A Baseline Assessment of Environmental and Ecological Conditions in the May River,

Beaufort County, South Carolina

                                                         

Submitted By:

 

Marine Resources Division

South Carolina Department of Natural Resources

217 Ft. Johnson Rd.

Charleston, SC 29412

 

in cooperation with the

 

United States Geological Survey

Water Resources Division, SC

720 Gracern Rd., Suite 129

Columbia, SC 29210

 

and the                                                            

 

National Ocean Service

National Oceanic and Atmospheric Administration

219 Ft. Johnson Rd.

Charleston, SC 29412

 

 

 

 

 

 

July 2001

 

 

 

 

Background and Rationale for Study

Over the next several decades, the coastal watersheds in the southeastern United States, especially those in Beaufort County, are projected to experience a high rate of human development (Edwards, 1989; Culliton and others, 1990; Cohen and others, 1997). The construction of infrastructure (roadway systems, commercial development, residential housing, and industrial facilities) that accompany human development will alter the rate and volume of freshwater inflow as well as the type and amount of pollutants introduced into estuaries (Driver and Troutman, 1989; Fulton and others, 1993; Mallin and others, 2000; Lerberg and others, 2000). Estuarine waters of particular concern are the tidal creeks and rivers, like the May River system (Fig. 1), that have little potential for diluting the effects of anthropogenic inputs. These creeks and small river drainage basins provide nursery habitat for many species of fish, shrimp, and crabs as well as feeding grounds for wading birds and adult fish (Weinstein and others, 1980; Wenner and Beatty, 1993; Dodd and Murphy 1996). Hydrologic processes in tidal creeks also trap fine-grained sediments, organic material, and chemical contaminants and serve as conduits and repositories for pollutants (Biggs and others, 1989; Sanger and others, 1999a; 1999b). During high tide, juvenile shrimp, crabs, and fish use the surface of the salt marshes and tidal flats as feeding grounds and refugia. At low tide, these biota are concentrated in the tidal creeks and rivers. Tidal creeks also serve as a primary pathway through which non-point source pollutants are delivered from terrestrial sources to the larger estuarine drainage systems.

 

In response to concerns about coastal development and the subsequent closure of large areas of shellfish beds in Beaufort County due to poor water quality, a group of concerned citizens formed the “Clean Water Task Force” (CWTF). The CWTF convened several meetings to discuss and evaluate concerns related to urban development and other land use practices on wetlands in Beaufort County. They summarized their findings in a final report entitled, A Blueprint for Clean Water. Strategies to Protect and Restore Beaufort County’s Waterways (Clean Water Task Force, 1997). This report listed several major steps and other recommendations that the County and its municipalities should undertake to improve Beaufort County water bodies, with assistance from appropriate state and other government agencies.

 

Figure 1.    Map of May River Study Area

 

One major recommendation was to perform baseline assessments of existing conditions in areas of concern. A high-priority area of concern identified at that time was Broad Creek and Okatee River. In response to this need, Beaufort County and the South Carolina Department of Health and Environmental Control (SCDHEC) and South Carolina Department of Natural Resources (SCDNR) supported a comprehensive baseline assessment of environmental and biological conditions in those two drainage systems. That study provided critical data on the water quality, sediment quality, and key biological resource conditions that could be used for comparison with future surveys (Van Dolah and others, 2000). The study also provided several recommendations that could help to improve conditions in those water bodies.

 

The May River represents a significant estuary in Beaufort County that may be adversely affected by planned developments. The Town of Bluffton (population about 1,200) is expected to undergo significant land-use changes in the next few years as residential and retirement communities continue to develop in the area. The May River is a treasured resource of this small coastal community. With its outstanding water quality, the river is a valued fisheries and shellfish nursery habitat. Oysters from the May River have historically been a significant economic resource for the area, as demonstrated each fall with the Bluffton Oyster Festival. Currently, the SCDHEC has designated the May River as an Outstanding Resource Water (ORW). Residents of Bluffton and the surrounding area are concerned that the health of the May River and its oyster beds might be compromised. by anticipated development in the area, With the exception of a few stations that have been sampled by SCDHEC and SCDNR as part of existing monitoring programs, limited data are available on the current condition of this estuarine river.

 

The proposed study is designed to provide a comprehensive baseline assessment of the May River estuary as a cooperative effort between the SCDNR Marine Resources Division, the U. S. Geological Survey (USGS), the National Oceanic and Atmospheric Administration’s (NOAA) National Ocean Service Charleston Laboratory, and the SCDHEC Bureau of Water. The study design will be similar to that used in the Broad Creek– Okatee River Environmental Study (Van Dolah and others, 2000), but will provide a substantially larger water-quality baseline. To maximize cost savings, we propose to conduct much of the primary sampling activities using protocols similar to those being used in two ongoing monitoring programs. These include a new comprehensive state-wide monitoring program of estuarine habitat quality entitled the “South Carolina Estuarine and Coastal Assessment Program” (SCECAP), which is a cooperative program being conducted by the SCDNR and SCDHEC (SCDNR, unpublished). Numerous sites are sampled each year throughout Beaufort County for the SCECAP study. These sites can be used to compare with conditions in the May River. The other ongoing SCDNR study is the Tidal Creeks Project (TCP) which is currently conducting monitoring in tidal creeks of the Okatee River through funding from SC Sea Grant’s Land Use – Coastal Ecosystem Study (LU-CES). 

