Methods Protocol

 

Benthic Macroinvertebrate

Assessment

Benthic Macroinvertebrate Monitoring Protocol

 

 

 

Instrumentation:

 

  1. Standard Kick Net, Open Mesh, 1.0 Meter Width
  2. Sieve Bucket
  3. 70% Ethanol
  4. Sample Containers – Plastic Screw Lid
  5. Standardized Data Sheets
  6. Garmin 3 GPS
  7. Velocity Meter
  8. Meter Stick and/or Tape Measure
  9. Surber Net, .3 meter by .3 meter`1

 

 

 

Quality Control/ Quality Assurance

 

  1. All sample containers are properly labeled with date, site location, contents, and monitors last name
  2. Standardized data sheets used to record identified benthic invertebrates
  3. After sampling in the field all nets are inspected to insure all debris, invertebrates, etc are included in the container, the nets are then thoroughly washed before next sampling

 

Freestone Sub Sampling Protocol

Grid Method

 

 

  1. stain and rinse sample
  2. place entire sample into a gridded pan
  3. use magnesium sulfate (Epson salt) mixture to make the organisms float
  4. pick all organisms from the pan
  5. pick a random number from a random numbers table or numbered caps: record number
  6. whatever number you choose is the number grid you will start working on
  7. pick and count all organisms out of the grid
  8. repeat steps 6-8 until there is 20% + or – 200 organisms

 

 

 Archived Samples-

  • If you are saving the remaining unidentified sample label sample jar:

Unsorted Sample Residue

 

  • If you are only saving the sub sample label small vile:

Sorted Sample Residue

Freestone Sub Sampling Protocol

Cookie Cutter Method

 

  1. strain and rinse sample
  2. place entire sample into a 28 grid pan (spread out evenly)
  3. pick 4 numbers from the random numbers dish
  4. write down the four numbers for the tier one of the sub sample
  5. place cookie cutter (soup can) around each grid packed
  6. using a plastic knife cute the debris within the cookie cutter
  7. to remove the debris and organisms a plastic spoon works well
  8. the debris and organisms are placed into a second 28 grid pan (spread out evenly)
  9. pick one number from the random numbers dish
  10. write down this number for tier two of the sub sample
  11. remove debris and organisms from the grid chosen in the second grid pan
  12. place debris in a Petri dish
  13. using a microscope; pick out all organisms and count
  14. repeat steps 9-13 until there is +/- 20% 200 organisms

 

Archived samples:

  • If you are saving the sub sample, label the small vile: Sorted Sub Sample

 

  • If you are saving the remaining unidentified sample, label the sample jar: Unsorted Sample Residue

Limestone Sub Sampling Protocol

Cookie Cutter Method

 

  1. strain and rinse sample
  2. place entire sample into a 28 grid pan (spread out evenly)
  3. pick 4 numbers from the random numbers dish
  4. write down the four numbers for the tier one of the sub sample
  5. place cookie cutter (soup can) around each grid packed
  6. using a plastic knife cute the debris within the cookie cutter
  7. to remove the debris and organisms a plastic spoon works well
  8. the debris and organisms are placed into a second 28 grid pan (spread out evenly)
  9. pick one number from the random numbers dish
  10. write down this number for tier two of the sub sample
  11. remove debris and organisms from the grid chosen in the second grid pan
  12. place debris in a Petri dish
  13. using a microscope; pick out all organisms and count
  14. repeat steps 9-13 until there is +/- 20% 300 organisms

 

Archived samples:

  • If you are saving the sub sample, label the small vile:

Sorted Sub Sample

 

  • If you are saving the remaining unidentified sample, label the sample jar: Unsorted Sample Residue

Method Protocol for Velocity and flow determination:

 

 

  1. Choose a cross-section of the stream where current and depth are most uniform and measure the width
  2. Split the stream into three equal segments and determine the current velocity of the midpoint of each segment (both surface and bottom velocity)
  3. Use the formula R=WDaV to determine the volume of flow for each segment of the cross section where R is the volume of flow, W is width, D is depth, as is a bottom factor constant (.8 for rocks; .9 for sand, mud, hardpan, or bedrock area and V is the surface velocity. Determine the total volume of flow by adding the three R values.

 

 

 

 

 

 

 

 

 

 

 

References:

 

Zimmerman, Mel. Biology 224 - Ecology Lab Manual. Williamsport: Lycoming College,                        1998. 1-9.

