- 1 Introduction
- 2 Insect and Arthropod Collection Content
- 3 Collection
- 4 Locality and Important Information
Methods to collect insects and other arthropods are almost as numerous as kinds of insects, and new techniques are still being developed. This page will cover the general collection of various groups of insects and arthropods, and offer specific hints and tips where available. However, collecting a specific taxon for a specific purpose may require a particular protocol which should be followed.
Insect and Arthropod Collection Content
Any collection event consists of two elements:
1) the sample,
2) locality and other relevant information concerning the sample and the circumstances in which it was collected. These are collectively referred to as data.
If one of these elements is missing then the remaining element is worthless and the entire endeavor was a wasted effort. If you have the greatest sample in the world, but don’t know when or where that sample was collected, then the sample, for the purpose of documentation, is worthless. Likewise, if you have a long and detailed account of specific circumstances surrounding the collection of a sample, but the related sample hasn’t been properly preserved, then you have wholly failed.
This may seem harsh, but people tend not to record relevant information, like locality, at the time they collect samples, thinking instead, “I’ll remember it.” But they do not remember and the effort was wasted. At other times people collect a sample, but do not properly preserve the specimens and the sample is worthless (see Preservation for more information).
Collection of insects and arthropods may be general or targeted, casual or formal, qualitative or quantitative. Typically collections are made to answer specific questions (what is here, does the number of species X change in relation to the number of species Y, how many of species Z are on 10 plants?). Collecting tools and techniques will differ based on which category or combination of categories a desired observation falls under. Some techniques used to collect specimens may not be appropriate for specific questions (e.g., qualitative collection will not allow a comparison of the density of species X among three fields).
1a. General Collection: no specific species or group of insect or arthropod is targeted. All specimens are of equal interest. Typically general collections are used to survey a particular habitat or location, for example, the community of insects living in roadside grasses next to a corn field.
1b. Targeted Collection: a specific species or group of insects or arthropods is targeted. Only certain specimens (e.g., corn rootworm) or a group (insects attracted to yellow sticky traps in an apple orchard one week before bloom) are of interest.
2a. Casual Collection: no specific intent to obtain specimens. Typically the insect or arthropod is encountered serendipitously and is collected to determine its identity or document its presence.
2b. Formal Collection: collection of specimens is intentional and often based on a specific protocol.
3a. Qualitative Collection: no attempt is made to quantify collecting effort, sample size, catch, etc. Typically only the presence or absence of a particular species or group is of interest. Collection events (samples) are not equivalent and cannot be compared beyond presence/absence of a particular taxon.
3b. Quantitative Collection: one or many of the aspects of the collecting is held constant, such as area, time, effort, etc. Collection events (samples) can be compared among one another.
Several publications offer detailed collection and preservation information:
|1||Collecting and Preserving Insects and Mites: Tools and Techniques||Schauff, M. E. (Ed.). 2001. Collecting and preserving insects and mites: techniques and tools. Update and modified WWW version of: G. C. Steyskal, W. L. Murphy, and E. H. Hoover (eds.). 1986. Insects and mites: techniques for collection and preservation. Agricultural Research Service, USDA, Miscellaneous Publication 1443: 1-103.||Excellent resource for nearly all types of collection and preservation.||Available online: http://www.ars.usda.gov/Main/site_main.htm?docid=10141 – PDF at |
|2||Collecting, Preparing, and Preserving Insects, Mites, and Spiders||Martin, J. E. H. 1977. Collecting, preparing, and preserving insects, mites, and spiders. Part 1. The insects and arachnids of Canada. Canadian Department of Agriculture publication 1643. 182 pp.||Excellent resource for nearly all types of collection and preservation.||Available online: http://www.esc-sec.ca/aafcmono.html - PDF at |
