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WI DNR Field Procedures Manual
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Part B: Collection Procedures

702.2 Benthic Invertebrate Survey - Artificial Substrates

  1. Scope
  2. Artificial substrates are useful for collecting macroinvertebrates associated with the water column and can be used to demonstrate water quality and effects of contaminated sediments on the overlying or downstream water column. They can be used in almost any stream or lake habitat, either suspended in the water column or anchored to the bottom. They are left in the water for relatively long time periods (2-8 weeks) to be colonized by drifting or swimming aquatic organisms.

    The main advantage of artificial substrates is the minimization of the effects of physical variables between sites such as: substrate type, depth, and light penetration. Data comparisons between stations are, therefore, simplified.

    Some disadvantages include: large effort and time to results (two separate field trips, 2-8 weeks to colonize), loss of sample because of weather or vandalism, and unquantifiable loss of invertebrates if samplers are accidentally disturbed.

    For sediment assessments, the most significant attribute is the bias for aquatic rather than benthic invertebrates. Invertebrate colonies from artificial substrates generally reflect water quality rather than sediment quality unless the sediment is directly affecting the quality of the water above it. So, artificial substrates should be used to demonstrate a change in water quality near or downstream from a sediment and in conjunction with other sediment data, but should not be used as a direct measure of sediment quality alone. Discussions of both the advantages and disadvantages of artificial substrate samplers appear in, Klemm et al (1990), Rosenberg and Resh (1982), Weber (1973) and Chapter 5 of Downing (1984).

  3. Equipment
  4. Several different designs of artificial substrates are available, but only two common types will be described here. Other types are described in the scientific literature (Klemm et al, 1990; Rosenberg and Resh, 1982).

    1. Multi-plate Samplers

      Multiplate artificial substrate samplers consist of a series of square or circular hardboard discs, separated by spacers and fastened together through their centers to a threaded eyebolt. The standardized, reproducible and easily measured substrate surface areas allow for uniform replicates and the collection of quantitative data. The substrates can either be suspended or anchored a predetermined distance from the water or sediment surface.

      The Fullner (1971) modification to the Hester-Dendy (1962) multiple-plate design is widely used. The modified Hester-Dendy sampler is constructed of 0.3 cm (0.125 in) thick tempered hardboard with 7.6 cm (3 in) diameter round plates and 2.5 cm (1 in) round spacers that have 5/8 in center-drilled holes. The plates are separated by spacers on a 1/4 in diameter eyebolt, held in place by a nut at the top and bottom. A total of 14 large plates and 24 spacers are used. The top nine plates are each separated by a single spacer, plates 9 and 10 are separated by two spacers, plates 11 and 12 are separated by three spacers, and plates 13 and 14 are separated by four spacers. The hardboard sampler is about 14 cm (5.5 in) long and has a surface area of about 1,160 cm2 (0.116 m2). The surface area and dimensions will change from absorption of water. Hardboard samplers exposed to toxicants, oils and/or preservatives (alcohol, formalin) cannot be reused.

    2. Rock Baskets

      The other common type of artificial substrate is the rock basket sampler most commonly composed of a cylindrical, chrome plated basket (barbecue basket) filled with 1 to 3inch diameter rocks or rocklike material. See Mason et al. (1967) or Klemm et al (1990) for a more detailed description of a sampler and suspension system. It is more difficult to calculate surface area for rock basket samplers, but the rocks simulate rubble type natural substrates better than multiplate samplers.

    3. Alternative substrate materials

      Alternative substrate materials that have been used in samplers include: 3M plastic mesh (conservative webbing, 3M corporation, St. Paul, MN), natural rubble, natural leaf litter or any other suitable substrate for colonization of invertebrates. Samplers constructed from rotisserie baskets and 3M mesh have been described by Stauffer et al. (1976) and Swift (1985).

    4. Equipment Checklist Artificial Substrates
      Artificial substrates.
      Anchoring or suspension equipment for artificial substrates.
      Dip net or net bag for retrieval of artificial substrates.
      Shallow, white pan for cleaning artificial substrates.
      Wash bottle and forceps.
      Labeled sample containers large enough to enclose assembled, multiplate artificial substrates,
      or
      Labeled sample containers large enough to contain contents of rock basket type artificial substrates.
      Labels for inside sample jars.
      Preservative - 95% EtOH, formalin.
      Tools for assembly and deployment or retrieval (cutting tool, pliers, etc.)
      Permanent marker and pencils.
      Field sheets and/or Forms 3200-81, one per sample site.

