Life Cycles, Collecting & Identifying
Aquatic "Bugs"

Insect Life Cycles

Insects have an exoskeleton, or rigid skin, rather than an internal skeleton like humans. Since the exoskeleton cannot stretch appreciably to accommodate increases in size larval insects grow in stages referred to as instars. Between each instar they must moult the old exoskeleton away by first absorbing as much material as they can from it then swallowing water or air until the old skin breaks open and the larva crawls out. The old exoskeleton left behind is called an exuvium (plural, exuviae). The larva will continue to swell until the new skin hardens thus ensuring room for a period of growth. Only larvae go through this process. Once the adult emerges from the final larval skin growth ceases.

In general the life cycles of aquatic insects are of two types referred to as either hemimetabolous or holometabolous.

Hemimetabolous insects include mayflies, dragonflies, damselflies, stoneflies and true bugs. In this type of life cycle after hatching the larva (sometimes referred to as "nymphs") develops through a series of instars until maturity. After each successive moult between instars the external wing buds become larger. In the final moult the adult emerges with fully developed wings. The above image is of a series of larvae and the adult of the Giant Water Bug (Lethocerus americanus). Note the increase in size and similarity to the adult the larval stages have. The adult is sexually mature although usually some period of time before mating occurs for the teneral adult to feed, disperse, set up territories and/or develop adult color patterns. Mayflies are unique in having a subadult (subimago) emerge from the last larval moult. This stage, although it can fly, has cloudy wings and is not sexually mature in most cases and requires a further moult to reach sexual maturity.

Holometabolous aquatic insects include the caddisflies, beetles, spongillaflies, alderflies, true flies, wasps and moths. In this type of life cycle the larval growth is similar to hemimetabolous larvae but wing pads and other adult features are not present. The larva remains relatively unchanged in appearance other than an increase in size. There are also usually a definite number of larval instars. The last larval instar moults into a pupa. In this stage major restructuring of the body occurs to reach the adult form. Once this tissue development is complete the fully winged adult emerges. Above are the life stages of a mosquito.

Collecting Aquatic "Bugs"

Collecting aquatic insects and macroinvertebrates requires only a limited amount of equipment:

  • rubber boots, hip or chest waders
  • a long handled net or kitchen sieve
  • a shallow white pan or bottom of a white plastic pail
  • an assortment of small liquid proof vials and jars
  • forceps
  • "pooter"
  • field notebook
  • 90% to 100% alcohol for specimens collected from water
  • 70% to 80% alcohol for specimens collected in aerial sweeps
  • field bag to carry things such as the vials, forceps, notebook etc.

Other accessories:

  • special aerial net
  • a camera to record pictures of the site
  • GPS unit for long/lat info
  • thermometer
  • fine meshed aquarium net
  • turkey baster and/or eye dropper

These supplies can be made or purchased in various forms from retail hardware and drug stores or biological supply companies.

It is wise to go collecting with another person or a group. Not only does this provide a safety factor because slimy rocks are slippery but it also enables the collecting to be done more efficiently.

To make collections in ponds and along lake shores the net is passed through the weeds and open water. In streams and rivers the net can be held downstream of the feet and the feet shuffled to disturb pebbles and small stones. Macroinvertebrates on the stones will fall off and be carried by the current into the net. The net is then swirled in the water to remove fine silt and mud. The contents of the net are then dumped into the pan with about 2 cm of water in it. The macroinvertebrates will be seen crawling out of vegetation, sticks and pebbles and can be easily picked up with forceps or a small net and transferred to a vial or small jar with preservative. Macroinvertebrates can also be collected from submerged logs and stones by hand.

I find one of the most useful collecting tools for shallow water is an ordinary kitchen sieve about 20 cm in diameter. The kitchen sieve can be swept through the vegetation, mud and gravel with no damage. I carry a hinged lid condiment container filled with alcohol and a pair of forceps tied around my neck with a string. I find that I can sweep the sieve through the water and quickly examine the material collected. When I find interesting specimens I open the lid of the container and put the specimens in the alcohol and snap the lid back on. Periodically I transfer the specimens into a larger leak-proof sampling jar.