 

General Sampling Design and Scope of Work

Three general zones representing the upper (zone 1), middle (zone 2) and lower (zone 3) portions of the river (approximately 7-kilometer-long segments of the May River) have been identified for sampling (Figure 2). Within each zone, two subtidal stations will be randomly located in the mainstem of the river for open-water sampling and oyster bar investigations. One to two randomly located subtidal stations in each zone will be sampled. For the purposes of this study and the SCECAP program, a large tidal creek represents estuarine water bodies of less than 100 meters (m) in width from marsh bank to marsh bank and greater than 3m deep at mean high water. In addition, two headwater tidal creeks will be intertidally sampled within each zone. For the purposes of this study and the TCP, a headwater tidal creek is defined as the point where water depth in the channel is approximately 1 m deep at mean high water and continuing down the creek for 600 m.  This sampling regime yields 16 sampling areas (Fig. 2). All stations will be located using Global Positioning System (GPS) equipment.

 

 

 

 

 

 

Figure 2.   Candidate Sampling Sites

 

X
 

     USGS Continuous Stns.     Headwater Creeks     Large Creek Sites     Open-water Sites
 

X
 

X
 

X
 

X
 

Zone 2
 

Zone 3
 

Zone 1
 

X
 

X
 

 

 

During summer 2001 (USGS FY01), USGS and SCDNR will perform reconnaissance to identify and locate six sampling sites within the headwaters of the tributary creeks of the May River. The SCDNR will locate 6 sampling sites within the mainstem of the May River and 4 sites within the associated tidal creeks. During the same period, USGS will install and commence operation of three continuous water-monitoring instruments (described later in this document). Data collection by these three instruments will continue for a minimum of 24 months, with a post-study option available to the cooperator to continue for a longer period.

 

From fall 2001 to summer 2002 (USGS FY02), USGS will collect water-quality and phytoplankton samples, at least quarterly, at each of the 16 sites. Additional water-quality samples will be collected during the summer months (June, July, August) which is the period when water quality problems can be most severe and when many of the estuarine biota of concern are using the creeks and river as nursery habitat. This season is also consistent with the SCECAP program, which will provide comparative data. In addition, USGS will collect quarterly water samples for the enumeration of bacteria (E. coli) at each of the 6 headwater primary sites. Additional collections for bacterial enumeration may be made at a frequency determined by weather and tide conditions, in order to gain a more complete picture of the relative contributions of bacterial contamination by each tributary’s drainage area, under a variety of conditions.

 

Biological- and sediment-quality sampling, and water quality monitoring will be conducted by SCDNR during the summer of 2002 (USGS FY02 Q4). Sampling of the biota (fish, benthic invertebrates), and sediments, as well as water quality monitoring will be conducted once in each of the 16 sites. Oyster bed sampling will be conducted in 2-4 representative beds in each zone of the mainstem of the May River. All sampling will be coordinated among cooperating agencies to ensure maximum cost savings. A general description of the methods to be used is provided for each major program element.

 

USGS Water Quality Monitoring Program

Water-column samples will be collected quarterly for the determination of nutrient and suspended sediments concentrations as well as for the determination of the phytoplankton community, which will be analyzed by SCDNR. Two different sampling methods will be employed depending on whether the samples are taken in the headwaters of tidal creek or in the subtidal tidal creek and open-water sites.

 

United States Geological Survey methods for the collection of water-column samples in streams (in this case, headwater tidal creeks) require that isokinetic samples be collected across the entire width of the stream or creek at equally spaced points (equal width increment or EWI), when the discharge is greater than 1.5 cubic feet per second (ft3/sec). An isokinetic sample is collected by a compositing device that results in a sample that is proportional in volume to the amount of discharge at each point in the stream or creek (U. S. Geological Survey, 1999). If the discharge is less than 1.5 ft3/sec, a dip or discrete (point) sampling method will be used. Water samples at each site will be composited into a single sample, using a USGS-designed cone splitter. The splitter apportions samples into containers specific for each type of analysis. These procedures involve “parts per billion” sampling techniques designed to eliminate sources of contamination. Samples collected in the tributaries will be analyzed for the same suite of components as the water column samples collected from the May River (listed below).

 

Water-quality samples will be collected in the subtidal tidal creeks and open-water May River mainstem sites by inserting pre-cleaned water bottles to a depth of 0.3 m, inverting the bottles, and then filling them directly at that depth.

 

All near-surface water-quality samples collected in the May River system will be analyzed for concentrations of total nutrients, total organic carbon (TOC), total alkalinity, and turbidity. In addition, sub-samples will be shipped to SCDNR for phytoplankton composition measurements as described later in this document.