|3||A Field Guide to Insects: America North of Mexico||Borror, D. J. and R. E. White. 1998. A Field Guide to Insects: America North of Mexico. 2nd edition. Houghton Mifflin Harcourt, New York. 416 pp. [apparently a reprint of the 1970 version, but still very good]||Excellent resource for general collection and preservation. The best value of the printed resources.||Book available new and used|
|4||Borror and Delong’s Introduction to the Study of Insects||Triplehorn, C. A., and N. F. Johnson (eds). 2005. Borror and Delong’s introduction to the study of insects. 7th Edition. Brooks/Cole Publishing, Kentucky, U.S.A. 868 pp.||Good general resource for collection and preservation.||Book available new and used|
|5||An Introduction to the Aquatic Insects of North America||Merritt, R. W., M. B. Berg, and K. W. Cummins. 2008. An Introduction to the Aquatic Insects of North America. Kendall Hunt Publishing Dubuque, IA. 1214 pp.||Excellent resource for nearly all types of collection concerning aquatic insects.||Preview of the 3rd edition available at Google Books: , 4th edition book available new and used|
|6||How to Know the Immature Insects||Chu, H. F. 1949. How to know the immature insects: an illustrated key for identifying the orders and families of many of the immature insects with suggestions for collecting, rearing and studying them. Pictured Key Nature Series. WM. C. Brown Company, Dubuque, IA. 234 pp.||Excellent resource for working with immature insects. Out of print, but inexpensive used copies are abundant.||Book available used|
|7||Soil Biology Guide||Dindal, D. L. (ed). 1990. Soil Biology Guide. John Wiley & Sons, New York. 1349 pp.||Excellent resource, especially for non-insect or spider arthropods.||Book available new (on-demand reprint) and used|
|8||A Manual of Entomological Techniques||Peterson, A. 1953. A Manual of Entomological Techniques. 7th edition. Edwards Brothers, Inc., Ann Arbor, MI. 376 pp.||Excellent resource for nearly all types of collection. Out of print, may be difficult to find.||Book sometimes available used|
Mass Collection Devices
Light Trapping (ultraviolet lights (“black lights”), mercury vapor, and various others) is a general term describing the use of lights to collect insects. It exploits some insect’s behavior of traveling to a light at night. This is a well-known phenomenon, although definitive explanations of this behavior are lacking. Insects tend to be best attracted to the shorter wavelengths given off by ultraviolet and mercury vapor lights.
Several different types of light traps (see Light Trapping, right) are routinely used. Some are designed to attract, collect, and preserve insects (1–3) while others only attract insects (4). Traps 1–3 each use alcohol or soapy water to kill specimens. The sample is collected the next day, labeled and sorted. Traps 1 and 3 are homemade and operate from a 12 volt battery. Trap 2 is commercially available and can be purchased as an AC or DC unit.
Trap 4 is a light sheet. This particular example is homemade but contains the basic design elements. The white sheet reflects light and acts as a structure for insects to land on. The sheet can be hung on a tight rope if trees/structures are available. Keep the hanging sheet tight (unlike what is illustrated in the figure) and do not bump it. A sheet on the ground is important as many insects will accumulate there. Mercury vapor (top) and ultraviolet (hanging) lights can be used together or in concert. Insects at light sheets are easily collected with aspirators (see below).
Different groups of insects tend to come to light traps at different times of the night, for example bark beetles (Scolytinae) tend to arrive in the early evening, while saturniid moths (Saturniidae) may not arrive until 1 or 2 am. The best strategy is to turn the light on at dusk and keep it on until at least midnight.
Flight Intercept Traps
Flight Intercept Traps: FITs (also known as barrier traps) are passive traps that are used to collect insects flying through an environment. Many insects when encountering a barrier while flying will either fly up to go over the barrier, or drop down to the ground. This behavior can be exploited to collect insects. Ground level FITs consist of a vertical barrier, collection containers and a horizontal rain fly (see Flight Intercept Trap, right). The vertical barrier is typically made of fine mesh, such as mosquito netting, although it could be made of clear plastic or glass. The rain fly is typically plastic. It obviously functions to keep rain from flooding the collection containers, but also functions as a top barrier to reduce escapes, thus enhancing trap success.
