Please use the following guidelines when preparing samples in the field. Feel free to contact our Project Coordinator, Shanda McGraw, if you have difficulty finding any of the materials mentioned below.
The best sample containers are heavy duty wide mouth plastic jars such as Nalgene® HDPE or equivalent with screw on lids. The most common sample containers range in size from 500 to 1000 ml.
Good jars can seem expensive but are actually cheap protection for your investment. If at all possible do not use glass jars. In a pinch, ziploc type bags can be used but should be triple bagged. We also provide sampling kits on request. You can choose to rent sampling kits or purchase them outright.
Clear and accurate labels are critical. Use two identical labels per container, one placed inside the container and one attached to the exterior. Use whatever information is appropriate to your needs on the labels, but make sure it clearly defines each sample. Maintain a simple labeling system.
External labels are convenient when organizing samples and provide a safeguard against loss of the internal label. However, exterior labels are subject to much abuse and are often rendered illegible so they should only be used in conjunction with an internal label. Attach the labels to the outside of the jar with clear packing tape, making sure to cover the label entirely. Please note that the ink from a “sharpie” marker or a pen will wash away when exposed to ethanol. If a sample requires multiple containers each container should have two labels and should be clearly marked as 1 of 5, 2 of 5, etc.
Internal labels can be printed on a laser printer using heavy card stock. Use a heavy weight cotton fiber paper (Rite in the Rain) and a No. 2 pencil or Pigma Micron pen if labeling by hand. Heavy card stock and pencil also work well for external labels.
The alcohol preservatives mentioned below are considered Hazardous Materials and MUST be packed and shipped according to Department Of Transportation rules.
The two most common preservatives/fixatives you will encounter are formalin (a solution of 5–10% formaldehyde and water) and ethyl alcohol (ethanol). Ethanol is preferred when shipping samples to the EcoAnalysts lab. Formalin is a toxic substance and requires special handling. Formalin’s acidic qualities may degrade many macroinvertebrates if not properly buffered.
Preserving Macroinvertebrate Samples
Ethanol is a potent enough fixative for most applications and is much safer to deal with. Careful attention must be paid to the final concentration of alcohol in the sample. Too high a concentration will cause the invertebrates to dehydrate and become brittle, and therefore make them very difficult for our macroinvertebrate taxonomists to identify. If the alcohol concentration is too low, the samples will not be adequately preserved and cause your invertebrates to decompose in their containers! For long term storage we recommend a solution of 70-80% ethanol. Since the water entrained in the sample usually dilutes the alcohol considerably, we recommend using a 95% ethanol solution in the field.
Adjust the amount of alcohol to the matrix of the sample. A good rule of thumb is a 1:1 ratio of preservative to sample material. If the sample is large and just barely fits in the jar, you are better off splitting it into separate containers to allow adequate preservation.
Invertebrates die and decompose remarkably fast, so don’t delay preserving them. If for some reason you are unable to use preservative in the field, put the samples on ice, out of direct sunlight, and preserve them as soon as possible. You may find that isopropyl alcohol (rubbing alcohol) is more readily obtainable. Isopropyl is a good alternative to pure ethanol, but make sure you get the technical grade (99%). Denatured alcohol is also available but is a less attractive alternative as it often contains toxic additives to that could cause the macroinvertebrates to become more difficult to identify.
Preserving Fish Samples
Fish samples should be placed in wide mouth plastic jars with screw on lids, please do not use glass jars.
The fish should originally be preserved in a 10% formalin solution for at least a week. For large specimens, it is a good idea to open the body cavity to allow the fixative to penetrate. After the specimens have been fixed by the formalin, they should be transferred to alcohol for long term storage. Do not use alcohol as a fixative, it does not preserve internal organs well and it tends to dehydrate the muscle tissue making the fish brittle.
Preserving Periphyton Samples
There are several preservatives available, Lugol’s solution and “M3″ fixative are the best. You may also use buffered 4% formalin or 2% glutaraldehyde. For M3 a ratio of 20mL to 1L of sample is adequate for preservation.
- M3 Fixative
- 5g potassium iodide
- 10g iodine crystals (sublimed)
- 50mL formalin
- 1L distilled water
- Combine and dissolve.
Preserving Zooplankton Samples
Ethanol or isopropanol (30-40%) are the preservatives of choice for samples that will be analyzed less than a year after they were collected. For long term storage (greater than one year) 2-4% formaldehyde is the only permanent preservative routinely used for zooplankton, though it is generally modified by the addition of several other chemicals – sodium borate, sugar, and/or BHT usually – for different purposes. Keep in mind that formaldehyde is recognized as a toxic, carcinogenic substance. Use alcohol when ever possible.