 

The bottles will be stored on ice until they are preserved either in the field or immediately upon return to the laboratory (nutrient and TOC samples only). Sampling protocols follow standards described by the SCDHEC (1997) and/or the SCECAP/Coastal 2000 QAPP for sampling, specific to the Coastal 2000 program (SCDNR, unpublished). The dissolved nutrient samples will be collected in triplicate and filtered through 0.45 micrometer (um) pore cellulose acetate filters prior to preservation and delivery to the laboratory. At the time of sample collection, measurements of dissolved oxygen, pH, specific conductance, and temperature will be made using a HydroLab® DS-3 or DS-4 datasonde attached to a Scout 2[1].

 

Laboratory processing will be completed at the USGS National Water Quality Laboratory (NWQL) using standardized U.S. Environmental Protection Agency (USEPA) - accepted procedures. The following constituents and parameters will be quantified: total nitrate/nitrite nitrogen, total Kjeldahl nitrogen, ammonia, total phosphorus, total organic carbon, total alkalinity, dissolved nitrogen species (NH4, NO3/NO2), PO4, dissolved organic nitrogen (DON), dissolved organic phosphorous (DOP), dissolved organic and inorganic carbon (DOC, DIC), dissolved silicon/silica, turbidity, biological oxygen demand (BOD5), total suspended sediments (TSS), and five-day biochemical oxygen demand.

 

The SCDHEC currently collects monthly samples for bacterial enumeration at 11 sites in the May River, as part of a shellfish monitoring and management program. Data from several years past are available, and future data will be made available. Therefore, for purposes of this study, extensive sampling for bacterial enumeration will not be required. However, collections at selected tributary headwater sites are warranted during this study to focus on specific bacterial contributions by those creeks. Such collections will follow standard USGS- or EPA-approved protocols (for example, a maximum 6-hour, holding time, on-ice, before inoculation of m-TEC nutrient plates, and incubation at 35 degrees Celsius for 2 hours, followed by 22 to 24 hours at 44.5 degrees Celsius).

 

Continuous monitoring

Three sites (one in each of the three zones; see fig. 3) will be selected for continuous monitoring of tidal streamflow, water level, temperature, specific conductance, and dissolved oxygen concentration. Water-level and flow data will be collected at 15-minute (min) intervals. Tidal stream flow will be determined by using an acoustic Doppler velocity meter. Continuous tidal creek flows will be determined by relating cross-sectional area to a mean velocity at each site using methods described by Simpson and Bland (1999), Lipscomb (1995), Laenen (1985), and Rantz and others (1982). Flow measurements will be made at each site to relate the measured index velocity to the mean velocity. Measurements of water temperature, specific conductance, and dissolved oxygen concentration will be made on a 30-min interval. Field operation, calibration of water-quality monitors, and the computation of water-quality continuous records will follow procedures described by Wagner and others (2000). One gaging station will include a precipitation gage.

 

The continuous monitoring sites will be incorporated into the real-time gaging network of the South Carolina District of the USGS. Data will be relayed via the GOES satellite to the S.C. District Office, at four-hour intervals. Provisional data will be available on the USGS, South Carolina District web page http://wwwsc.er.usgs.gov/ and will be updated on a four-hour interval.

 

Continuous monitoring of streamflow and physical water-quality parameters generates a time series of responses to the riverine/estuarine system due to meteorological changes that are difficult to measure by using a periodic sampling schedule. From the continuous streamflow record for the three sites, a water budget will be determined to calculate the amount of runoff per unit area of the watershed. This measurement will be critical to an understanding of the transport of materials from the watershed and how the watershed is responding to changing meteorological and land-use conditions.

 

SCDNR/NOS Biological, Sediment and Water Quality Monitoring Program

SCDNR Water Column Phytoplankton Monitoring

Water samples will be collected each quarter in triplicate by USGS and transported immediately on ice to the SCDNR laboratory for analyses. Phytoplankton biomass, phytoplankton community, and harmful algal species enumerations will be made by SCDNR following standard protocols established for the SCDNR Harmful Algal Bloom (HAB) Project. Estimates of phytoplankton biomass will be made by using chlorophyll-a measurements on a fluorometer. High Performance Liquid Chromatography (HPLC) pigment analysis also will be conducted. This analysis is an effective method for analyzing phytoplankton community composition because it offers advantages over standard microscopic techniques in that numerous samples can be processed relatively rapidly, and the potential for subjectivity in phytoplankton identification is reduced. Dr. Alan Lewitus has applied several modifications to current HPLC pigment analytical methods to develop a protocol that optimizes the detection of 20 pigments of known chemotaxonomic importance to algal identification (Van Heukelem and others, 1994; Van Heukelem and Thomas, 2001).

 

In addition, the SCDNR staff will analyze the water samples for the presence of harmful algal species. The SCDNR has generated a list of known estuarine phytoplankton that are classified as “harmful algae” based on precedence in forming blooms that cause adverse effects on ecosystems, fisheries, or humans. Accurate identification requires microscopic inspection of fresh, unpreserved samples. Precise and accurate quantification requires microscopic enumeration using fixed (preserved) water samples. In conjunction with the HPLC pigment analyses, the relative contribution of harmful algae to overall algal composition will be determined.

