Processing and Point Mounting Chalcidoidea
by
Patrick B Beauzay'
Department of Entomology
North Dakota State University
Fargo, ND
Introduction Processing Mounting References Links
The Hymenoptera superfamily Chalcidoidea is a taxonomically diverse and economically important insect group. Currently, about 22,000 chalcidoid species have been described (Noyes 2003). Estimates on the total number of species approach 400,000 (Noyes 2000). Many species are used as biological control agents of agriculturul pests, usually as part of an integrated pest management plan. Their natural population control service results in annual savings of billions of dollars to the agriculture industry worldwide. The reduced need for costly pesticide application also has a positive environmental benefit, particularly for aquatic systems. The Chalcidoidea is a cosmpopolitan group, and there is obviously a great potential for discovery of new species that could be economically beneficial. The identification and use of chalcidoid wasps in biological control requires accurate species identification. Currently there are relatively few taxonomists who work with Chalcidoidea, especially considering the diveristy of the superfamily. These same taxonomists provide chalcidoid identification services to research and extension entomologists worldwide. Because many chalcidoid species are minute and relatively soft-bodied, it is critical for the purpose of identification that the specimens are properly prepared. This page explains and illustrates a simple protocol for processing and mounting Chalcidoidea (and other microhymenoptera) collected from Malaise trap and flight intercept trap samples. The procedures explained below should work well for any insect sample collected in a liquid killing agent regardless of trap type.
Processing Samples from Liquid Killing Agents
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| A. 500 ml, 250 ml, 125 ml sample bottles | I. 3/0, 5/0, 10/0 fine brushes | |
| B. 500 ml squirt bottle containing 80 % EtOH | J. Transfer pipette | |
| C. 2 dram vials with neoprene stoppers | K. #5 straight and #7 curved forceps | |
| D. Small glass petri dish | L. Shellac glue and applicator | |
| E. 5 ml beaker | M. Sharp dissecting scissors | |
| F. 95 mm x 17 mm glass petri dish with grid | N. #100 standard mesh sieve | |
| G. From l to r: 80 % EtOH, 95 % EtOH, 100 % EtOH, HMDS | O. Plastic funnel | |
| H. Pins, points, temporary labels, scrap cardstock | P. #4 standard mesh sieve | |
Figure 1. Processing and mounting equipment
Townes style Malaise traps (Townes 1972) are usually equipped with a 500 ml plastic sample bottle containing a liquid killing agent to collect insects caught in the trap. Water with a drop or two of non-ionic surfactant, propylene glycol, and 70 % to 80 % ethanol can all be used as killing agents. In the first two cases, samples should be thoroughly washed with deionized water and then stored in 80 % ethanol until processing. If specimens are collected directly into ethanol, the collecting ethanol should be replaced with fresh 80 % ethanol for storage. In any case, simply dump the raw sample into a 100 gauge wire mesh sieve and run water over the sample for about a minute, occasionally swirling the sample with a larval forceps to ensure thorough washing of the sample. Once the sample is washed, use running water to collect the sample into a tight bolus along the curved edge of the sieve. Next, tilt the sieve at a slight angle so that excess water can run out through the mesh and so that the sample bolus stays along the edge of the sieve. Once excess water has been allowed to drain, transfer the sample to a suitable sized plastic storage bottle. I have found that 125 ml and 250 ml bottles work well for small and medium-sized samples, and a 500 ml bottle is required for large samples. The sample is easily transferred to the storage bottle by placing a clean plastic funnel with a six inch diameter into the storage bottle, and then placing the sieve (which still contains the sample) vertically on top of the funnel. Tilt the sieve slightly forward so that the sample will dump into the funnel, and carefully transfer the sample into the funnel by squirting 80 % ethanol along the sieve mesh between the sample and the mesh (not through the mesh from the bottom of the screen). The sample bolus should slide into the funnel and into the storage bottle. Carefully examine the sieve to make sure all contents have been transferred, and then use the ethanol squirt bottle to transfer any specimens that may have become stuck on the funnel. Be sure to label the identity of the sample. It is a good practice to place one label inside the sample (written in alcohol-proof ink or #2 pencil) and affix one label on the outside of the sample bottle. With large samples, the ethanol may need to be replaced again if the sample will not be processed immediately, or if only certain insects will be immediately removed from the sample.
