Aquatic Invertebrate Community Study
Total Page：16
File Type：pdf, Size：1020Kb
..,, ------------------------- U. S. Fish & Wildlife Service Aquatic Invertebrate Community Study at Prime Hook NWR Bombay Hook NWR Long Island NWR Complex Supawna Meadows NWR Summer2000 Prepared by: Cassy Lewis Lisa Johnson NON-TARGET AQUATIC INVERTEBRATE COMMUNITY STUDY 2000 Prime Hook National Wildlife Refuge Bombay Hook National Wildlife Refuge Long Island National Wildlife Refuge Complex Supawna Meadows National Wildlife Refuge PURPOSE The immediate goal of this study in the summer of 2000 was to collect baseline data on the aquatic invertebrate communities present in wetlands where mosquito control activity occurs, or may occur in the future, on National Wildlife Refuges. We surveyed the invertebrate communities characterizing the water column and the benthos at four refuges in Delaware, New Jersey, and New York. This study is a component of the long term attempt to minimize the potential negative effects of mosquito control activities, particularly the use of larvicides, on the 1ion-target invertebrates that share the habitat of mosquito larvae. Knowledge of the invertebrate taxa that are present (or abundant relative to other taxa) at a site will assist refuge staff in mosquito control decision making, particularly in choosing specific larvicides that will minimally impact a site's particular non-target invertebrates. Applying information about the local invertebrate communities to the selection of appropriate larvicides is an important step toward conserving invertebrate biodiversity and maintaining this important food resource for other species that use the wetlands. METHODS General Approach The four refuges that were sampled during July and August of 2000 were Prime Hook NWR, Bombay Hook NWR, Long Island NWR Complex (Wertheim NWR), and Supawna Meadows NWR (Table 1). At each refuge, we sampled three sites representing different habitat types where mosquito breeding was likely to occur. At ten to sixteen sample locations per site, we collected a benthic core sample and a water column sample, counted mosquito larvae/pupae present in three dip samples, and recorded vegetation/habitat type and water parameters (water temperature, water depth, pH, and salinity). In the lab, we sieved and sorted ten benthic and ten water column samples per site, counted and identified (to family-level) all aquatic invertebrates found, and dried and weighed the invertebrates. Any fourth-instar mosquito larvae that we found were identified to species level. Detailed Procedures Study Site Selection: The refuge biologists at each NWR that participated in the study identified three sites for sampling. We used sites that were representative of areas where mosquito control activity occurs or may occur. If both salt marshes and fresh water wetlands were present at the NWR, we suggested that the three sites include both habitat types. Some biologists included in their choices a pair of two similar sites, one that had been sprayed with mosquito larvicides and one that had not, for comparative purposes. The refuge staff provided maps of the area, and insecticide spraying history information (Table 1). Depending on transportation time, sampling each site took between 1.5 and 3 hours. Once we arrived at a site, we selected a semi-discrete area to use as our sampling area. If the site was a salt marsh, we chose a representative portion of the marsh to sample that contained at least IO different potholes 01: wet areas. If the site was an open water body, we chose a sampling area that was bounded by terrestrial vegetation and that could be sampled in an afternoon or morning period of time. Field Data Collection: We selected a center point for each sampling area, and took a GPS location (recording the waypoint on the plugger unit for later downloading). Our pattern for sampling was to go out in 3 directions from the center point in radiating transects. We randomized our selection of sample locations at each site by spinning the compass dial to come up with an initial directional reading for the first transect. 120 and 240 degrees were then added onto the initial reading for the other transect directions. We took a GPS location at the end of each transect to define our sample area boundaries. 2 In order to space out sample points on the transect, we estimated the number of paces needed to evenly fit 3 to 5 points l:,etween the center and the perimeter of our area, with our perimeter sampling point ending up in the edge vegetation if we were in open water. We attempted to have at least 5m between sample points to minimize disturbing subsequent sample point areas. The following list details the data we collected in the field: 1. At each site, we recorded air temperature, sky condition (Sun/Rain/Cloudy/Partly Cloudy), and estimated wind speed. 