 

SCDNR Open-Water and Large Tidal Creek Monitoring

The water quality, biological, and sediment monitoring conducted by SCDNR will occur in the summer of 2002 (USGS FY02 Q4). Sampling at the six subtidal sites in the mainstem of the estuary and the four sites in the subtidal portions of the tidal creeks in each zone will follow standardized SCECAP sampling protocols. These protocols are briefly described below:

 

Water-quality measurements will be collected at all stations prior to deployment of other sampling gear. Instantaneous measurements will include near-surface, mid-depth and near-bottom measurements of dissolved oxygen, salinity, and temperature using Yellow Springs Instrument (YSI®) Inc. Model 85 water-quality meters and near-surface measures of pH using a pHep 3 field microprocessor meter. The near-surface measurements will be collected approximately 0.3 m below the surface and the near-bottom measurements will be collected from approximately 0.3 m above the substrate.  More complete time-profile measurements of all four parameters also will be obtained from the near-bottom waters of each site using either YSI® Model 6920 multiprobes or Hydrolab® DS-3 and DS-4 datasondes. Measurements will be logged at 15-min intervals for a minimum of 25 hours (hrs) to record readings over two complete tidal cycles. If possible, longer 96-hr time series may be used.

 

Secchi disk readings will be collected during water quality sampling. All readings will be taken to the nearest 0.1 m using a solid white disk with measurement protocols standardized to reduce or eliminate readings that may be affected by glare or surface wave chop.

 

Several replicate grab samples of sediments will be collected, by SCDNR personnel, at each station to evaluate sediment characteristics, sediment contaminant levels, and benthic community composition. A total of 8 to 10 grab samples will be collected at each site using a stainless steel 0.04 square meter (m2) [or 400 square centimeters (cm2)] Young grab sampler from an anchored boat. The boat will be repositioned after each sample to ensure that the same bottom is not sampled twice and to spread the samples over a 10-20 m2 bottom area. All grab samplers will be thoroughly cleaned prior to field sampling and rinsed with isopropyl alcohol and seawater between stations.

 

Three of the grab samples will be collected by SCDNR personnel for analysis of benthic community composition. These samples will be washed through a 0.5 millimeter (mm) sieve to collect the benthic fauna and preserved in a 10 percent buffered formalin-seawater solution containing rose bengal stain. The remaining grab samples will be used to obtain a sediment-composite sample for analysis of sediment composition, contaminants, and sediment toxicity. Only the surficial sediments [upper 5 centimeters (cm)] will be collected from these grabs and combined to produce a composite sediment sample, which will be thoroughly stirred and subdivided into separate containers for use in sediment bioassays (seed clam, Microtox® tests), sediment characterization analyses (sand and silt/clay composition), total organic carbon, porewater analyses (pH, salinity, and ammonia), and analyses of sediment contaminants (metals, organics). The composited samples will be kept on ice until taken to the NOAA National Ocean Service (NOS) Charleston Laboratory, and stored, either at 4oC (toxicity, porewater) or frozen (contaminants, sediment composition, TOC), until analyzed.

 

Benthic samples will be sorted in the laboratory to remove the organisms from sediments remaining in the sample. All organisms will be identified to the species level, or the lowest practical level possible if the specimen is damaged or incomplete. A reference collection is maintained at the SCDNR Marine Resources Research Institute.

 

Particle size analyses will be performed using a modification of the pipette method described by Plumb (1981). Percentages of sand, grain size greater than or equal to 63 um in diameter will be determined by separation through a 63 mm sieve. Silt/clay, grain size less than 63 mm, will be determined by timed pipette extractions. Pore water ammonia will be measured using a Hach Model 700 colorimeter and TOC is measured on a Perkin Elmer Model 2400 CHNS Analyzer.

 

Contaminants to be measured in the sediments and tissues are listed in Table 1. All contaminants will be analyzed by the NOS Charleston Laboratory using the following protocols. Extraction and sample preparation for organics are similar to those described by Krahn and others (1988) and Fortner and others (1996). Samples are then extracted with CH2Cl2 using accelerated solvent extraction [ASE], concentrated by nitrogen blow-down, and cleaned by gel permeation chromatography where necessary. Polynuclear aromatic hydrocarbons (PAHs) will be quantified by capillary GC-ion trap mass spectrophotometry (ITMS) and HPLC Organochlorine pesticides and polychlorinated biphenyls (PCB) are analyzed using dual column gas chromatography with electron capture detection (GC-ECD using methods described by Kucklick et al (1997). Trace metals will be analyzed using methods described by Long and others (1998) using inductively coupled plasma spectroscopy (ICP) for aluminum, chromium, copper, iron, manganese, nickel, tin, and zinc, and by graphite furnace atomic absorption for arsenic, cadmium, lead, selenium, and silver. Mercury concentrations will be analyzed by cold-vapor atomic absorption.