Removing Chalcidoidea from raw Malaise trap samples can be problematic and frustrating. Lepidoptera, particularly Noctuidae, are commonly collected in Malaise traps and shed scales making the ethanol murky so that it is difficult to discern minute insects. Also, the large surface area of Lepidoptera wings makes visual detection of minute insects challenging. Large Diptera are also problematic because minute insects can easily become lodged in stiff setae or tarsal claws. To overcome these problems, I use different sieves to separate the large insects from the small. To do this, place a #100 fine mesh sieve beneath a #4 mesh sieve and pour the sample into the #4 sieve. Place the sample under running water, occasionally agitating the sample to ensure separation of large and small insects. After doing this, place the #4 sieve still containing the large insects into a white enamel coated dissecting pan filled with water so that the water level is above the mesh of the sieve. Gently agitate the sieve by swirling and lifting the sieve in and out of the water. This ensures that any small insects still clinging to large insects will become dislodged and separate out from the sieve. Agitate for about 30 seconds and then carefully strain the contents of the dissecting pan through the #100 sieve which already contains some small insects from the raw sample. Repeat the procedure using the dissecting pan 2 or 3 times, depending on the size of the sample. You should be able to visibly determine when most of the small insects are removed because they will no longer show up in the dissecting pan. Once the small insects have been removed, they can be transferred to a clean sample bottle with 80 % ethanol using a funnel and ethanol squirt bottle as described above. The refined sample containing the small insects may contain a large number of moth scales. Gently swirl the sample and allow it to settle for about 5 or 10 minutes. Most of the moth scales should float to the top of the sample where they can be siphoned off with a turkey baster or some other siphoning device. Once most of the moth scales have been removed, Chalcidoidea are now ready to be removed from the sample.
This narrative assumes that the worker is familiar with the chalcidoid families, or the other insect families that are of interest to the worker as this method is suitable for many other insect groups that are collected in liquid killing agents. Swirl the sample and pour a part of its contents into a petri dish so that the petri dish is about half full. I use a petri dish bottom about 17 mm deep with a 95 mm diameter and with a grid on the bottom. Place the petri dish on the stage of a dissecting scope and position the dish so that the extreme left hand side of the dish is centered in the scope. I use a Nikon SMZ-1B with 10x eyepieces set at the lowest magnification. This scope is relatively inexpensive and is easy to work under. Before sorting, prepare a small glass petri dish with about 10 ml of 95 % ethanol. This dish will temporarily hold the Chalcidoidea you are removing from the sample. Begin systematically moving the large petri dish back and forth under the dissecting scope, removing any Chalcidoidea. Use a #6 or #7 fine forceps to remove the wasps. I have found that the best way to do this is to grab a wasp by a forewing. DO NOT grab the body or a leg as this will almost certainly damage the wasp. Minute and wingless individuals can be removed using a plastic transfer pipette. Transfer the wasps to the small holding dish. Once the large dish has been thoroughly examined, dump the unwanted insects into another clean bottle for later processing of other insect groups. Repeat the above procedure until the entire sample has been examined. Once the sample has been sorted, pour the wasps that are in the small holding dish into a 2 dram vial (with neoprene stopper) using a small funnel. Be sure again to properly label the 2 dram vial. I use pre-cut pieces of card stock and alcohol-proof ink for my labels. You now have a sorted sample of Chalcidoidea stored in 95 % ethanol.
Chalcidoidea, especially soft-bodied forms, should not be stored in ethanol for too long (see Noyes 1982). Specimens that are to be slide mounted can remain in 95 % ethanol until they are ready for preparation (see Platner et al. 1999). Specimens that are to be dry mounted on points or cards must be completely dehydrated to avoid collapse of body segments, particularly the head. After complete dehydration using 100 % ethanol, the chalcidoid samples can be dried using the critical-point drying method (Gordh and Hall1979) or by using hexamethyldisilazane (HMDS) following the method of Brown (1993). In both methods, the body tissues are suitably prepared for electron microscopy. Chemical drying using HMDS is much less expensive than critical-point drying and, except in the case of minute soft-bodied Chalcidoidea, gives specimens that are in excellent condition for microspcopic examination. I have had good success in preparing small genera of Mymaridae, notably Anagrus and Erythmelus, using HMDS. I have also found that specimens dried using HMDS are less brittle than those that have been critical-point dried.
