DEP uses the following procedures when conducting field sampling for freshwater
- Two electrofishing passes are conducted at each 75m site to obtain abundance data and to measure the ecological health of the fish community. The condition of the fish community in the stream reach will be determined from this sample.
- All long-term baseline and reference
stream freshwater fish inventories will be conducted between June
1 through the middle of October when spawning and migration effects
are minimal (Kazyak and Jacobson 1994). Sampling when flow variability
is low increases the likelihood that samples will be collected
under similar conditions (Jacobson and Kazyak 1993). In the event
of inclement weather, sampling should be conducted one to two
days after rain events that cause the stream to rise above baseflow channel bars. Sampling should be done two to three days after
bankfull storm events, or when water recedes to baseflow levels
and water clarity returns.
- At least one biologist will have a valid State collecting permit. For safety reasons a minimum two-person team will be used at all times. All members of the sampling team should use polarized glasses to reduce glare and improve capture probability (Ohio EPA 1987; Meador et al. 1993; and Kazyak and Jacobson 1994).
- Channel block nets are securely placed across the stream at both ends of the inventory segment. Block nets should be placed avoiding unnecessary stream disturbances. Gaps between the lead line and the stream bottom should be filled with streambed material (i.e. rocks). The ends of the block net should be firmly anchored to each stream bank, using rebar if necessary. It is essential that the assumptions of removal sampling (i.e. no immigration or emigration during the removal sampling) be maintained.
- All quantitative sampling will be done with a backpack electrofishing unit with programmable output waveforms (POW) (e.g. Model 12-A Smith Root or equivalent). POW electric pulse was selected to minimize skeletal damage to stream fish. Users will adjust the output based on the conductivity levels measured in the stream segment (Table 1). Conductivity should be measured with a conductivity meter following manufacturer's instructions for calibration and use.
Table 1. Recommended Voltage Settings for Smith Root Model 12-A Backpack Electrofisher based on Ambient Conductivity Levels
||400 to 500
|100 to 400
||200 to 400
||100 to 200
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Smaller streams with low baseflow will require lower voltage settings. The user should start at the lowest voltage settings on the above table. In all cases, electrofishers should be set up by observing fish behavior and recovery times, not by voltage or current measurements.
Student intern sets up live wells with bubblers to oxygenate the water and keep the sample fish alive during the process.
- Two passes will be made in an upstream direction within the station segment (from the 0m end to the 75m end). The entire segment should be electrofished, including any backwater areas, and shallows (Kazyak and Jacobson, 1994). For every three meters of wetted width there needs to be an additional electroshocker. All fish will be netted and removed during each pass, with fish caught kept in five gallon buckets. Each bucket containing fish will be equipped with a portable oxygen bubbler.
To minimize mortality, fish will be identified after each pass has been completed. Identified fish can either be kept in a float box (or live well), with the float box anchored well outside of electrofishing range in a shaded run area or immediately released back to the stream outside of the sampling segment. Eels may be kept to prevent them from reentering the sampling segment. The float box should have the end facing the current elevated so oxygenated water can flow through the box. Fish are kept in the float box until the station has been completed. Total number of individuals of each species per pass will be recorded on a field data sheet. The bottom block net should be thoroughly checked for fish after the second pass and be added to the second pass fish counts. Fish will be returned to the station reach after completion of the inventory.
Fish are identified, inventoried and immediately released.
The identification and inclusion of young-of-year (YOY) fish in the sample has not been recommended by other agencies (Ohio EPA 1989, Plafkin et al. 1989; Barbour et al. 1999). These fish are generally <15 to 20 mm in length. The collection technique and difficulty in field identification was not found to be consistently effective for all species (Ohio EPA 1989), raises sampling costs and increases the need for laboratory analysis (Karr et al. 1986). Including YOY has had large effects on IBI scores in years and habitats in which YOY were especially abundant and probably introduces a larger margin of error in IBI interpretation (Angermeier and Karr 1986). DEP identifies all adult fish (approximately >30mm). Most YOY fish in a stream segment will not be captured due to their surface to volume ratio; they recover quickly, avoid capture, and may pass through 1/4" mesh. Toward the later months of the fish sampling period, some YOY species will grow beyond the 30mm guideline. Staff will use their professional judgment during sampling to determine YOY. Examples of some of these species are sunfish, bullhead, and creek chubs. All YOY salmonids will be identified, measured and weighed in the field.