 

Sediment toxicity will be measured using two assays. The Microtox® assay utilizes the photoluminescent bacterium, Vibrio fischeri, to provide a sublethal toxicity measure that is based on the attenuation of light production by the bacterial cells when they are exposed to a toxic material. Solid-Phase Microtox® assays follow the protocols described by the Microbics Corporation (1992). Toxicity is based on criteria described by Ringwood and others (1997), which accounts for variations in response due to sediment composition. The seed clam assay involves exposing juvenile clams, Mercenaria mercenaria, to sediments for a 7-day period using protocols described by Ringwood and Keppler (1998). Toxicity is measured using both sublethal (growth rate) and lethal end points compared to exposure in control sediments.

 

Fish and large crustaceans (primarily penaeid shrimp and blue crabs) will be collected at each site following the benthic sampling to evaluate community composition, and beginning in 2002, to analyze for fish tissue chemistry and identify the presence of pathologies. Two replicate tows will be made at each site using a 4-seam trawl (18-foot (ft) rope, 15 ft head rope and 0.75-inch (in). bar mesh throughout). Trawl tow lengths will be standardized to 0.5 kilometer (km) for open-water sites and 0.25 km for creek sites. Tows will be made only during daylight hours with the current and speeds standardized as much as possible. Tows made in tidal creeks will be limited to periods when the marsh is not flooded (approx. 3 hrs + mean low water). This limitation also is generally applied to open water sites. Catches are sorted to lowest practical taxonomic level, counted, and checked for gross pathologies, deformities or external parasites. All organisms will be measured to the nearest 0.1 centimeter (cm) and weighed to the nearest 0.1 kg. When more than 25 individuals of a species are collected, the species is subsampled.

 

SCDNR Headwater Tidal Creek Monitoring

The water quality, biological, and sediment monitoring conducted by SCDNR will occur in the summer of 2002 (USGS FY02 Q4). Alternate sampling protocols will be used for the headwater portions of the creeks because these habitats are largely intertidal. The laboratory protocols will primarily follow the same procedures as described for the subtidal sampling and will not be reiterated below unless they differ. The intertidal sampling of the headwater tidal creeks will occur throughout the upper 600 m of each headwater creek.. Within this area, six benthic community samples will be collected in each creek with a 45.6-cm2 hand core at 1 m below mean high water. In addition, sediment samples will be collected for analyses of grain size, TOC, porewater ammonia, and chlorophyll-a, corresponding to each of the six benthic community samples in each creek. These ancillary data will be used to evaluate chemistry and the benthic community data. The surface sediment samples for chlorophyll-a analysis will be analyzed following Strickland and Parsons (1972) using a spectrophotometer.

 

One seine will be pulled in each creek for 25 m to evaluate the fish and crustaceans utilizing the tidal creeks. The seine sample will be located at approximately 500 m from the top of the defined creek. The fish and crustaceans will be preserved in formalin and processed to the lowest taxonomic level in the laboratory. At the primary benthic community sampling site in each creek, the following samples also will be collected: (1) sediment samples for chemical contamination; (2) sediment samples for toxicity tests (i.e., Microtox and seed clam); (3) one water sample for fecal coliform concentrations and typing; and (4) water samples to measure nutrient concentrations, turbidity, BOD5, fecal coliform concentrations and typing, and phytoplankton biomass.

 

Water quality will be monitored in each creek for four to five days prior to sampling for the other parameters. Water-quality measurements will be made using a Hydrolab or YSI instrument which collects temperature, pH, salinity, dissolved oxygen, and depth data at 30 min intervals.

 

SCDNR Oyster Bed Assessments and Recruitment Potential

Oyster beds are of special concern due to their value as a recreational and commercially harvested species. Oysters (Crassostrea virginica) serve an important ecological role as EFH since the beds form living reef structures that support a host of other associated organisms generally not found in surrounding sand or mud habitats (Coen and Luckenbach, 1999; Coen et al., 1999a; b). This is especially true in SC where seagrass (SAV) beds are absent. SCDNR staff have been examining size‑frequency relationships, recruitment potential, and disease (MSX and Dermo) levels of native oyster populations as indicators of habitat health, along with estimates of transient and selected resident species (Coen et al., 1999a; Coen and Luchenbach, 1999). These measures appear to provide an excellent indication of habitat health and oyster status and recruitment potential (Broad-Okatee Report). Therefore, we propose to use similar approaches in the May River study to:

 

Two to four representative beds in the May River drainage system adjacent to fringing marsh will be randomly selected in each zone. A 20-meter transit line will be placed along each shoreline at approximately mean sea level [level with densest oyster populations (approximately 4' below MHW)]. Five samples (5/site x 6-12 bed sites) will be collected at each site by placing a 0.143m2 quadrat at pre-selected random locations along the transit line. Digital photos will be taken prior to sampling for strata identifications. If no oysters are located within a pre-assigned random location, another random position will be chosen to ensure that five representative oyster samples are collected from each site. All oysters and sediment will be removed to a depth of ~11 cm and then placed in a labeled bag. The specific quadrat placement for each sample will be adjusted to maximize the percentage of oyster shell (live and dead) cover within the quadrat.

 

In the laboratory, 25 oysters will be removed from the above samples at each site for oyster disease (n = 25/site). These oysters will be measured for shell height so that they would be included in the remaining size frequency analysis. Bag contents will then be washed to remove sediment and non-molluscan biota. All mussels and crabs will be counted and scored categorically from 0-3 using preexisting abundance levels. Using calipers, each live oyster (including spat) will be measured for shell height (defined as the distance from the umbo to the outermost edge) to the nearest millimeter. All oysters will be examined to eliminate dead individuals.

