WI DNR Field Procedures Manual
Intranet Edition

Scope

"Benthic invertebrates [are]... the most appropriate biological indicators [of in situ toxic effects] because they are most directly associated with contaminants in sediments through their feeding and behavioral activities." (Reynoldson et al, draft).

Benthic invertebrate communities (living in and directly on the surface layers of sediment) are directly affected by the chemical and physical integrity of the sediment and overlying water. Invertebrates living in the water column are often not directly affected by contaminants in sediment deposits except when contaminants leach into the water column or invertebrates are exposed to suspended sediments moved from the deposit by some event. For this reason, the most direct way to measure the effects of sediment associated contaminants is to survey the invertebrates inhabiting the top 10-15 cm of soft sediment.

Sampling the benthos of surface waters involves many considerations, many of which are not within the scope of this manual. There is a considerable body of scientific literature dealing with the theoretical basis of benthic population distribution and proper study design, other sampling techniques, taxonomy, and statistical analysis of benthic samples. The intent of this section is not to condense the scientific literature but to present collection techniques commonly used in Department investigations. It is assumed that the investigator already has a working knowledge of fresh water benthos and has developed a sound study design prior to collecting samples in the field.

Safety

Formalin (formaldehyde) is a known human carcinogen and should be handled with caution. Wear gloves whenever handling formalin and always choose a well ventilated area to work in; upwind of the samples if outside or under a ventilated hood if processed inside. Samples containing formaldehyde should not be sent through the mail. Any samples containing formalin should be carefully handled and packed during transport to avoid breakage, leaks and spills. Also see section 701.2 on safety.

Equipment

Specific Study Design Considerations

For a summary of basic study design considerations, refer to section 701.1 Planning a Sediment Survey.

Quantitative versus qualitative invertebrate samples (Klemm et al. 1990)

A basic consideration in developing a plan of study for an invertebrate survey is to decide on the type of data needed, either quantitative or qualitative, which then determines the types of sampling and analysis methods used. Quantitative techniques allow the use of statistical techniques in the comparison of site data. Quantitative sampling and analysis also allows the estimation of standing crop (numbers of invertebrates), invertebrate species composition and diversity, biomass, and productivity by measuring the number of invertebrates per unit area of substrate. Three to five or more replicate samples per site (habitat niche, microhabitat or strata) are needed to produce reliable data. The more replicates per site, the greater the precision of the data, i.e., more precise comparisons can be made between sites allowing the detection of more subtle population differences.

Qualitative sampling and analysis techniques cannot generate density information but require fewer samples per site and can provide species presence/absence and richness (relative abundance) information, but not information on standing crop or biomass. Collection of qualitative samples usually entails the use of different collecting devices in all habitats encountered at a site to collect as many different species as possible. Klemm et al. (1990) strongly recommends that a habitat assessment (recorded in the field notebook) be performed at each sampling station because habitat can so strongly affect the presence or absence of species. Only sites with similar habitats where similar efforts should be compared. Artificial substrate samplers which minimize the effects of habitat on species collected can be used at sites with differing habitats if sites are to be compared.

Preparation

Preservative - Prepare the proper strength preservative ahead of time. Be aware that the unpreserved sample will contain some water which will dilute the preservative to some extent.

NOTE: Formalin is the name used for the commonly sold ~40% (by weight) formaldehyde solution. So, 40% formaldehyde = 100% formalin. 10% formalin = 4% (w/w) formaldehyde.

Label all containers with contents (such as distilled water, 10% formalin, 80% ethanol, etc.) to avoid confusion and accidents while in the field.

Refer to the preparation section (C.1.) in 701.4 General Sediment Sampling Equipment and Procedures.

Sample Collection and Processing

The following is derived from Klemm (1990) and this Manual.

Collection procedures will vary, depending upon the type of collecting device used. A coring device is advised when substrate texture allows (not too sandy) and quantitative samples of a particular depth are desired, and/or when different strata of sediment are needed. A grab may also be used for quantitative samples and is advised in very sandy sediment types, but the depth of penetration may vary somewhat but the collection of discreet sediment strata is not possible. Follow procedures described in section General Sediment Sampling Equipment and Procedures for the use of sampling equipment and collection of the sediment samples.

Procedures are described below for processing a benthic invertebrate sample once the sediment is obtained.

1. Sieving Samples

   Sieving invertebrate samples reduces the volume of sediment that must be sorted through in the lab. A #60 sieve (250 µm openings) is recommended for most all new projects in Wisconsin because the smaller invertebrates will be retained by the #60 sieve and should yield more complete invertebrate community data for a site. Number 30 (500 mm) sieves should be used for continuing projects where data using the #30 size sieve already exists.