Collection containers are filled with a killing agent and/or preservative agent. Water, with soap to act as a surfactant, may be used as a killing agent, but the traps must be serviced and specimens removed daily, otherwise specimens will begin to rot. Propylene glycol based antifreeze (or pure propylene glycol) can be used as a killing and preserving agent. DO NOT USE ethylene glycol antifreeze because of its toxicity to mammals (the sweet taste also increases the likelihood of mammals disturbing the collection containers). If propylene glycol is used, traps can be left for long periods without servicing. Ground level FITs collect an enormous number of specimens over time (100+ per day), and will collect numerous non-flying ground-dwelling insects and arthropods as well.
A Malaise Trap is a type of FIT with the collection container at the top (see Malaise, FIT Combination, right). Generally the vertical barriers form an “H” shape when viewed from above, and the rain fly is higher on one end to act as an inverted funnel for insects that fly up when encountering the central barrier. A collection jar is placed at the peak of the trap. Specimens in the top collection jar are generally protected from the elements and can be killed with aerosols (and left to dry), or alcohol or other liquid preservatives. Generally Malaise style traps are used to collect flies and wasps, which tend to fly up when encountering a barrier, but they are effective for a wide variety of insects.
Pitfall Traps are used to collect ground dwelling insects and arthropods. The concept is simple; a container with preservative is placed in a hole in the ground and organisms fall in. The best design is a double cup design (see Pitfall Trap, right). After the hole is dug, two cups (typically robust disposable plastic drink cups, the expensive brands) are placed in the hole and soil packed around it to the level of the lip of the inner cup. The inner cup can now be removed and emptied while the outer cup keeps the hole from back filling with soil. It is important that the lip of the inner cup is level with the surrounding soil, otherwise small organisms may see it as a barrier and go around the cup or burrow beneath it.
An elevated cover is placed over the pitfall to keep out water, falling debris, and partially protect against curious mammals. The cover only needs to be ~10 cm (3 inches) above the cups. Covers can be made of almost any material and if the pitfall is not meant to be left out for very long a disposable plastic plate held up with kabob skewers will work. A drift fence, often made with aluminum flashing, can be used to direct individuals to the pitfall.
Install pitfalls using a bulb planter when the soil is moist, even if this is months before any collecting will be done. Wet soil increases the ease of installation, and also allows dirt to be packed up against the lip of the inner cup. When pitfalls are not in use, remove preservative, and either cap the pitfall or place several sticks in it to allow escape of organisms that fall in (you do not want to have to deal with a two week post-mortem mouse drowned in rain water).
Sifting and Berlese Funnels
Sifting and Berlese Funnels are used to extract small insects and arthropods from leaf litter and other debris (dung, mushrooms, flood debris, etc.). A sifter is a device used to concentrate individuals by removing large pieces of substrate. This is done using a container with a wire mesh bottom. The debris is placed in the container, agitated, and the smaller particles and individuals fall through (see Sifting, right). If you sift over a white pan (see below) or white sheet you can immediately begin collecting specimens with an aspirator or brush (see below).
A large amount of sifted debris can be collected if you place a cloth bag, such as a pillow case, under the sifter. NEVER use a plastic bag. Carbon dioxide buildup in sealed plastic will kill all the organisms. If you must use plastic bags, cool the sample quickly and/or process it within a few hours to minimize this effect. Always put a label in the bag with the sample. Keep the bag cool and moist until you are ready to extract specimens with a Berlese Funnel. Samples in cloth bags will remain viable for a week or longer if kept cool and moist.
A Berlese funnel uses a heat source, typically a light, to warm and dry a sample from above. The sample is placed on wire mesh and suspended over a collection container. As the top of the sample warms and dries individuals migrate down eventually falling into the collection container. Take care to not overheat the sample. Drying should take 1.5-3 days depending on the size of the sample. Alcohol (80% ethanol) is a good preservative to use in the collection container. Always put a label in the collection container. If live specimens are desired, place a moist napkin in the collection container and check often.
Bait Traps, using fruit, dung, carrion, pheromones, heat, color, etc. as attractants can be general or targeted collection devices. Many bait traps are homemade, while others are available commercially.
The general design of a bait trap includes two elements (see Bait Traps, right):
1) A place to hold the bait: Attractants such as pheromones or colors can be incorporated in the structure of the trap, while fruit, dung, carrion, etc. typically need a cup or compartment in which to be placed.