 

The oysters designated for disease analysis will be examined for two common oyster diseases (Dermo and MSX). For Perkinsus marinus (Dermo), cell counts will be determined using Ray’s technique (Ray, 1952) and examined following the procedures outlined in Bobo and others (1997). For Haplosporidium nelsoni (MSX) identifications, prevalence will be determined from oyster histological sections fixed in Davidson’s AFA, embedded in paraffin and stained in Harris hematoxylin and eosin (Howard and Smith, 1983; Bobo and others, 1997).

 

A subset of oysters, approximately 20, also will be evaluated to assess subcellular level effects using three assays including lysosomal integrity, lipid peroxidation, and glutathione concentrations. A neutral red (NR) assay will be used to evaluate lysosomal integrity in the digestive gland cells of oysters (Lowe and others, 1992; Ringwood and others, 1998a). Cells with NR retained in lysosomes are scored as stabile and those with NR leaking into the cytoplasm are scored as destabilized. A minimum of 50 cells will be counted for each preparation, and the data will be expressed as destabilization indices (percent destabilized lysosomes per individual oyster).

 

The thiobarbituric acid (TBA) test will be used to measure lipid peroxidation (Gutteridge and Halliwell, 1990). Digestive gland tissues will be homogenized in 50 milli Molar potassium phosphate buffer (pH 7.0) and centrifuged (14,000 Revolutions per minute, 4oC, 5 minutes). A subsample of the supernatant will be mixed with tricholoroacetic acid containing TBA and butylated hydroxytoluene, heated at 100oC for 15 min and centrifuged to remove the precipitate. The resulting malondialdehyde (MDA) will be detected at 532 nanometers (nm) on a spectrophotometer. Standards will be prepared as described by Csallany and others (1984), and the data will be expressed as nanno moles (nM) MDA / gram (g) wet weight.

 

Glutathione concentrations of individual oyster digestive gland tissues will be determined by the DTNB-GSSG Reductase Recycling Assay (Anderson, 1985). GSSG reductase will be added and the rate of TNB formation monitored at 412 nm at 30-sec intervals for 90 seconds. Concentrations of GSH will be estimated from a standard curve and reported as nM GSH / g wet weight.

 

After the randomly selected quadrats have been collected, a minimum of 30 oysters will be collected by hand for tissue analysis from the mid-intertidal portion of the endemic reefs. Oysters will be larger than 7 cm in length when possible. Oysters will be stored on ice until transport to the laboratory. All contaminants will be analyzed by the NOAA National Ocean Service (NOS) Charleston Laboratory using the protocols described in Fortner and others (1996).

 

To evaluate recruitment potential and as a measure of integrated habitat growth (growth, mean size and post-settlement recruitment) we will be deploying replicate (n = 3) standardized trays of weighed oyster shell (‘cultch’) covered with large mesh for shell retention at least six sites for a period of 7-12 months. After recovery all live oysters will be measured to assess habitat ‘quality’ and oyster population recovery status.

 

NOS Supplemental Analyses of Fecal Coliform Bacteria

 

In addition to the basic fecal coliform measures, we recommend additional analyses be conducted of bacterial samples collected in the summer of 2002 to better document sources of these bacteria. A multi-tiered approach to differentiating sources of fecal coliform bacteria is proposed that includes: 1) Most Probable Numbers (MPN) and/or Membrane Filter for enumeration of fecal coliform bacteria; 2) Analytical Profiling Index (API) Biotyping to isolate E. coli bacteria; and 3) Multiple Antibiotic Resistance (MAR) of E. coli. This, in conjunction with the spatial watershed based sampling design, will be used to facilitate isolating sources with respect to land use density, land use type, and creek tributary.

 

Application of the MAR technique in the surface waters of South Carolina would be useful in differentiating animal and human bacterial pollution sources. The rationale for this approach is that bacteria from human sources will be more resistant to antibiotics than bacteria from wild animals since humans receive antibiotic treatments, and wild animals do not. Domestic animals are treated with only a limited number of antibiotics and will generally be intermediate in terms of MAR and will have different MAR patterns than humans. Dr. Geoffrey Scott and co-workers at NOAA are currently working with Dr. Mark Tamplin and co-workers at USDA to characterize samples from several South Carolina watersheds. In previous studies this method has been shown to be useful in differentiating point from nonpoint pollution sources in South Carolina (Van Dolah and others, 2000), Florida (Parveen and others, 1997) and Maryland (Kaspar and others, 1990). We propose to use MAR to differentiate E. coli bacteria isolated by API from positive fecal coliform samples collected from each site and known pollution source.