   1. After collecting a sample, the sample should be placed into a #60 or #30 mesh (250 µm and 500 mm openings, respectively) sieve bucket. Any large debris should be cleaned (remove invertebrates and add them to the sample) and removed from the sample. The sample is then washed through the sieve over the side of the boat or in a tub with site water until no more fine sediment washes through the mesh. Take care not to allow site water into the bucket from the top as this could allow non-sample organisms to contaminate the sample. Washing can be accomplished by adding small portions of the sample at a time to the sieve or the whole sample at once. Many invertebrates are fragile, so this should be done as gently as possible while still getting the job done. If the sample clogs the mesh so water does not drain out, submerge the bucket half way into the water and lower and raise the bucket with enough thrust to push water in from the bottom and suspend the clogging sediment. Performing this action while also sliding the bucket sideways should clean a sample relatively quickly without much damage to the invertebrates. Sometimes the sample will contain chunks of clay that must be carefully broken up with your gloved hands (remember, most toxic contaminants are associated with fine sediments). Another method of cleaning the sample is to use a gloved hand to swish the water while the bucket is partially submerged to resuspend the sample and allow fine sediment to fall out.
   2. After no more fine sediment will wash through the sieve, place sample in sample jars. To move the sample to a jar, tip the sieve bucket so a bottom corner is lowest and wash the remaining sample off the screen and into the bottom corner of the bucket. Wash this into a flat light colored wash pan or directly into the sample jar if possible (a wash bottle and forceps are handy for this). Make sure not to use so much wash water that the final preservative concentration is too dilute.
   3. Quart-size sample jars made of glass (possible breakage and sample loss problem) or plastic make good sample containers. Whirl-pacs or other well sealed bags can also be used, but invertebrates become even more fragile after being preserved, so the container should protect the integrity of the sample during handling and transport.
2. Sample Preservation and Processing

   1. Add 95% ethanol to the sieved sediment sample to a final concentration of ~ 70-80% ethanol. Ethanol is most commonly used to preserve benthic invertebrate samples. If identification of oligochaetes is desired, samples should be preserved in 10% formalin (final concentration) for 10 minutes or longer before transferring them into 70-80% ethanol (Klemm et al, 1990). Samples do not need to be transferred to alcohol if the laboratory doing the identification will accept formalin. If time and space is available in the field, it may be easiest to preserve the samples in formalin in the sorting pan (keep formalin downwind, see safety section), then decant formalin (into appropriate container) and rinse any remaining formalin out with sieved site water (unsieved site water may introduce additional invertebrates).
   2. To exchange ethanol for formalin in sample jars: 1) drain the formalin into a suitable container using a funnel if necessary. This can be accomplished with a #30 or #60 mesh screen placed over the sample jar or by carefully dumping the sample back into the sieve bucket. 2) Rinse out any remaining formalin with sieved site water or tap water. 3) Move sample to sample jar and add ethanol to a final 70-80% concentration.
   3. After addition of the preservative, gently invert sample jar to mix. Invertebrates can become fragile after preservation. Do not shake sample.
   4. Label each sample with tape and permanent marker on the outside of the container. Also place a water resistant label made of part bond paper written in pencil on the inside. Information should include the date, time, site identification, name of person taking sample and replicate number/total samples at the site (sometimes the order the samples were taken in can make a difference).
3. Sorting

   The specific sorting procedure may vary for each study, depending upon the needs of the investigation. In general, sorting of samples obtained with grabs and cores is similar to those obtained with other devices that collect both substrate material and organisms. Sorting is usually accomplished by placing sample material in a shallow, white pan and picking invertebrates from the sample debris with the aid of a low power magnification device (binocular scope or scanning lens). Sorting is made considerably easier when the invertebrates are stained a bright red color from ETOH with Rose Bengal dye added. However, ask the laboratory performing the identifications before using Rose Bengal to make sure it will not interfere with their identification procedures. Prior knowledge of the taxonomic references to be used in identifications is essential before staining, since staining may obscure color patterns sometimes used as diagnostic features for identification.

   The following procedures may be done by the contracted laboratory or by the investigator.

   1. To remove the alcohol or formalin preservative prior to processing, place the sample material on a U.S. Standard #30 or #60 mesh screen or sieve, and wash with water. Be careful to remove all invertebrates from the screen after washing.
   2. Small amounts of sample material are then placed into the sorting pan and searched for invertebrates as noted above.
   3. Invertebrates are removed with a forceps and placed into suitably sized, labeled storage containers with 70-80% ETOH for longterm storage.
4. Subsampling

   When sorting samples with large amounts of sample material or organisms, sorting and analysis time can be considerably reduced by subsampling.