Measuring young of year game fish (Smallmouth Bass).
Weighing sample salmonid fish in the field (Brown Trout).
Gross external pathology and anomalies will be observed and recorded when the fish are identified for each pass to minimize repeated handling of the fish. The body surface and fins will be examined for discoloration, hemorrhaging, cloudiness, raised scales, scale deformities, spots, visible external parasites, growths, ulcerations, fin erosion, and other abnormal conditions. Eyes will be examined for cloudiness, hemorrhaging, exophthalmia (popeye), cataracts, or orbital depression (Kazyak and Jacobson 1994).
Fish are examined for anomalies when they are identified.
- Biomass will be recorded at all sites in the eastern piedmont (this includes all watersheds east of Great Seneca). Biomass should be measured after each pass. A scale is used for this process and weight is taken to the nearest tenth of a gram. If there are substantial amounts of fish, more than one measurement may be taken. It is important not to injure or kill the fish in this procedure.
- Additional sampling procedures must be followed for sites that contain trout. All trout are anesthetized with a 1:10 solution of clove oil and ethanol alcohol, identified to the species level, measured for total length (TL) to the nearest millimeter, weighed in grams, and returned alive to the stream at the end of the survey. The pelvic fins of all trout will be examined for fin wear to determine if it is a hatchery or wild fish. Note any hooking or predation injuries. Any trout that are removed from the station must be returned to the microhabitat from which it was collected. All trout data will be submitted to Maryland Department of Natural Resources (MD-DNR), Freshwater Fisheries Division Region III staff.
Representatives of any species unable to be identified will be placed in labeled Nalgene containers with 10% formalin solution and taken to the laboratory for identification. The label will include the date, watershed, stream segment identification number, pass number, and the name of the collector.
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- 1 POW backpack electrofishing unit, or equivalent, per 3m of wetted width.
- 1 anode ring and pole per backpack electrofishing unit
- 1 cathode ("rat tail") per backpack electrofishing unit
- 12-volt rechargeable batteries
- 5-gallon buckets
- 1 oxygen bubbler per fish holding bucket
- 1 fish measuring board (in. and cm)
- 1 10kg scale (for biomass sites)
- 1 2kg scale (for trout streams)
- Field data sheets, pencils
- Hip chain
- 1 dip net per person (1/4" stretch mesh)
- Hip boots or chest waders
- Polarized sunglasses
- Cell phone
- Nalgene container with 10% formalin
- ID Labels for formalin containers
- TMS-fish tranquilizer or equivalent (for trout streams)
- Aluminum tree tags
- Collection permit
- 2 or more block nets
- Field identification aids
- First Aid Kit
- Multi-parameter probe that measures conductivity, pH, temperature, and dissolved oxygen (HydroLab, YSI, etc.)
- Staple gun (extra staples)
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DEP uses the following procedures when conducting laboratory experiments for
- Fish preserved for subsequent identification or for watershed vouchers are immersed in a buffered 10% formalin solution as soon as possible after capture. Large specimens should be slit on the right side prior to preserving. Vouchers will remain in buffered formalin for at least two weeks for fixation. The vouchers will then be immersed in tap water for twenty-four hours. After this period, the water is discarded and 70% ethanol is added to the container for permanent storage. Unidentified specimens can be examined and identified at this stage.
- Fish are identified using published keys for the Mid-Atlantic region. Some of these keys may include: Schwartz (1963); Loos et al. (1972); Davis (1974); Lee et al. (1976); Eddy and Underhill (1978); Lee et al. (1981); Knopf (1983); Cummins (1987); Page and Burr (1991); and Jenkins and Burkhead (1994); Raesly and Kayzak (2003). Scientific nomenclature follows the recommendations of the American Fisheries Society (Robins et al. 1991). The use of three or more publications is recommended to verify identification and reduce the possibilities of false identification.
- DEP staff verifies identifications. After taxonomic verification is completed, the information is transferred to the appropriate fish data sheet. If identification questions remain, the specimen will be taken to either the office of the Maryland DNR Freshwater Fisheries Division Region III biologist or the MBSS for identification.
- A cumulative list of freshwater stream fish found in Montgomery County will be maintained by DEP. Additions to the cumulative list are invited, and will be accepted with the submission of a properly identified voucher specimen. Required submittal information should include name of researcher, date of collection, location of collection, and identification of the specimen.
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