 

Each of the 16 samples collected in the summer of 2002 (USGS FY02 Q4) will be enumerated for fecal coliform densities using Most Probable Numbers or Membrane Filter methods. All positive samples will be further analyzed to isolate the E. coli bacteria portion. All positive E. coli bacteria will be further evaluated using MAR Testing. All sampling and analyses will follow appropriate chain of custody procedures for handling and tracking samples. Appropriate Quality Control/ Quality Assurance Methods (blanks and positive/negative controls) will be used for each analytical procedure (API, MAR). All original notebooks and data summaries will be maintained in secure areas and electronic data bases at NOAA/NOS/Charleston Center for Environmental Health and Biochemical Research (CCEHBR). Each method is briefly described as follows.

 

Most Probable Numbers of bacteria in the samples will be determined using Multiple Tube or Membrane Filter techniques (American Public Health Association, 1984; 1992). The resulting tubes will be picked, streaked onto Plate Count Agar (PCA), and incubated. Following incubation, some of the bacteria from each isolate will be inoculated into a saline solution to enable the exposure of the bacteria to each of the biochemical tests of the API. After another incubation, the APIs will be “read” to determine the API codes for each isolate. The API (bioMérieux Vitek, Inc., Hazelwood, MO) is a series of 21 biochemical tests placed on a strip. Each test is assigned a numerical value from which an API code is determined for the bacterial isolate being tested. The code can be found in an index that corresponds to a bacterial species (Escherichia coli, Klebsiella pneumoniae, Pseudomonas aeruginosa, and so on).

 

Escherichia coli isolates, confirmed using API, will be further tested for MAR following the method of Parveen and others (1997). Isolates will be transferred to a 96 well plate containing tryptic soy broth (TSB) and incubated. The broth cultures are then transferred in duplicate with a 48-prong replicator to Mueller-Hinton agar plates, each containing one of 10 antibiotics, or to a control plate without antibiotics. Plates are incubated. Resulting growth will be measured and compared to the size of the same isolate on the control plate (no antibiotic added). The following end points for MAR will then be determined: 1) Antibiotic Resistance = less than 15 percent reduction in colony size on the antibiotic plate compared to the control plate; and 2) Antibiotic Sensitivity = greater than or equal to 15 percent reduction in size compared to the control plate. Based on these colony measurements, the number of MAR indices will be calculated for each isolate.

 

Web Site

The USGS South Carolina District will place a web page on its public site that will provide up-to-date information on the progress of the project, contacts for questions, and data that has been through a quality assurance review process.

 

 

Report Preparation

A cooperative, integrated report will be prepared, describing all findings within 14 months of summer 2002 (USGS FY02 Q4) data collection. The cooperative report will document baseline conditions of the water quality, sediment quality, and living resources of the May River tidal creek ecosystem and compare those conditions with other typical, non-polluted sites in Beaufort County that are sampled in the same year. Included in the report will be the policies required by SCDNR to ensure that water quality and living resources of tidal creeks as habitat for representative important species are protected. An additional report will be prepared by USGS, at the discretion of the cooperator, for the continuous monitoring of water quality and streamflow that will occur in the fall of 2002, and winter and spring of 2003 (USGS FY03 Q1, Q2, Q3). The additional USGS report will document the longer term baseline conditions or potentially altered conditions in water quality and streamflow.

 

 

Selected References

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American Public Health Association, 1992, Standard Methods for the Examination of Water and Wastewater, 18th Edition. American Public Health Association, Washington, DC.

 

Anderson, M.E., 1985, Determination of glutathione and glutathione disulfide in biological samples. Methods in Enzymology, v. 113, p. 548-555.

 

Biggs, R.B., DeMoss, T.B., Carter, M.M., and Beasely, E.L., 1989, Susceptibility of U.S. estuaries to pollution. Aquatic Sciences, v. 1, p. 189-207.

 

Bobo, M.Y., Richardson, D.L., Coen, L.D., and Burrell, V. G., 1997, A report on the protozoan pathogens Perkinsus marinus (Dermo) and Haplosporidium nelsoni (MSX) in South Carolina shellfish populations, with an overview of these shellfish pathogens. SCDNR-MRD-MRRI Technical Report, Charleston, SC.

 

Clean Water Task Force, 1997, A blueprint for clean water. Strategies to protect and restore Beaufort County’s waterways. Final report of the Clean Water Task Force. Beaufort County, SC, 70 p.

 

Coen, L.D., and Luckenbach, M.W., 1999, Developing successful criteria and goals for evaluating oyster reef restoration: ecological function or resource exploitation? Ecological Engineering, in press.

 

Coen, L.D., Knott, D.M., Wenner, E.L., Hadley, N.H., and Ringwood, A.H., 1999a, Intertidal oyster reef studies in South Carolina: Design, sampling and experimental focus for evaluating habitat value and function. In: Luckenbach, M.W., Mann, R., Wesson, J.A. (eds) Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. Virginia Institute of Marine Science Press, Gloucester Point, VA.

 

Coen, L.D., Luckenbach, M.W., and Breitburg D.L., 1999b, The role of oyster reefs as essential fish habitat: a review of current knowledge and some new perspectives. Pages 438-454 in Benaka, L.R. (ed) Fish Habitat: Essential Fish Habitat and Rehabilitation. American Fisheries Society, Symposium 22, Bethesda, Maryland.