 

 

 

 

Methods~ Kick Net Sampling

 

1. Site selection should include visibly moving water as well as a riffle

2. The kick net should be set about 1 meter downstream from the riffle

3. The first person supports the net while the second kicks a one meter by one meter area behind the riffle ensuring to stir up sediment beneath rocks allowing debris to flow downstream into the net (any large rocks in the area should be rubbed free of debris in front of the net and then removed to make kicking easier)

4. The “kicker” lifts the net by the ends previously submerged in the water being careful not to allow any debris to float away and places the net in a sieve bucket.

5. A second regular plastic bucket is used to rinse the contents of the net into the sieve bucket, forceps or fingers may be used to pull off anything that sticks to the net

6. The contents of the sieve bucket are rinsed and placed in plastic containers (because of the qualitative nature of the sample, it is important that all debris caught in the net is included in the final sample)

7. 70% ethanol is added to kill and preserve macro-invertebrates.

8. Plastic containers should be labeled with monitors last name, site location, date, and contents

 

 

 

 

 

 

 

 

7 Benthic Macroinvertebrate Protocols

*** For the full file go to http://www.epa.gov/owow/monitoring/rbp/ch07main.html ***

This Chapter is divided into two parts: Part A (this file), and Part B

Rapid bioassessment using the benthic macroinvertebrate assemblage has been the most popular set of protocols among the state water resource agencies since 1989 (Southerland and Stribling 1995). Most of the development of benthic Rapid Bioassessment Protocols (RBPs) has been oriented toward RBP III (described in Plafkin et al. 1989). As states have focused attention on regional specificity, which has included a wide variety of physical characteristics of streams, the methodology of conducting stream surveys of the benthic assemblage has advanced. Some states have preferred to retain more traditional methods such as the Surber or Hess samplers (e.g., Wyoming Department of Environmental Quality [DEQ]) over the kick net in cobble substrate. Other agencies have developed techniques for streams lacking cobble substrate, such as those streams in coastal plains. State water resource agencies composing the Mid-Atlantic Coastal Streams (MACS) Workgroup, i.e., New Jersey Department of Environmental Protection (DEP), Delaware Department of Natural Resources and Environmental Control (DNREC), Maryland Department of Natural Resources (DNR) and Maryland Department of the Environment (MDE), Virginia DEQ, North Carolina Department of Environmental Management (DEM), and South Carolina Department of Health and Environmental Control (DHEC), and a workgroup within the Florida Department of Environmental Protection (DEP) were pioneers in this effort. These 2 groups (MACS and FLDEP) developed a multihabitat sampling procedure using a D-frame dip net. Testing of this procedure by these 2 groups indicates that this technique is scientifically valid for low-gradient streams. Research conducted by the U.S. Environmental Protection Agency (USEPA) for their Environmental Monitoring and Assessment Program (EMAP) program and the United States Geological Survey (USGS) for their National Water Quality Assessment Program (NAWQA) program have indicated that the rectangular dip net is a reasonable compromise between the traditional Surber or Hess samplers and the RBP kick net described the original RBPs.

STANDARD BENTHIC MACROINVERTEBRATE SAMPLING GEAR TYPES FOR STREAMS (assumes standard mesh size of 500 ΅ nytex screen)

  • Kick net: Dimensions of net are 1 meter (m) x 1 m attached to 2 poles and functions similarly to a fish kick seine. Is most efficient for sampling cobble substrate (i.e., riffles and runs) where velocity of water will transport dislodged organisms into net. Designed to sample 1 m2 of substrate at a time and can be used in any depth from a few centimeters to just below 1m (Note -- Depths of 1m or greater will be difficult to sample with any gear).
  • D-frame dip net: Dimensions of frame are 0.3 m width and 0.3 m height and shaped as a "D" where frame attaches to long pole. Net is cone or bag-shaped for capture of organisms. Can be used in a variety of habitat types and used as a kick net, or for "jabbing", "dipping", or "sweeping".
  • Rectangular dip net: Dimensions of frame are 0.5 m width and 0.3 m height and attached to a long pole. Net is cone or bag-shaped. Sampling is conducted similarly to the D-frame.
  • Surber: Dimensions of frame are 0.3 m x 0.3 m, which is horizontally placed on cobble substrate to delineate a 0.09 m2 area. A vertical section of the frame has the net attached and captures the dislodged organisms from the sampling area. Is restricted to depths of less than 0.3 m.
  • Hess: Dimensions of frame are a metal cylinder approximately 0.5 m in diameter and samples an area 0.8 m2. Is an advanced design of the Surber and is intended to prevent escape of organisms and contamination from drift. Is restricted to depths of less than 0.5 m.