Aerial net sweeps of the shoreline vegetation will collect adult aquatic insects and swarming adults. The net contents are examined and the insects removed using a pair of forceps dipped in alcohol or a "pooter"/aspirator. Pooters come in a variety of styles. In the simplest form a tube is put in your mouth and the insects are sucked up. A fine screen prevents the insect from being sucked into your mouth. In pooter A the insect(s) is sucked into a tube. Your finger is placed over the collecting end to keep the insects from escaping and then the insects are blown into a collecting jar. Another pooter type (B) sucks insects into a jar which can be be replaced when full. These methods are not very sanitary and can result in allergic reactions. An healthier and safer alternative is to attach a turkey baster (C) to the sucking end of pooter A. A modification to the B style pooter is to incorporate a one way flow rubber bulb (used for blowing dust of camera lenses) into the desgin. Cut a small diagonal slot to insert the blowing end of the bulb into the tube. Squeezing the bulb creates suction in the collecting tube which draws the bug into the jar. Putting a small funnel on the collection end of any pooter will increase the capture effeciency.

caddisfly & mayfly larvae on rock

Examining branches and rocks along the shore, snags in the water or taking sweeps of the water surface with a fine-meshed net will collect exuviae. Adult insects can also be collected directly from the water which they emerge from using various types of emegerence traps. This method not only provides many specimens in good condition but also information on the habitat/microhabitat the insect lives in, the timing of the adult emergence and if the collecting area of the trap is known, a rough estimate of the numbers emerging.

Many adult insects are attracted to lights, especially black lights, so checking lighted windows at night or looking around street lights can result in many interesting specimens. Of course biting flies, most of which have aquatic origins, can be collected from one's person as they attempt to feed using forceps or a pooter.

For each collection a label should be made indicating the location, date, and collector's name. Resistall paper seems to work well in alcohol and does not deteriorate. I have found an HB Pencil or Pigma art pens are alcohol permanent. If mass produced labels are needed it is best to test the printer ink in alcohol for a number of weeks to ensure it does not run or fade.

It is useful to make notes in a field note book describing the method of sampling and the sample location. Things like time, weather, water and air temperature, and other measurements if taken, and biological and habitat observations can all provide valuable information for future reference. Photographs of the habitat also provide useful information.

bug vials

The alcohol should be changed about 24 to 48 hours after sampling to ensure the specimens will be thoroughly preserved and not start decomposing. If the jar has a large number of specimens or large specimens in it the alcohol may have to be changed a number of times to ensure proper preservation. Try to keep the samples in a cool place

The next step is to sort the specimens into groups and begin to identify and study them. Small vials (2 to 3 drams in size) are ideal for storing sorted macroinvertebrates. Screw capped vials with plastic inner seals or neoprene/rubber stoppered vials work well. (Corks should not be used as they inevitably allow the alcohol to evaporate.) The sorted macroinvertebratesare preserved in 70% alcohol so they will not become brittle. A label of the location, collection date and name of collector is included in each of the vials with the name of the specimen if it has been identified. Try a store collections in a cool place and in a cabinet out of direct sunlight.

pinned insects

Alcohol is a good all purpose preservative for most aquatic macroinvertebrates, although there are many, some extremely vile, concoctions that have been devised for particular groups. Ethyl alcohol (ethanol) seems to be a better preservative than isopropyl (proponal). (Note: All alcohols should be used in a well ventilated area to avoid inhaling the fumes. Methanol should NOT be used for health issues.) All alcohols cause colours to fade which makes it less than perfect for some groups such as adult dragonflies and damselflies. Also, adult mosquitoes and moths have their wings covered in scales which will wash off in alcohol so these groups should never be placed in liquid. For these groups pinning is a better way of preserving the specimens. The captured specimens are placed in a killing jar (This is an air tight jar that has absorbent material in the bottom with a small amount of ethyl acetate (finger nail polish remover) poured on it.) until they die. Another killing option for insect samples is to place them in a freezer over night. This kills the specimens while keeping them in good condition. Once killed the insects are sorted and an insect pin is placed through the thorax using a pinning block and then is placed (positioned if needed) on a styrofoam board until dry. A label is attached to the pinned specimen for reference.

No matter what method of preservation used the collections should be examined periodically to ensure the alcohol has not evaporated from the vials and that pinned specimens have not be infested with dermestid beetles which can reduce a pinned collection to dust in short order.

Taxonomic Rearing: A very worthwhile and rewarding activity is the "rearing" of immature insects. Only a small percentage of aquatic insects in the province can be identified to species in the larval stage so they have to be associated with the adult stage to be identified.

For this procedure mature larvae, or pupae, are collected and kept alive in a container of water; (one specimen/container). Rearing containers can be as simple as plastic beverage cups with a loose fitting lid or pill bottles to more elaborate trays with mesh sided cages and aerators to provide oxygen and current to the larvae. Each group of aquatic insects requires different rearing conditions to successfully complete their life cycles. For some groups the larvae may need to be fed. With luck the larva will develop to maturity and the adult insect will emerge. The larval and pupal skins and the adult stage can then be preserved together and become a valuable record of the association as in the case of this successfully rearing of a chironomid (non-biting midge).