To dry Chalcidoidea using either HMDS or critical-point drying, it is first necessary to completely dehydrate the specimens using 100 % ethanol. It is important that the 100 % ethanol is 100 % dry. Repeated openings of a container to the atmosphere will eventually result in partial hydration of the ethanol. To avoid this, I recommend storing the 100 % ethanol used for sample drying in a 500 ml glass bottle with about 1 inch of a molecular filter in the bottom of the bottle. This will remove any water that enters the 100 % ethanol. To dehydrate the specimens, remove the label from within the 2 dram storage vial containing the specimens and set it aside. Remove the 95 % ethanol using a transfer pipette and being careful not to remove any wasps. Add about 2 ml of 100 % ethanol, replace the stopper, and allow the specimens to soak for about 30 minutes. Remove the 100 % ethanol and repeat using fresh 100 % ethanol. Pour the ethanol and wasps from the 2 dram vial into a 5 ml beaker. Remove the ethanol using the transfer pipette. Any wasps that might remain in the vial can be removed with a #1 or smaller natural hair brush (camel or marten hair works well) or the transfer pipette. Under a fume hood, add 1.5 to 2 ml of HMDS to the 5 ml beaker so that the specimens are completely covered. For very large samples, it may be necessary to use a small petri dish with the wasps arranged in a single layer on the bottom of the dish. In this case, use enough HMDS to completely submerge all of the wasps. Allow the HMDS to evaporate under the fume hood for about four hours. Once the sample is dried, the specimens are ready for mounting or dry storage.
Dried specimens can be stored unmounted until it is convenient for the worker to prepare mounts. I recommend storage of dry unmounted material in jewelry boxes measuring 69 mm x 43 mm x 24 mm (2.5 in x 1.5 in x 7/8 in). Using boxes of these dimensions, 96 samples can be stored in a standard Cornell drawer. Once again, be sure to properly label the storage boxes.
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A. Mounted wasps |
| B. Sample box with dried wasps | |
| C. Mounting platform (Bristol board glued to corkboard) | |
| D. Nikon SMZ-1B stereoscope with fiberoptic light source | |
| E. Glue applicator (pin vise with head of pin exerted), card scrap with glue | |
| F. 3/0 and 5/0 fine brushes | |
| G. Pin vise with minuten exerted (for positioning body parts) | |
| H. Sharp dissecting scissors | |
| I. Prepared Bristol board points attached to #2 insect pins | |
| J. 80 % EtOH | |
| K. Shellac glue |
Figure 2. More mounting equipment
Dried specimens for microscopic examination should be mounted on points. I have found the card mounting method described by Noyes (1982) to be inadequate for two main reasons. Firstly, many specimens die in rather inconvenient positions to facilitate card mounting and are difficult or impossible to position properly on the card. Secondly, viewing morphological characters necessary for identification and/or description is problematic because such characters are sometimes not visible at all or cannot be properly viewed from different perspectives. Such specimens need to be soaked from the card and then remounted which increases the risk of specimen damage. Also, I have found that I break more specimens when trying to position them properly on a card. Point mounting is as efficient as the card mounting process and results in specimens with more easily observable characters. Specimens should be relaxed prior to mounting, especially if they have been critical-point dried. I have found that specimens that have been chemically dried with HMDS retain some flexibility long after they have been dried. If one is extremely careful, HMDS-dried specimens can be mounted without relaxing them first. Specimens can be relaxed in an atmosphere of acetic acid (see Noyes 1982) or in an atmosphere of ethyl acetate or Barber's solution.
I have found alcohol soluble shellac to be the best adhesive for point mounting Chalcidoidea. The glue does not dry quickly and thus allows for repositioning of the specimen once it has been mounted. It is also thick enough that the specimen will stay in position until the glue dries. Specimens can be removed from dried shellac by soaking them in chloroform. The shellac will become rubbery and peel away from the specimen (Gary Gibson, pers. comm.). Here is a recipe for shellac glue from Gary Gibson, Ph.D. of the Canadian National Collection of Insects, Arachnids and Nematodes (http://canacoll.org/Hym/Staff/Gibson/Gibson.htm):
Ingredients: 250 ml. pure white shellac; 20 ml. 70% ethyl alcohol. Boil shellac in a porcelain dish that has a pourout lip. Heat shellac to a rolling boil, stirring constantly with a glass rod while boiling for about 20 minutes or until shellac becomes foamy. Add ethyl alcohol, stirring constantly, and boil for another 5 minutes or until mixture becomes foamy again. Remove from heat and immediately pour into screw cap vials. We use 1 dram (15 x 45 mm) vials and the recipe fills approximately 22 vials. The resulting gel should have the consistency of Vaseline and be soft enough to allow movement of specimens when these are pointed. If the gel gets too thick over time it can be thinned by adding a drop of 75% ethanol and stirring the vial with a pin. The dish and rod can be cleaned using methyl alcohol.
