Appendix I: Special Considerations in Field Sampling and Laboratory Processing

Fixatives and Preservatives

The terms "fixative" and "preservative" have been used interchangeably throughout this protocol. However, formalin and Kahle’s fluid (see 1 and 2 below) are properly referred to as fixatives, and are used to kill and fix organisms in the field, whereas ethanol (see 3 below) is properly referred to as a preservative, and is intended for long-term use.

1. 10% formalin (100% formalin = 37% aqueous formaldehyde solution; 100 mL of 100% formalin + 900 mL of water = 10% formalin) - A good general-use fixative, although specimens will lose color and become brittle and the shells of molluscs will dissolve after long-term storage. Buffered formalin will avoid the latter. It is best to use formalin as a field fixative and transfer specimens into 70% ethanol for long-term storage. Many investigators prefer not to use formalin at all because it is a skin and eye irritant and may be carcinogenic under prolonged exposure (National Cancer Institute 1996).

2. 10% solution of Kahle’s fluid in water (to make 1L: 15 parts by volume (pbv) of 95% ethanol [290 mL]; 6 pbv of 100% formalin [115 mL]; 1 pbv of glacial acetic acid [20 mL]; and 30 pbv of distilled water [575 mL]; see Wiggins 1977) - This is a good fixative for most insects but should be replaced within hours with 70% ethanol for molluscs and crayfish. It is acidic and will remove calcium from shells and carapaces.

3. 70% ethanol in water (750 mL of 95% ethanol topped up to 1 L with water » 70% ethanol) - A good preservative, especially for crustaceans and insects, but unlike formalin it does not fix tissues. Its main advantage is low toxicity to humans. However, large volumes are required, it is expensive, and concentrations >40% can only be obtained with a permit. Denatured ethanol may be available in higher concentrations. Isopropyl (rubbing) alcohol or the more toxic methanol (methyl hydrate) can often be used as substitutes.

The volatility of ethanol means that it evaporates readily. Thus, collections stored in ethanol must be periodically monitored. Addition of a few drops of glycerine to vials containing alcohol-preserved material will protect them from desiccation, and will keep the specimens from becoming brittle.

Field labels must be added to the inside and outside of vials, jars, or plastic bags. Labeling can be done directly on the vessel or, if it is to be reused, on a piece of duct tape stuck to the outside. Inside labels should be written with a soft-lead pencil on non-recycled, high-quality paper (Appendix II). Labels should include the station number/location, the lake or stream, and the date. It is important that persons receiving the samples for processing understand any date abbreviations used. The samples taken should be entered in the field data book.

Laboratory labels must be added to the inside of vials containing organisms sorted to higher taxa as well as those that have been identified to lower taxa and verified by an expert. Such labels should be written in indelible ink on high-quality or waterproof paper (Appendix II), and should include the following information: (1) collection site, (2) date of sampling, (3) identity of taxon, (4) number of specimens, and (5) identifier.

Blank labels for field and laboratory use can be prepared en masse on a sheet using a word processor and a laser printer or laser photocopier. (Ink-jet printing runs in water or ethanol). They are then cut apart for use. Packs can be prepared for field use by applying rubber cement to one end of a bundle of labels. Sheets of blank labels can be photoreduced for use in small vials.

Standard Field/Laboratory Forms

Large-scale biodiversity surveys are well-served by the development of standard field and laboratory forms, which document characteristics of the habitat sampled, include site drawings or photos, and track samples from the point at which they were taken through to processing and identification of organisms. Examples of standard forms can be found in Cuffney et al. (1993) and Kellogg (1994). (A site form developed for a large biomonitoring study on the Fraser River, British Columbia, is available from either D.M. Rosenberg or A.P. Wiens).

The value of sweep-net collections of adult insects along shorelines to the identification of immature aquatic forms has been discussed above. Another, more time-consuming endeavor is to establish rearing programs to provide associated stages. The taxonomy of aquatic insects is based mainly on the adult form, although the immatures are the forms most frequently collected in aquatic sampling. If the adult, cast pupal skin, and cast last larval skin are available for holometabolous insects (i.e. those with complete metamorphosis such as midge flies), or the adult and a series of cast nymphal skins are available for hemimetabolous insects (i.e. those with incomplete metamorphosis such as mayflies), then the immature forms can often be identified by working backward from the adult. Merritt et al. (1996) review field-based and laboratory rearing methods for major insect groups.

The specimens sorted into major taxa and stored in 10-ml glass vials (see above) can be identified to lower levels by using two excellent, North American texts: Thorp and Covich (1991) for non-insect benthic macroinvertebrates and Merritt and Cummins (1996) for the insects. Both texts provide keys to genera and references to the more specialized literature for species-level determinations.

Once identified, specimens belonging to the same taxon should be stored in their own vial or in a group of shell (tiny) vials plugged with cotton and placed together in a larger vial. Accurate labeling is essential. Counts should be carefully entered onto the sorting sheets described above, or onto sheets specially designed for lower taxa. These data can eventually be entered into an electronic database.

Non-specialists may find it difficult to identify most benthic invertebrates to the species level. Hence, it is wise to send representative, identified material to qualified systematists for verification or get the systematists directly involved in the study. The experts listed above will be able to recommend systematists who specialize in taxa of great relevance to the study. A voucher collection of identified/verified material should be prepared (and curated) for future reference. Curation is important because vials containing alcohol will dry out over time. Voucher collections often prove invaluable in rechecking data, and in taxonomic revisions.