Classification and Identification

When people formally identify a plant, animal or insect, they are going beyond just giving it a name. In reality, the specimen is being placed in an hierarchical classification of living things. Humans classify everything in the world. For example, books can be divided into fiction, non-fiction, etc. Each of these groups can in turn be further subdivided (romance, comedy, self-help and so on). The same has be done for living organisms. Initially classifications were for practical purposes: edible, inedible, poisonous, dangerous, medicinal, etc. As biology developed, similar looking organisms were grouped together. With the synthesis of ideas in the 18th and 19th centuries by workers such as Linnaeus, Lamarck, Wallace, Darwin, classifications began grouping organisms together that shared a common evolutionary heritage (common ancestry). The process of developing this "natural" classification continues to evolve as new methods such as DNA analysis are developed to supplement morphological, anatomical, behavioural and ecological information.


As can been seen from the table above there are seven major levels of classification. At the finest level is the species name. Organisms are usually referred to by their genus and species name together. Always written with the genus capitalized, the species name lower case and in italics, Aeshna eremita. These two names (binomial name) together are unique to that organism. Often included with the genus species name are the author, the person who is credited with first describing the organism and the publication date for the description. For example the full citation for the Lake darner would be, Aeshna eremita Scudder 1866. If there are parentheses around the author's name this indicates it was originally described as belonging to another genus and a subsequent worker assigned it to its current place in the classification scheme. There are very strict rules for naming new organisms and assigning organisms to new groups.

For the general public naming aquatic "bugs" does not go much beyond common names (dragonflies, mosquitoes, beetles, blackflies, snails, clams, etc.). These names may refer only to groups and vary in their precision. For example, dragonflies refer to the suborder Anisoptera of the order Odonata (Dragonflies and Damselflies). There are about 45 different species of dragonflies reported from Saskatchewan. Mosquitoes refer to the family Culicidae with 47 types present in the province. The term beetle refers to the order Coleoptera which has over 230 aquatic representatives in Saskatchewan. Blackflies refer to the family Simuliidae, which has 32 species recorded from the province. Snails and clams are two classes (Gastropoda and Pelecypoda) in the phylum Mollusca. There are over 30 aquatic species of each in Saskatchewan. And so on....

"The species is the thing": Typical generalized identifications, such as those above, carry little real information regarding life history, ecology, and environmental requirements, as these vary, sometimes significantly, among the species of the "group". It is the species level identification that carries the detailed information regarding life cycle and biology that researchers use to communicate, develop experiments and test hypotheses. Accurate species identifications and ecological information is especially critical in the case of insects that carry diseases or are pests. For example, not all mosquitoes feed on humans and mammals, some feed on birds, others will feed on both birds and mammals and still others feed on amphibians and reptiles. Only certain species carry West Nile Virus and are able to transmit it to humans. To monitor the potential of disease spread and develop effective control measures it is vital to be able to identify the species, know its life history, its preferred breeding habitats, feeding behaviours and its general ecology. Furthermore, biodiversity studies require species level work in order to accurately document the faunal (and floral) richness of areas or regions of concern. And, the monitoring of invasive species, as the name implies, requires being able to identify the specimens in question to the species level in order to know its detailed biology so the invasion may be prevented, or if the invasion as already occurred, controlled effectively.

The species category is commonly the most detailed level of classification used in biology. Species information in itself is a generalization of life history, biology, ecology and distribution data of populations that make up a potentially interbreeding group. It is the basis for all generalizations of higher levels of classification. In many instances species within the same genus have significantly different life histories, biologies, and environmental requirements. Without first determining the species in a study it is impossible to assess the accuracy of genus or family level reporting. Therefore, studies at the supraspecific level have their accuracy fundamentally compromised especially in cases that involve a number of multispecies genera. Unfortunately, genus and family level analysis without supporting species level data seems to be the current vogue of aquatic ecology and impact monitoring even though the value of genus/species level taxonomy can provide significant additional information (Lenat and Resh 2001). Supraspecific research (family and generic level studies) not only affects the quality of data available to researchers and decision makers but has also curtailed the development of aquatic insect systematics and species level biodiversity research.

Lethocerus americanus

To identify some aquatic macroinvertebrates to species can be as simple as comparing pictures in a guidebook similar to "birding". If the specimen looks like the "Giant Water Bug" and is over 7 cm long and it was collected in Saskatchewan then you can be reasonably sure it is Lethocerus americanus as they are so distinctive in shape and size.

Unfortunately, easy species identifications like the above are rare. In most cases the process of identifying an aquatic macroinvertebrate to species is an arduous task that requires a great deal of knowledge and years of experience to do correctly and effectively. As E.O. Wilson wrote in his book "Naturalist" (Island Press, Washington D.C. 1994) "[Taxonomy] is a craft and a body of knowledge that builds in the head of a biologist only through years of monkish labor..... If a biologist does not have the name of the species, he is lost. As the Chinese say, the first step to wisdom is getting the right name."