Points used for point mounting should be punched from quality Bristol board with a small teardrop point punch. Bristol board points hold up better and are less ragged than card stock points. #2 or #3 enameled insect pins should be used.
Points should have the tip bent down at a ninety degree angle to accommodate very large Chalcidoidea such as large Perilampidae and Chalcididae (figure 3). Small to mid-size Chalcidoidea should be mounted directly to a truncated point (figures 4 and 5). This type of point is made by simply cutting off the very tip of the point with a sharp dissecting scissors. The width of the truncation should be about the width of the thorax of the specimen. Minute Chalcidoidea can be mounted directly to the tip of the point without the need of truncating the point. It is useful to prepare several dozen points before mounting a series of specimens.
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| Figure 3. Euperilampus triangularis (Say) | Figure 4. Steffanolampus salicetum (Steffan) | Figure 5. Torymus sp. |
Prepare a mounting platform by gluing or taping a 3 inch x 4 inch piece of Bristol board to a piece of corkboard of the same dimensions. This platform is placed on the stage of the dissecting microscope. Place the insect pin (with attached Bristol board point) into the center of the corkboard platform at an angle away from you, so that the point is tilting about 30 degrees upward. From this point on, the procedure for mounting the wasp is virtually the same as that described by Noyes (1982). Place the wasp to be mounted near the point in the position it will occupy on the point (figure 6). Make sure that both the wasp and the point are visible in the microscope field of view and focus on the tip of the point. Use the glue applicator to place a small amount of shellac glue on the tip of the point. The amount of glue used should be roughly equal to ½ the volume of the thorax. To mount the wasp, dab the tip of a fine brush to your tongue to gather a small amount of saliva. Touch the brush to the thorax of the wasp. The saliva on the brush should provide enough adhesion to pick up the wasp. Place the thorax of the wasp laterally into the glue. The glue will provide more adhesion than the saliva on the brush and the wasp will stay attached to the point. Make any minor positional adjustments necessary to produce a good quality mount. Once the specimen has been mounted and properly positioned, set it aside to dry for about 15 minutes. After the specimen has allowed to set, position the point in its proper place on the pin using a #4 forceps. Labels can now be placed on the pin, and the specimen is ready for examination and/or storage.
Figure 6. Perilampus sp. ready for
point mounting
Brown, B.V. 1993. A further chemical alternative to critical-point drying for preparing small (or large) flies. Fly Times 11:10.
Gordh, G., and J.C. Hall. 1979. A critical point drier used as a method of mounting insects from alcohol. Entomological News 90(1):57-59.
Noyes, J.S. 1982. Collecting and preserving chalcid wasps (Hymenoptera: Chalcidoidea). Journal of Natural History 16:315-334.
Noyes, J.S. 2000. The Encyrtidae of Costa Rica. Memoirs of the American Entomological Institute 62:355 pp.
Noyes, J.S. 2003. Universal Chalcidoidea Database. URL: http://www.nhm.ac.uk/research-curation/projects/chalcidoids/
Platner, G.R., R.K. Velten, M. Planoutene, and J.D. Pinto. Slide-mounting techniques for Trichogramma (Trichogrammatidae) and other minute parasitic Hymenoptera. Entomological News 110(1):56-54.
Links to Chalcidoidea Websites
USDA Systematic Entomology Lab, Chalcidoidea webpages: http://www.sel.barc.usda.gov/hym/chalcid.html
CNC Chalcidoidea website: http://canacoll.org/Hym/Staff/Gibson/chalcid.htm#Resources
CNC Chalcidoidea morphology website: http://res2.agr.gc.ca/ecorc/chalcid/intro_e.htm
University of California, Riverside: http://cache.ucr.edu/~heraty/index.html
Texas A & M University: http://hymenoptera.tamu.edu/
The Ohio State University world listing of chalcidoid workers: http://iris.biosci.ohio-state.edu/newsletters/cmen.html
Nomina Nearctica: http://www.nearctica.com/nomina/wasps/hymenop.htm
' Patrick B Beauzay
Graduate Studies, Department of Entomology
217 Hultz Hall, 1300 Albrecht Dr. Box 5346
North Dakota State University
Fargo, ND 58105
TEL (701) 231-9491
Send questions or comments to: patrick.beauzay@ndsu.edu
Updated: 16 February, 2006