2) A means of killing and preserving the specimens: If the killing/preservative agent is liquid a cup or basin is needed, typically this is separate from the compartment used to hold the bait. Another common killing agent, especially for traps that use color or pheromones, is the use of a sticky substance, such as “tanglefoot”. This is not a preservative, and “sticky traps” should be checked often if quality (or even identifiable) specimens are desired, and special solvents must be employed to remove the sticky material from the specimens.
Nets: Nets used to collect insects and arthropods fall into three general categories; aerial nets, sweep nets, and aquatic nets (see Nets, right). Each is specifically designed to interface with the environment in which the sampling will take place. Nets consist of three elements, the handle, the hoop, and the bag.
Note on the use of nets: Many insects are vigilant. If you intend to sample with a net do so with competence, do not blunder around, cast a shadow willy-nilly, shake branches and vegetation, or stomp around in the water or on the bank before collecting. Otherwise your intended specimens will fly/swim away, drop to the ground, or otherwise disappear long before you take a single swipe.
Aerial nets are used to collect flying insects such as mayflies, dragonflies, butterflies, moths, true flies, bees, and wasps. Aerial nets are deliberately built light, with long thin handles, sometimes flexible hoops, and a soft bag with a large mesh size. These are ideal for collecting delicate specimens, such as mayflies or butterflies. An aerial net should never contact any solid things, but only “scoop” the insect from the air. The bag is long so it may be flipped over the hoop trapping the specimen after a catch. Generally specimens collected in aerial nets are removed from the net with the hands or transferred from the net to a killing jar (see below). Aerial nets should NEVER be used for beating vegetation or in the water (unless you absolutely have to. The only thing worse than ruining a fine net is not collecting a rare specimen).
Sweep nets, or beat nets, are used for sweeping vegetation or beating dead limbs, etc. They are very robust, with short thick handles, heavy inflexible hoops, and have hardy bags of very small mesh or light canvas cloth. The durability of the net allows the user to assault the substrate with such force that insects are jarred loose and collected in the net. Sweep net samples can be “quantified” if the area swept per sweep and number of sweeps per sample are held constant. Generally specimens are collected from a sweep net using an aspirator. If small flies, wasps, or other fast flying organisms are desired, the entire sample can be dumped into a plastic bag or other container, the specimens can be killed, cooled, frozen, etc. and dealt with later. Sweep nets can be used as aquatic nets in a pinch, although sweeping with a wet bag can be difficult.
Aquatic nets are used to sample aquatic environments ranging from lakes, ponds, rivers, streams, abandoned tires, puddles, tree holes, etc. Aquatic nets range from a standard size, with a ~1.5 m (5 ft) long handle, to diminutive sizes, such as aquarium nets with 5 cm (2 inch) wide hoops. Standard sized aquatic nets are robust and can be used somewhat forcefully, but be careful not to rip the bag on hidden underwater debris. Generally hoops of aquatic nets are flat on the distal (away from the handle) edge to facilitate collection against the benthos (bottom) of the water column. Do not use an aquatic net like a shovel; scooping too much muddy substrate, gravel, or aquatic vegetation will make separation of specimens from the sample difficult or impossible. After you have collected a sample, it can be “washed” by bobbing the sample in the water, making sure that the lip of the net doesn’t go below the water and allow specimens to escape. Samples placed in a water tight container and kept cool will retain live specimens for hours. Otherwise samples should be preserved in 80% ethyl alcohol (with replacement 24 hours later if the sample was particularly watery) or can be sorted immediately. Sorting an aquatic sample in the field is best done using a white pan (see below) but care should be taken that small or cryptic organisms are not overlooked.
Beat Sheets: Beat Sheets are an important way to survey specific plants for insects (see Beat Sheet, right). Foliage should be struck soundly with a stick so insects are jarred loose and fall onto the sheet. Rather than holding the beat sheet horizontal, tip the far corner up, insects will walk upslope and are less likely to immediately fly away. Specimens are easily collected with an aspirator.