From the testing and implementation efforts that have been conducted around the country since 1989, refinements have been made to the procedures while maintaining the original concept of the RBPs. Two separate procedures that are oriented toward a "single, most productive" habitat and a multihabitat approach represent the most rigorous benthic RBP and are essentially a replacement of the original RBP III. The primary differences between the original RBP II and III are the decision on field versus lab sorting and level of taxonomy. These differences are not considered sufficient reasons to warrant separate protocols. In addition, a third protocol has been developed as a more standardized biological reconnaissance or screening and replaces RBP I of the original document.

Kicknet
Kick net

D-frame Dipnet
D-frame Dip net

Rectangular Dipnet
Rectangular Dip net

Hess sampler
Hess sampler
(Mary Kay Corazalla, Univ. of Minnesota)

 

7.1

SINGLE HABITAT APPROACH: 1 METER KICK NET

The original RBPs (Plafkin et al. 1989) emphasized the sampling of a single habitat, in particular riffles or runs, as a means to standardize assessments among streams having those habitats. This approach is still valid, because macroinvertebrate diversity and abundance are usually highest in cobble substrate (riffle/run) habitats. Where cobble substrate is the predominant habitat, this sampling approach provides a representative sample of the stream reach. However, some streams naturally lack the cobble substrate. In cases where the cobble substrate represents less than 30% of the sampling reach in reference streams (i.e., those streams that are representative of the region), alternate habitat(s) will need to be sampled (See Section 7.2). The appropriate sampling method should be selected based on the habitat availability of the reference condition and not of potentially impaired streams. For example, methods would not be altered for situations where the extent of cobble substrate in streams influenced by heavy sediment deposition may be substantially reduced from the amount of cobble substrate expected for the region.

7.1.1    Field Sampling Procedures for Single Habitat

 

        1.

 

                FIELD EQUIPMENT/SUPPLIES NEEDED FOR BENTHIC

                MACROINVERTEBRATE SAMPLING

n       SINGLE HABITAT APPROACH

  • standard kick-net, 500 ΅ opening mesh, 1.0 meter width
  • sieve bucket, with 500 ΅ opening mesh
  • 95% ethanol
  • sample containers, sample container labels
  • forceps
  • pencils, clipboard
  • Benthic Macroinvertebrate Field Data Sheet*
  • first aid kit
  • waders (chest-high or hip boots)
  • rubber gloves (arm-length)
  • camera
  • Global Positioning System (GPS) Unit

* It is helpful to copy field sheets onto water-resistant paper for use in wet weather conditions

 

A 100 m reach representative of the characteristics of the stream should be selected. Whenever possible, the area should be at least 100 meters upstream from any road or bridge crossing to minimize its effect on stream velocity, depth, and overall habitat quality. There should be no major tributaries discharging to the stream in the study area.

 

2.     Before sampling, complete the physical/chemical field sheet (see Chapter 5; Appendix A-1, Form 1) to document site description, weather conditions, and land use. After sampling, review this information for accuracy and completeness.

3.    Draw a map of the sampling reach. This map should include in-stream attributes (e.g., riffles, falls, fallen trees, pools, bends, etc.) and important structures, plants, and attributes of the bank and near stream areas. Use an arrow to indicate the direction of flow. Indicate the areas that were sampled for macroinvertebrates on the map. Estimate "river mile" for sampling reach for probable use in data management of the water resource agency. If available, use hand-held Global Positioning System (GPS) for latitude and longitude determination taken at the furthest downstream point of the sampling reach.

  1. All riffle and run areas within the 100-m reach are candidates for sampling macroinvertebrates. A composite sample is taken from individual sampling spots in the riffles and runs representing different velocities. Generally, a minimum of 2 m2 composited area is sampled for RBP efforts.
  2. Sampling begins at the downstream end of the reach and proceeds upstream. Using a 1 m kick net, 2 or 3 kicks are sampled at various velocities in the riffle or series of riffles. A kick is a stationary sampling accomplished by positioning the net and disturbing one square meter (continued below)