  5. Collection Procedure
    1. Setting the samplers

      Artificial substrates are anchored or suspended at the sample site to be colonized by drifting and swimming invertebrates. Placement within the water column depends upon the objectives of the study. For sediment assessments, the samplers should usually be placed close to the sediment, but samplers from all sites should be at similar depths. For water quality assessment, samplers are normally placed in the euphotic zone (1-3 feet from water surface) to collect the highest diversity and abundance of invertebrates possible (Mason et al. (1973) in: Klemm et al. (1990)).

      Care should be taken to prevent silt from building up and thus reducing the colonizable surface area of the substrate.

      To prevent vandalism, place samplers out of the view of passers by and away from boat traffic and heavily used fishing areas. If water levels at the site will fluctuate, place the sampler so it will be midway in the water column at low-flow or at least will not ever be exposed to the air.

      It is recommended to place at least three samplers at each site. There is no exact standard for exposure time, but six weeks is recommended by many authorities (Klemm et al, 1990, American Public Health Association, 1985, and Weber, 1973). Whatever the exposure time, data should not be compared between samplers exposed for different lengths of time.

      The best times for sampling invertebrates is spring/early summer, and fall. This is when there is the most activity and the highest abundance of invertebrates can be obtained. Remember that invertebrate assemblages will change through the seasons, so the time of year will affect the numbers and species of invertebrates obtained.

    2. Retrieving Samplers

      The method used to retrieve an artificial substrate is critical to the resulting quality of data. Some invertebrates will abandon the substrate as soon as the sampler is disturbed. For this reason, samplers should be retrieved with great care to reduce the loss of invertebrates from the sample. Ideally, the sampler is approached from downstream and enclosed with a U.S. Standard #30 or finer mesh bag or dip net before it is moved and brought to the surface. SCUBA or a remote enclosure system can be used in deep water. But, if these are not viable options, samplers should be pulled up with the least amount of disturbance and netted before reaching the surface.

      The same procedures to deploy samplers should be used for all samples and sites that will be compared (e.g., depth, exposure period, current velocity, sunlight exposure, habitat type).

  6. Sample Processing
  7. After retrieval, multi-plate samplers can be placed directly into containers large enough to hold the sampler plates (one quart, large mouth jars or sturdy plastic bags) and preserved with 70-80% ethanol until processing; or they can be cleaned (organisms removed) in the field. Rock baskets are almost always cleaned in the field. The contracting laboratories will normally clean samplers if asked, but the work must be included in the contract, and will cost slightly more per sample.

    Remember: If stored in preservative, substrates absorb the preservative and are considered unsuitable for reuse.

    Alternatively, samplers can be stored in sieved, refrigerated water from the sampling site. Sorting and preservation should proceed within 24 hours of storage.

    With all artificial substrate types, the enclosing net used during retrieval should be inspected for invertebrates and the recovered invertebrates included in the sample.

    Always place a water resistant sample label inside the bag or jar as well as on the outside. Information should include: Site ID, location, habitat type, time, date and names of collectors.

    1. Cleaning samplers

      1. Disassemble the sampler and place plates, spacers or rocks in a tub of water (sieved to prevent introducing non-sample organisms if site water is used) and gently scrub plates or rocks into the tub with a soft bristled brush to remove all invertebrates.
      2. Concentrate the resulting sample by pouring the tub of water through an appropriate sized sieve, and then place into sample containers with the appropriate preservative (10% formalin or 70-80% ethanol). With multiplate samplers, the sampler is disassembled and each plate and spacer is inspected with the aid of a low power magnifier. All target invertebrates are then removed for preserved storage in 70-80% ETOH. The ETOH remaining within the sample containers is then poured through a #30 or #60 mesh sieve and all retained macroinvertebrates are included in the sample. The inside of the emptied sample container should also be inspected for invertebrates to be included in the sample.
      3. For rock basket samplers, rocks stored in preservative are treated similarly to the plates in the above example. If only the organisms and associated materials from the rock surfaces were retained in ETOH, the ETOH/organism mixture is passed through a #30 or #60 mesh sieve. The retained organisms and debris are then transferred to a white pan for sorting, with the aid of a low power magnification device.

      For quantitative work, the surface area of the artificial substrate sampler must be determined to allow for calculation of organism density per unit surface area.

    2. Sorting and Subsampling

      Refer to the sorting and subsampling parts of 702.1 Benthic Invertebrate Surveys - Benthic Samples.

    3. Taxonomic Identification

      Refer to the taxonomic identification part of 702.1 Benthic Invertebrate Surveys - Benthic Samples above for procedures and a list of invertebrate taxonomy laboratories.