   For routine investigations and when sample sizes are too large to sort 100%, subsampling is accomplished by the following method (Weber, 1973):

   1. Thoroughly mix and distribute the entire sample evenly over the bottom of a shallow, white tray.
   2. Place a divider in the tray which delineates quarter sections.
   3. Sort the two opposite quarters delineated by the divider, or one randomly selected quarter in extremely large samples.
   4. Combine the two remaining quarters and store as reference material or discard.
   5. Follow the same procedure for individual taxonomic groups, if present in excessively large numbers, to reduce analysis time.

   A similar procedure is followed for sorting Hilsenhoff Biotic Index (HBI) samples, except a gridded sorting tray is used and only 100 arthropods are subsampled. Refer to Hilsenhoff (1987) for a complete description.

   Other subsampling methods are possible, depending upon the needs of the study. A method to statistically check on the validity of the subsamples withdrawn and to predict upper and lower confidence limits for the estimated total is presented in Elliott (1977).

5. Taxonomic Identification

   The taxonomic level to which organisms are identified may vary with the objective of the study and should be discussed in each project's Plan of Study, Quality Assurance Project Plan, or similar document. Benthic invertebrate samples are normally sent out to a laboratory for sorting and identification (see below). Only someone with training in the field of benthic invertebrate taxonomy should perform sample identifications. Follow procedures below if samples are to be sorted before being sent to a lab.

   There are two laboratories that the Department currently uses for invertebrate sample analysis (see below). Contracts for invertebrate sample analysis are coordinated by Mike Miller-FH/2 at Central Office. Each lab can only process a limited number of samples in a year.

   1. Dr. Stanley Szczytko's laboratory at the University of Wisconsin-Stevens Point. This laboratory currently processes most or all of the Department's basin assessment and HBI invertebrate samples. They do not perform oligochaete identification beyond family. His address is:

      Dr. Stanley Szczytko
      College of Natural Resources
      UW-Stevens Point
      Stevens Point, WI 54481

   2. Dr. Kurt Schmude directs the invertebrate laboratory at the University of Wisconsin-Superior's Lake Superior Research Institute. This laboratory was used by the DNR for the first time in 1993 with better than satisfactory service. Because of the location of this lab, they may be or become more experienced with the identification of Great Lakes benthic invertebrates (Especially Lake Superior). Services offered by this lab may in the future include data analysis for each sample and project. Special services such as specific identifications or data analysis must be negotiated. His address is:

      Dr. Kurt Schmude
      Lake Superior Research Institute
      Hawkes Hall, Rm 153
      1800 Grand Avenue
      Superior, WI 54880

Documentation

Information about the sample site and collection procedures are normally recorded on Department Form 3200-81, Macroinvertebrate Field and Bench Sheet, and sent with the samples to the lab doing the taxonomy work. They are compatible with the DNR invertebrate (BUG) computer program utilized by Dr. Stan Szczytko's lab. But, because these bench sheets were designed for stream locations and not specifically for soft sediment deposits, and they must be sent with the samples to the lab, you may want to develop a separate field sheet for a specific project. Whichever field sheet is used, it is imperative that all pertinent site information is written down. It is also very important to label the sample container(s) from each site with the corresponding Sample ID number that appears on Form 3200-81 for that site. Copies of this form are available from the download form page.

Quality Assurance

The main concern with quality assurance of benthic core or grab samples is to prevent carryover of organisms from one sample to the next. Carryover is prevented by careful washing and inspection of collection devices and sieve screens after each sample collection.

Another concern is disturbance of sample sites prior to and during sample collection. Anchoring a boat over the sample site must be done carefully to avoid physical disturbance of the area to be sampled. Whenever possible, anchor the boat upstream at least several feet and drift to the site or anchor a few feet away and gently paddle the boat to the sample site. In windy conditions, two anchors may be necessary on different sides of the boat for stabilization. Mobile invertebrates may leave the area or others may be carried offsite by water currents generated by anchoring, causing sample error. A similar problem can occur from the core or grab impacting the substrate.

If the jaws of a grab don't close successfully on retrieval, another sample must be attempted at the same site. It is very important to move successive sampling attempts several feet away from all preceding attempts to avoid sampling previously disturbed substrates.

Invertebrates become brittle in preservative. Samples should be handled as gently as possible to avoid breaking up individual invertebrates.

Refer to EPA 1992, Klemm 1990 and the Quality Assurance Guidance for In-place Pollutant Monitoring Activities (WDNR, 1990) for more detailed quality assurance procedures.

References

See 702.2 section on references for macroinvertebrate sampling references.