ALTERNATIVES FOR STREAM REACH DESIGNATION

  • Fixed-distance designation--A standard length of stream, such as a reach, is commonly used to obtain an estimate of natural variability. Conceptually, this approach should provide a mixture of habitats in the reach and provide, at a minimum, duplicate physical and structural elements such as a riffle/pool sequence.
  • Proportional-distance designation-- Alternatively, a standard number of stream "widths" is used to measure the stream distance, e.g., 40 times the stream width is defined by EMAP for sampling (Klemm and Lazorchak 1995). This approach allows variation in the length of the reach based on the size of the stream.
  1. (continued) upstream of the net. Using the toe or heel of the boot, dislodge the upper layer of cobble or gravel and scrape the underlying bed. Larger substrate particles should be picked up and rubbed by hand to remove attached organisms. If different gear is used (e.g., a D-frame or rectangular net), a composite is obtained from numerous kicks (See Section 7.2).
  1. The jabs or kicks collected from different locations in the cobble substrate will be composited to obtain a single homogeneous sample. After every kick, wash the collected material by running clean stream water through the net 2 to 3 times. If clogging does occur, discard the material in the net and redo that portion of the sample in a different location. Remove large debris after rinsing and inspecting it for organisms; place any organisms found into the sample container. Do not spend time inspecting small debris in the field. [Note -- an alternative is to keep the samples from different habitats separated as done in EMAP (Klemm and Lazorchak 1995).]
  2. Transfer the sample from the net to sample container(s) and preserve in enough 95 percent ethanol to cover the sample. Forceps may be needed to remove organisms from the dip net. Place a label indicating the sample identification code or lot number, date, stream name, sampling location, and collector name into the sample container. The outside of the container should include the same information and the words "preservative: 95% ethanol". If more than one container is needed for a sample, each container label should contain all the information for the sample and should be numbered (e.g., 1 of 2, 2 of 2, etc.). This information will be recorded in the "Sample Log" at the biological laboratory (Appendix A-3, Form 2).
  3. Complete the top portion of the "Benthic Macroinvertebrate Field Data Sheet" (Appendix A-3, Form 1), which duplicates the "header" information on the physical/chemical field sheet.
  4. Record the percentage of each habitat type in the reach. Note the sampling gear used, and comment on conditions of the sampling, e.g., high flows, treacherous rocks, difficult access to stream, or anything that would indicate adverse sampling conditions.
  5. Document observations of aquatic flora and fauna. Make qualitative estimates of macroinvertebrate composition and relative abundance as a cursory estimate of ecosystem health and to check adequacy of sampling.
  6. Perform habitat assessment (Appendix A-1, Form 2) after sampling has been completed; walking the reach helps ensure a more accurate assessment. Conduct the habitat assessment with another team member, if possible.

12.   Return samples to laboratory and complete log-in form (Appendix A-3, Form 2).

QUALITY CONTROL (QC) IN THE FIELD

  1. Sample labels must be properly completed, including the sample identification code, date, stream name, sampling location, and collector's name, and placed into the sample container. The outside of the container should be labeled with the same information. Chain-of-custody forms, if needed, must include the same information as the sample container labels.
  2. After sampling has been completed at a given site, all nets, pans, etc. that have come in contact with the sample should be rinsed thoroughly, examined carefully, and picked free of organisms or debris. Any additional organisms found should be placed into the sample containers. The equipment should be examined again prior to use at the next sampling site.
  3.  Replicate (1 duplicate sample) 10% of the sites to evaluate precision or repeatability of the sampling technique or the collection team.

 

FIELD EQUIPMENT/SUPPLIES NEEDED FOR BENTHIC MACROINVERTEBRATE SAMPLING--MULTI-HABITAT APPROACH

  • standard D-frame dip net, 500 opening mesh, 0.3 m width (~ 1.0 ft frame width)
  • sieve bucket, with 500 opening mesh
  • 95% ethanol
  • sample containers, sample container labels
  • forceps
  • pencils, clipboard
  • Benthic Macroinvertebrate Field Data Sheet*
  • first aid kit
  • waders (chest-high or hip boots)
  • rubber gloves (arm-length)
  • camera
  • Global Positioning System (GPS) Unit

* It is helpful to copy field sheets onto water-resistant paper for use in wet weather conditions

7.2

MULTIHABITAT APPROACH: D­FRAME DIP NET

 Streams in many states vary from high gradient, cobble dominated to low gradient streams with sandy or silty sediments. Therefore, a method suitable to sampling a variety of habitat types is desired in these cases. The method that follows is based on Mid-Atlantic Coastal Streams Workgroup recommendations designed for use in streams with variable habitat structure (MACS 1996) and was used for statewide stream bioassessment programs by Florida DEP (1996) and Massachusetts DEP (1995). This method focuses on a multihabitat scheme designed to sample major habitats in proportional representation within a sampling reach. Benthic macroinvertebrates are collected systematically from all available instream habitats by kicking the substrate or jabbing with a D-frame dip net. A total of 20 jabs (or kicks) are taken from all major habitat types in the reach resulting in sampling of approximately 3.1 m2 of habitat. For example, if the habitat in the sampling reach is 50% snags, then 50% or 10 jabs should be taken in that habitat. An organism-based sub sample (usually 100, 200, 300, or 500 organisms) is sorted in the laboratory and identified to the lowest practical taxon, generally genus or species.