Aspirator: An aspirator (also known as a “pooter” and more colloquially as a “sucky uppy thing”) is a small device used to collect specimens (see Aspirator, right). A collecting vial is sealed by a stopper that has two tubes. The fixed metal tube is the collection tube and the flexible yellow tube is the suction tube. Notice in the figure’s insert that the opening to the suction tube is covered with very fine mesh. This is important. Small insects are sucked through the collection tube and trapped in the collection vial when suction is applied to the suction tube. The fine mesh over the suction tube keeps the specimens from being eaten by the collector. If you use a collection vial with a lid, then multiple collections can be kept separate by capping one vial and getting another.
The collector does not need to fully inhale in order to create proper suction to capture a specimen, in fact inhaling through the device is not recommended. A “kissing” technique may be used where air is taken only into the mouth and not into the lungs. Use caution aspirating near fungal spoors, dust, or dead animals. Some people attach a new car gasoline in-line filter to the end of their suction tube to keep from inhaling dirt and debris.
Aspirators can be used for any and all types of collection and should be available whenever one is in the field.
DO NOT put any chemicals in the collection vial, such as ethyl acetate or alcohol, otherwise you will be huffing the fumes.
DO NOT capture spiders (unless you are specifically after them). Spiders, even small ones, lay a trail of silk as they move around the vial. Other specimens will get caught in this and untangling specimens without damaging them is difficult.
DO put a little bit of napkin in the collection vial. It soaks up moisture and gives the specimens something to hold on it.
Your Collection Kit should consist of everything you need to have a successful collecting experience. Whenever you are collecting there are a minimum of items you should have available, either immediately in the field, in the car, or at the field house. Below is a list of recommended materials needed to collect insects. Other materials than may be important when in the field, such as sun screen, water, a bear whistle, etc. are not listed but should be taken.
I. Take the appropriate Traps, Nets, Sifter and cloth bags, and an Aspirator.
II. Midsized Materials (see figure, right):
1. White Pan: a very important piece of collection equipment. Fungi, limbs, grasses, crops, aquatic samples, etc. can be beaten, shaken, or tossed into a white pan and specimens can be collected immediately, usually with an aspirator or fine tipped paint brush.
2. Camera: important for documentation of specimens, samples, sample area, damage, etc. See How to Properly Photograph and Submit Images for Digital Diagnostics.
3. FIELD NOTEBOOK: This is where you will record, in detail, important information pertinent to your collections/observations.
4. Kill Jar: used to kill specimens that should not or cannot be killed in alcohol. Be sure your kill jar is labeled “Poison.”
5. Crow bar, or large screw driver, etc.: used to access difficult to collect locations, such as under bark, in logs, behind walls, dry dirt, etc.
III. Small Equipment: A general all-purpose small equipment kit should be available either immediately in the field (in your backpack) or in the vehicle, at camp, etc. It contains supplies that might be needed for any given collection opportunity.
The figure Small Equipment (left) illustrates the minimum small Equipment needed for the field.
1. Label paper
2. Fine tip ink pen with archival quality ink that will not run in alcohol
3. Larger permanent marker
4. Snap blade knife
6. A) labeled ethyl acetate vial to recharge kill jars; B) labeled vials with 80% ethyl alcohol; C) empty (dry) vials
7. A) vial with safety pins; B) vial with precut labels; C) vials with extra lead and erasers if you use a mechanical pencil
8. Assortment of insect pins sizes 1–4
9. Insect pinning block
10. Syringe with removable tip to inject large specimens with alcohol (e.g. moths)
11. Extra aspirator vials with lids
12. Hand lens, 10x magnification
13. A) fine tipped forceps; B) larval (soft) forceps; C) fine paint brush (can be used to collect small insects); D) tools specific to equipment (here, an Allen wrench used to remove sweep net hoop from the handle).
14. Collection Kit storage container.
IV. Miscellaneous Containers: You never know what you may encounter in the field and wish to bring back to the lab: a few mites, an entire plant, something that must be kept dry, something that must remain wet, a specimen you want to keep alive, something fragile, something smelly, etc. Always have a bunch of containers at the ready. They should be: large, small, pliable, rigid, able to seal in liquids, waterproof, etc. Many small containers three fourths full of with 80% alcohol are recommended.
Locality and Important Information
The absolute minimum information that must be associated with samples is an unambiguous location and the date. Feel free to record any other information that you think might be important regarding the sample.