  8. Documentation
  9. Information about the sample site and collection procedures are recorded on Department Form 3200-81, Macroinvertebrate Field and Bench Sheet. It is very important to label the specimen container(s) from each site with the corresponding Sample ID number that appears on Form 3200-81 for that site. See 701.3 for additional information on documentation. See the download section for an electronic copy of form 3200-81.

  10. Quality Assurance
  11. To ensure quality data from artificial substrate samples, the investigator should be aware of the following:

    1. Samplers should be retrieved very carefully to prevent the loss of organisms. Samplers should not be disturbed at all prior to retrieval (approach site from downstream) and should be enclosed during retrieval with a net having a mesh size at least as fine as U.S. Standard #30 or #60 or small enough to retain target organisms.
    2. Multiplate samplers or others made from porous materials should not be reused after exposure to oils/toxins in water or storage in preservatives.
    3. Samplers to be reused should be inspected carefully to remove all organisms to prevent carryover to successive sample sites or exposure periods.
    4. Artificial substrates may not sample all organisms found on natural substrates with equal efficiency. Use caution when interpreting data from artificial substrate samples. Refer to Chapter 5 of Downing (1984) for a more complete discussion.
    5. Sampler to sampler variability can be estimated from results for replicate samplers placed at each location. Samplers should be placed so that they do not interfere with each other by changing water flow, light, siltation, etc.

  12. References
  13. ASTM. 1992. E 1469-92. Standard practice for collecting benthic macroinvertebrates with multiple-plate samplers. Philadelphia, PA.

    American Public Health Association. Standard Methods for the Examination of Water and Wastewater (Washington D.C.: American Public Health Association, 1985), pp. 1268.

    Burton, G.A. Jr. (Ed.) 1992. Sediment Toxicity Assessment. Lewis Publishers, Boca Raton, Florida. 457pp.

    Downing, John A. and F. H. Rigler (eds.). 1984. Sampling the benthos of standing waters, a manual on methods for the assessment of secondary productivity in fresh waters. IBP Handbook 12, 2nd Edition. Blackwell Scientific, Oxford, U.K. 501 pp.

    EPA. 1992. Sediment Classification Methods Compendium. Office of Water, Washington DC. EPA 823-R-92-006.

    Klemm, D.J., P.A. Lewis, F. Fulk, and J.M. Lazorchak. 1990. Macroinvertebrate field and laboratory methods for evaluating the biological integrity of surface waters. Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency, Cincinnati, OH 45268. EPA/600/4-90/030.

    Mason, W.T., J.B. Anderson and G.E. Morrison. 1967. Limestone-filled, artificial substrate sampler-float unit for collecting macroinvertebrates in large streams. Prog. Fish-Cult. 29:74.

    Mason W.T., Jr., C.I. Weber, P.A. Lewis and E.C. Julian. 1973. Factors affecting the performance of basket and multiplate macroinvertebrate samplers. Freshwater Biology 3:409-436.

    Reynoldson, T.B., K.E. Day and R.H. Norris. (In review). Biological guidelines for freshwater sediment based on Benthic Assessment of Sediment (the BEAST) using a multivariate approach for predicting biological state. Submitted to: Australian Journal of Ecology.

    Rosenberg, D.M. and V.H. Resh. 1982. The use of artificial substrates to study freshwater benthic invertebrates. In: J. Cairns Jr. (ed.). Artificial Substrates. Ann Arbor Science Publ., Ann Arbor, MI. pp.175-235.

    Weber, C. I. (ed.). 1973. Biological field and laboratory methods for measuring the quality of surface waters and effluents. USEPA, Cincinnati, USA. EPA670/473001.

  14. References for Additional Information
    1. General Benthic Ecology:

      Hart, C.W., Jr. and S.L.H. Fuller (eds.) 1974. Pollution ecology of freshwater invertebrates. Academic Press. New York. 389 pp.

      Hynes,H.B.N. 1970. The ecology of running waters. Univ. Toronto Press. Toronto, Canada. 555 pp.

      Resh, Vincent H. and D. M. Rosenberg (eds.). 1984. The ecology of aquatic insects. Praeger Publ. New York. 625 pp.

      Thorp, J.H. and A.P. Covich (eds.) 1991. Ecology and classification of North American freshwater invertebrates. Academic Press, Inc. San Diego, CA. 911 pp.

    2. Study Design and Sampling Methods:

      ASTM. 1992. Standard practice for Collecting Benthic Macroinvertebrates with Multiple-plate Samplers. E 1469-92.