 

7.2.1

Habitat Types

 

The major stream habitat types listed here are in reference to those that are colonized by macroinvertebrates and generally support the diversity of the macroinvertebrate assemblage in stream ecosystems. Some combination of these habitats would be sampled in the multihabitat approach to benthic sampling.

 

Cobble (hard substrate) - Cobble will be prevalent in the riffles (and runs), which are a common feature throughout most mountain and piedmont streams. In many high-gradient streams, this habitat type will be dominant. However, riffles are not a common feature of most coastal or other low-gradient streams. Sample shallow areas with coarse (mixed gravel, cobble or larger) substrates by holding the bottom of the dip net against the substrate and dislodging organisms by kicking the substrate for 0.5 m upstream of the net.

Snags - Snags and other woody debris that have been submerged for a relatively long period (not recent deadfall) provide excellent colonization habitat. Sample submerged woody debris by jabbing in medium-sized snag material (sticks and branches). The snag habitat may be kicked first to help dislodge organisms, but only after placing the net downstream of the snag. Accumulated woody material in pool areas are considered snag habitat. Large logs should be avoided because they are generally difficult to sample adequately.

Vegetated banks - When lower banks are submerged and have roots and emergent plants associated with them, they are sampled in a fashion similar to snags. Submerged areas of undercut banks are good habitats to sample. Sample banks with protruding roots and plants by jabbing into the habitat. Bank habitat can be kicked first to help dislodge organisms, but only after placing the net downstream.

Submerged macrophytes - Submerged macrophytes are seasonal in their occurrence and may not be a common feature of many streams, particularly those that are high-gradient. Sample aquatic plants that are rooted on the bottom of the stream in deep water by drawing the net through the vegetation from the bottom to the surface of the water (maximum of 0.5 m each jab). In shallow water, sample by bumping or jabbing the net along the bottom in the rooted area, avoiding sediments where possible.

Sand (and other fine sediment) - Usually the least productive macroinvertebrate habitat in streams, this habitat may be the most prevalent in some streams. Sample banks of unvegetated or soft soil by bumping the net along the surface of the substrate rather than dragging the net through soft substrates; this reduces the amount of debris in the sample.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Methods ~ Surber

 

1. Site selection should include visibly moving water as well as a riffle, with a water level no higher than the top of the net

2. The Surber net should be set about 1 foot downstream from the riffle

3. The net is supported by the first person while the second person uses a scrub brush to rub the rocks in a 1 foot by 1 foot area in front of the net

4. The net is removed from the water; debris is rinsed to the bottom of the net using the current

5. The collecting bottle is unscrewed and the contents placed in a plastic container which is then filled past the level of the contents with 70% ethanol

6. Plastic containers should be labeled with monitors last name, site locations, date, and contents

 

 

 

 

 

 

Appendix III

 

1. Benthic Macroinvertebrate Sample Login Sheet

2. Sample Scoring Criteria

3. List of Potential Benthic Invertebrates

4. Standardized Macroinvertebrate Data Sheet

5. Tips and Pictures for Macroinvertebrate Analysis

 

 

 

 

 

Benthic Macroinvertebrate Sample

Log-in Sheet

Explanation of Categories

 

Date Collected – date the sample was collected

 

Collected by – collectors last name

 

Preservation – what the sample was preserved with (ethanol or formalin)

 

Stream name and Station ID – name of the stream and the sample id number

 

Location in lab – write the name of stream cabinet the sample is in

 

Sub-sampling protocol – limestone or freestone

 

Sub-sample date and initials – date of completion of sub-sampling and the initials of who did the sub-sampling

 

ID date and initials – date of completion of identification and the initials of who did the identification

 

Archived – mark if the sample was kept and preserved for a reference

 

 

 

 

 

Scoring Criteria


 

 

 

 

 

Potential Benthic

Invertebrates



 

 

 

 

Standardized Data

Sheet