2. We assessed the vegetation present at each site and chose one of the following to describe vegetative type: 0 = Open Water EV Emergent Vegetation p Salt Marsh Pothole (the nearest pothole to where the paced sample point ended up falling) SAV = Submerged Aquatic Vegetation All sample points at a site were usua!ly, but not always, the same vegetation type. We recorded vegetation types for individual sample points when they differed from the rest of the site. 3. Once arriving at a sample point, we took 3 separate dips for mosquito larvae before disturbing the area. We recorded the average number of mosquito larvae and pupae per dip, and assigned a percent for each developmental category present (instars 1, 2, 3, 4 and pupae) in the three dips. We collected samples of the fourth instar larvae and pupae and placed them in whirl-pak bags for later identification. 4. We took a water column sample using the D-frame net by sweeping above the benthos for 1 meter in length. If the water was too shallow to get the D-frame net through the water column, we used the 3 50-ml dipper to collect 10 dips of water, which we poured through the D-frame net. If there was less water than this present (such as in a small salt marsh pothole), we poured a total of 1 dip of water (obtained by getting small amounts of water in one dipper, and filling the sec.:ond dipper) through the D-frame net, and recorded that this was done. We placed the contents of 3 1 the net in a labeled zip-lock bag. Ifwe found a fish in the sample, we made a note of this and returned the fish to the water. (We also noted whether we observed evidence of fish in the water continuous with our sample point.) 5. We then took readings for water temperature (nearest degree Celsius), pH (±0.1 pH, using a Hanna Instruments pHep 3 Pocket-Sized Microprocessor), salinity (nearest part per thousand, using a Sper Scientific Salt Refractometer w/ATC 300011), and water depth (nearest whole centimeter). We took depth and temperature readings at every sample point. If the body of water was homogenous and connected, we collected pH and salinity data only three times if three consistent readings were obtained. However, if discrete potholes of water were encountered, we measured pH and salinity at each sample point. We calibrated the refractometer at the beginning of the season, and calibrated the pH meter prior to each trip. 6. Next we took the benthic core sample, using a plastic PVC core with a screen on one side. The core measured 5 cm in depth and 10 cm in diameter. The core was placed on the bottom and pushed into the benthos until the depth of 5 cm was achieved. We put the contents of the sample into a labeled zip-lock bag. When the root mat was too tough to dislodge by hand, we went around and under the core with a serrated trowel to get the sample. Storage o(Samples: While on the road, we kept the sample bags on ice in coolers. We put them into the refrigerator upon returning to the lab. If the samples had to be stored for over 1 week, we added enough anhydrous reagent alcohol (95% denatured alcohol formula 3A, 5% Isopropyl alcohol) to the bags to coat all of the material. [Before using alcohol to preserve the samples, one sample was kept refrigerated for eight days. Some soft-bodied insects (damselfly larvae and mosquito pupae) were beginning to decompose - other insects, including corixids and beetle larvae, were still whole. The water column samples tended to decompose more quickly than the benthic samples.] Processing Samples: We rinsed material from each sample in a 1 mm - mesh sieve, and placed the remaining material into white sorting trays. Concentrated sugar water (one five-pound bag of sugar per gallon of water) was poured into each tray with the sample, in order to 4 aid in sorting. Most invertebrates floated to the surface of the sugar water; however, snails remained on the bottom. Large samples (usually benthics) where split in half or thirds and placed into separate trays, so that the white tray bottom was visible during sorting as a contrasting background. Each tray of material was searched under light with forceps by teasing apart one clump at a time, and all macro-invertebrates were picked out. If no organisms were observed, we ceased searching that tray-load of material after 15 minutes. Thus, each tray received at least 15 minutes of scrutiny, and large benthic samples were sorted through for at least 30 total minutes. Using dissecting microscopes and keys, we identified each insect invertebrate individual to family level. We identified fourth instar mosquito larvae to species level, and if any adults developed from pupae in our bags, we also identified them to species. We identified annelids, nematodes, water mites and snails to more general taxonomic levels (as far as was possible with our resources).