The easiest way to maintain high quality locality information is to enter these data into a field notebook. This will save hours of fumbling around, trying to remember the difference between this field or that field, and will greatly reduce the chances of making a mistake. Incorrect information can be more costly than no information at all. You can even take a photograph of the information you recorded in your field notebook for easy reference.
LOCALITY: Record the locality in an unambiguous way. Coded information such as “Field B”, “Plot 32”, “E3f2” or any other field numbers are, by themselves, unacceptable. Remember specimens from your samples might get sent to an expert in New Zealand and/or it may be a useful record for years to come, long after the coded hieroglyphics have lost their meaning.
2) County, Parish, Borough or other local political unit
3) Distance and direction to the nearest “permanent” named place (e.g., town) that can be easily found on a map (not someone’s farm, a grocery store, or “the giant oak tree.”)
4) Latitude and longitude
5) Your name: to get credit and so people can contact you if necessary
6) Add information specific to the time and place, such as: “edge of newly flooded rice field,” “near ditch in corn field.”
If you don’t have a GPS device immediately available you can still obtain latitude and longitude information very simply. Using www.google.maps.com  find the location where you took the sample. Make sure you have found the correct location! Place your mouse pointer over the location and click the right mouse bottom. On the menu that pops up, click “What’s here?” The latitude and longitude will be displayed on the search bar at the top of the page: e.g., “30.41085,-91.177362”. The first number is the latitude, which ranges from 0 at the Equator, to 90 degrees at the North Pole, and -90 degrees at the South Pole. The second number is the longitude which is 0 at the Prime Meridian (a line passing from the North Pole through the town of Greenwich, England to the South Pole). Longitude moves from the Prime Meridian 180 degrees to the east, and -180 degrees to the west (both meet in the middle of the Pacific Ocean). Most of the contiguous United States falls within 25 to 50 degrees (North) latitude, and -65 to -125 degrees (West) longitude. You must add West or “W” if you report longitude as a positive number.
DATE: Writing a date that can be correctly interpreted is important. This may seem simple but it is not. For example what is the following date: 3/7/04 ?
July 3rd 1804, 1904, 2004, 2104, etc.?
March 7th 1804, 1904, 2004, 2104, etc.?
The standard arrangement of the day and month differs among people and over time. This can be a real pain if you are doing genealogical research or looking at specimens of plants or insects (that could have been collected over the last 100+ years anywhere on Earth by people of many nationalities). Just because you and all your friends write Month/Day/Year doesn’t mean someone else will read it that way.
The most unambiguous way to write a date is to give the day in numeric form, followed by the month written in letters, abbreviated or in full, and finally the full four digits of the year. Placing the month in the middle has the added benefit of separating the two elements of the date that are made of numerals.
7 November 1913
3 Jan. 1941
2 July 2018
It is also permissible to use Roman numerals to represent the month. This may be more appropriate if someone other than an English speaker will be using the data.
7 XI 1913
3 I 1941
2 VII 2018
Field and Sample Labeling
NEVER WRITE LABEL INFORMATION ON THE OUTSIDE OF A CONTAINER:
NOT ON THE LID,
NOT ON THE SIDE.
All label information should be written on label paper with heavy pencil or indelible ink and placed INSIDE the sample, even if the sample is fluid.
NEVER WRITE LABEL INFORMATION ON THE OUTSIDE OF A CONTAINER!
PERMANENT MARKER IS NOT PERMANENT!
Label information written on the outside of a container WILL become smudged, blurred, or even erased by alcohol. Labeling caps of vials is UNACCEPTABLE, because if caps are removed from two or more vials all label information becomes suspect.
Example: You spent a week and a good deal of money collecting insects and traveling in another state and jotted down the locality data on the outside of the alcohol vials using your permanent marker. You returned and upon unpacking your samples, discovered that one of the vials was broken, the spilled alcohol is dyed a rich cobalt blue, and all the information on the vials has dissolved. You just lost all your samples!
NEVER WRITE LABEL INFORMATION ON THE OUTSIDE OF A CONTAINER!
First Detector Entomology Training Project