The typical process of identification is to "run" a specimen through a dichotomous key that uses choices of character states to work through a series of couplet statements to reach an identification.

For Saskatchewan four good taxonomic keys are:

Merritt, Cummins and Berg (2008), An Introduction to the Aquatic Insects of North America. 4th Ed. Kendall/Hunt Publishing Co. Dubuque, Iowa.

Clifford (1991). Aquatic Invertebrates of Alberta. University of Alberta Press. Out of print.

Smith, DG. 2001. Pennak's Freshwater Invertebrates of the United States. 4th ed. John Wiley and Sons.

Thorp, JH and AP Covich. 2001. Ecology and Classification of North American Freshwater Invertebrates. 2nd ed. Academic Press.

The above texts are not designed for Saskatchewan exclusively so there are many families, genera and species included that do not occur in the province and there are a significant number of macroinvertebrates found in Saskatchewan that are not covered in Clifford's Alberta text. This last point is important as regional keys often are able to use shortcuts because certain taxa are not found in the particular region and therefore specimens worked through the key may be incorrectly identified. Many workers do not fully appreciate that rather than these texts being the end of the identification process they are actually only a good starting point to the identification process.

Even with the proper literature assembled identifications can still be difficult and frustrating because more often than not the specimen is damaged and critical parts are missing. For insects, most larval keys are designed for final instar larvae and will not work if the specimen is an early instar. This is particularly important when lengths of the body, or a body part, or a ratio of one part to another, are used as the distinguishing character. Often the adult stage is required for species level identifications. For many insect groups the adults, if not aquatic, are not collected in regular aquatic sampling and special effort and techniques must be used to collect them. Furthermore, without positively associating the larval stage with the adult stage it can be impossible to identify the larvae to species by itself.

Identifying many macroinvertebrates to order or family level can often be done with the unaided eye or a simple 10X hand lens. However, for many family, genus and species identifications some form of stereoscopic dissecting microscope capable of magnifications of 7 to 40 times is extremely useful. For most groups no elaborate specimen preparation is needed but other groups, such as the non-biting midges (Chironomidae), microdissection, clearing of tissues and mounting on microscope slides to study them under high magnifications with the aid of a compound microscope is required.

The above information is primarily related to the traditional morphological methods of identification where structural differences are used to distinguish taxa. However, in many cases, interspecific morphological differences may be very subtle in closely related species to the point of two (or more) species being essentially indistinguishable in all stages. The only suspicion of the presence of multiple species may be from distributional, ecological or life cycle differences and "gut feelings" of taxonomists. The use of DNA barcoding has proven to be quite promising in splitting such species groups and discovering unknown species within these groups (Ball etal 2005, Pfenninger etal 2007, Zhou etal 2009, Webb etal 2007). However, barcoding does have its disadvantages as it requires much more than just a microscope. So at present barcoding is essentially only really practical for taxonomic research, although preliminary attempts are starting to explore its use in ecological and monitoring researches (Pilgram 2011).

Once an identification is made, no matter what the procedure used, a series of specimens should be either critically compared with specimens previously verified by an expert or sent to a taxonomic expert to verify the identification. Unfortunately, in light of over 30 years of poor funding for taxonomic work, experts are very few in Canada.


Ball SL, Hebert PDN, Burian SK, Webb JM. 2005. Biological identifications of mayflies (Ephemeroptera) using DNA barcodes. Journal of the North American Benthological Society 24: 508-524.

Lenat, DR and VH Resh. 2001. Taxonomy and stream ecology: the benefits of genus- and species-level identifications. Journal of the North American Benthological Society 20:287–298.

Pfenninger, M, C Nowak, C Kley, D Steinke and B Streit. 2007. Utility of DNA taxonomy and barcoding for the inference of larval community structure in morphologically cryptic Chironomus (Diptera) species. Molecular Ecology 16:1957–1968.

Pilgrim EM, SA Jackson, S Swenson, I Turcsanyi, E Friedman L Weigt, and MJ Bagley. 2011. Incorporation of DNA barcoding into a large-scale biomonitoring program: opportunities and pitfalls. J. N. Am. Benthol. Soc. 30:217–231.

Webb JM, Sun LL, McCafferty WP, Ferris VR. 2007. A new species and new synonym in Heptagenia Walsh (Ephemeroptera: Heptageniidae: Heptageniinae) based on molecular and morphological evidence. 16pp. Journal of Insect Science 7:63.

Zhou X, SJ Adamowicz, LM Jacobus, RE DeWalt and PDN Hebert. 2009. Towards a comprehensive barcode library for arctic life - Ephemeroptera, Plecoptera, and Trichoptera of Churchill, Manitoba, Canada. Frontiers in Zoology 2009, 6:30.