      Downing, John A. and F. H. Rigler (eds.). 1984. A manual on methods for the assessment of secondary productivity in fresh waters. IBP Handbook 17, 2nd Edition. Blackwell Scientific, Oxford, U.K. 501 pp.

      Elliott, J. M. 1977. Some methods for the statistical analysis of samples of benthic invertebrates. 2nd Edition. Ambleside: Freshwater Biol. Assoc. Sci. Publ. 25.

      EPA. 1992. Sediment Classification Methods Compendium. Office of Water, Washington DC. EPA 823-R-92-006.

      Green, Roger H. 1979. Sampling design and statistical methods for environmental biologists. John Wiley & Sons. New York. 257 pp.

      Hilsenhoff, William L. 1987. An improved biotic index of organic stream pollution. Great Lakes Entomol. 20(1): 3139.

      Klemm, D.J., P.A. Lewis, F. Fulk, and J.M. Lazorchak. 1990. Macroinvertebrate field and laboratory methods for evaluating the biological integrity of surface waters. Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency, Cincinnati, OH 45268. EPA/600/4-90/030.

      Stauffer, J.R., Jr., H.A. Beiles, J.W. Cox, K.L. Dickson and D.E. Simonet. 1976. Colonization of macrobenthic communities on artificial substrates. Revista de Biologia 10:49-51.

      Weber, C. I. (ed.). 1973. Biological field and laboratory methods for measuring the quality of surface waters and effluents. USEPA, Cincinnati, USA. EPA670/473001.

      WDNR. 1990 (draft). Quality Assurance Guidance for In-Place Pollutant Monitoring Activities. Unpublished document on file at the Office of Technical Services, Bureau of Water Resources Management.

    3. Sample Analysis:

      Downing, John A. and F. H. Rigler (eds.) 1984. A manual on methods for the assessment of secondary productivity in fresh waters. IBP Handbook 17, 2nd Edition. Blackwell Scientific, Oxford, U.K. 501 pp.

      Elliott, J. M. 1977. Some methods for the statistical analysis of samples of benthic invertebrates. 2nd Edition. Ambleside: Freshwater Biol. Assoc. Sci. Publ. 25.

      Green, Roger H. 1979. Sampling design and statistical methods for environmental biologists. John Wiley & Sons. New York. 257 pp.

      Hilsenhoff, William L. 1987. An improved biotic index of organic stream pollution. Great Lakes Entomol. 20(1): 3139.

      Klemm, D.J., P.A. Lewis, F. Fulk, and J.M. Lazorchak. 1990. Macroinvertebrate field and laboratory methods for evaluating the biological integrity of surface waters. Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency, Cincinnati, OH 45268. EPA/600/4-90/030.

      Narf, Richard P., E. L. Lange and R. C. Wildman. 1984. Statistical procedures for applying Hilsenhoff's Biotic Index. J. Freshwater Ecol. 2(5)441448.

      Weber, C. I. (ed.). 1973. Biological field and laboratory methods for measuring the quality of surface waters and effluents. USEPA, Cincinnati, USA. EPA670/473001.

    4. General Taxonomy:

      Hilsenhoff, William L. 1981. Aquatic insects of Wisconsin. Nat. Hist. Council, UW Madison, No. 2. Madison, WI 60 pp.

      Hilsenhoff, William L. 1982. Using a biotic index to evaluate water quality in streams. Tech. Bull. No. 132. Dept. of Nat. Res. Madison, WI 22 pp.

      Merritt, R. W. and K. W. Cummins (eds.). 1978. An introduction to the aquatic insects of North America. KendallHunt Publ. Co. Dubuque, Iowa, USA.

      Pennack, Robert W. 1978. Freshwater invertebrates of the United States, 2nd Edition. John Wiley & Sons, New York. 803 pp.

      Thorp, J.H. and A.P. Covich (eds.) 1991. Ecology and classification of North American freshwater invertebrates. Academic Press, Inc. San Diego, CA. 911 pp.

      Many more taxonomic and other references are available in the scientific literature. An excellent summary of the literature for benthos is published by the North American Benthological Society in their annually published series "Current and Selected Bibliographies on Benthic Biology".

Rev. 0, April 1995

This document is intended solely as guidance and does not contain any mandatory requirements except where requirements found in statute or administrative rule are referenced. This guidance does not establish or affect legal rights or obligations and is not finally determinative of any of the issues addressed. This guidance does not create any rights enforceable by any party in litigation with the State of Wisconsin or the Department of Natural Resources. Any regulatory decisions made by the Department of Natural Resources in any matter addressed by this guidance will be made by applying the governing statutes and administrative rules to the relevant facts. (From Manual Code 1210.1)

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