 

Cohen, J.E., Small, C., Mellinger, A., Gallup, J., and Sachs, J., 1997, Estimates of coastal populations. Science, v. 278, p. 1211-1212.

 

Csallany, A.S., Guan, M.D., Manwaring, J.D., and Addis, P.B., 1984, Free malonaldehyde determination in tissues by high-performance liquid chromotography. Analytical Biochemistry, v. 142, p. 277-283.

 

Culliton, T.J., Warren, M.A., Goodspeed, T.R., Remer, D.G., Blackwell, C.M., and McDonough, J.I., 1990, The second report of a coastal trends series: 50 years of population change along the nation's coasts 1960-2010. U.S. Department of Commerce, National Oceanic and Atmospheric Administration, Rockville, MD, 41 p.    

 

Dodd, M.G., and Murphy, T.M., 1996, The status and distribution of wading birds in South Carolina, 1988-1996. Final Report. SCDNR Technical Report, 66 p.         

 

Driver, N.E., and Troutman, B.M., 1989, Regression models for estimating urban storm-runoff quality and quantity in the United States. Journal of Hydrology, v. 109, p. 221-236.

 

Edwards, S., 1989, Estimates of future demographic changes in the coastal zone. Coastal Management, v. 17, p. 229-240.

 

Fortner, A.R., Sanders, M., and Lemire, S.W., 1996, Polynuclear aromatic hydrocarbon and trace metal burdens in sediment and the oyster, Carssostrea virginica Gmelin, from two-high salinity estuaries in South Carolina. In: Vernberg, F.J., Vernberg, W.B., and Siewicki, T. (eds.) Sustainable development in the southeastern coastal zone. University of South Carolina Press, Columbia, SC.

 

Fulton, M.H., Scott, G.I., Fortner, A., Bidleman, T.F., and Ngabe, B., 1993, The effects of urbanization on small high-salinity estuaries of the southeastern United States. Archives of Environmental Contamination and Toxicology, v. 25, p. 476-484.

 

Gutteridge, J.M.C., and Halliwell, B., 1990, The measurement and mechanism of lipid peroxidation in biological systems. TIBS, v. 15, p. 129-135.

 

Hach Company, 1994, DR/700 Colorimeter Procedures Manual, #46014-88.

 

Howard, D.W., and Smith, C.S., 1983, Histological techniques for marine bivalve mollusks. NOAA Technical Memorandum NMFS-F/NEC 25:97.

 

Kaspar, C.W., Curgess, J.L., Knight, I.T., and Cowell, R.R., 1990, Antibiotic resistance indexing of Escherichia coli to identify sources of fecal contamination in water. Canadian Journal of Microbiology, v. 36, p. 891-894.

 

Krahn, M.M., Moore, L.K., Bogar, R.G., Wigren, C.A., Chan, S-L, and Brown, D.W., 1988, High-performance liquid chromatographic method for isolating organic contaminants from tissue and sediment extracts. Journal of Chromatography, v. 437, p. 161-175.

 

Kucklick, J.R., Sivertsen, S.K., Sanders, M., and Scott, G.I., 1997, Factors influencing polycyclic aromatic hydrocarbon distributions in South Carolina estuarine sediments. Journal of Experimental Marine Biology and Ecology, v. 213, p. 13-29.

 

Laenen, A., 1989, Acoustic velocity meter systems: Techniques of water resources investigations of the United States Geological Survey, Chapter A17, Book 3, 38 p.

 

Lerberg, S.B., Holland, A.F., and Sanger, D.M., 2000, Responses of tidal creek macrobenthic communities to the effects of watershed development. Estuaries, v. 23, p. 838-853.

 

Lipscomb, S.W., 1995, Quality assurance plan for discharge measurements using broad band acoustic Doppler current profilers. US. Geological Survey Open-file report 95-701, 7 p.

 

Long, E.R., MacDonald, D.D., Smith, S.L., and Calder, F.L., 1995, Incidence of adverse biological effects within ranges of chemical concentrations in marine and estuarine sediments. Environmental Management, v. 19, p. 81-97.

 

Long, E.R., Scott, G.I., Kucklick, J., Fulton, M.H., Thompson, B., Carr, R.S., Biedenbach, J., Scott, K.J., Thursby, G.B., Chandler, G.T., Anderson, J.W., and Sloane, G.M., 1998, Magnitude and extent of sediment toxicity on selected estuaries of South Carolina and Georgia. National Oceanic and Atmospheric Administration Technical Memorandum NOS ORCA 128. National Oceanic and Atmospheric Administration, Silver Spring, MD.

 

Lowe, D.M. Moore, M.N., and Evans, B.M., 1992, Contaminant impact on interactions of molecular probes with lysosomes in living hepatocytes from dab Limanda limanda. Marine Ecological Progress Series, v. 91, p. 135-140.

 

Mallin, M.A., Williams, K.E., Esham, E.C., and Lowe, R.P., 2000, Effect of human development on bacteriological water quality in coastal watersheds. Ecological Applications, v. 10, p. 1047-1056.

 

Microbics Corporation, 1992, Microtox® Manual. Vol. 1. 1992 Edition, Carlsbad, CA.

 

Parveen, S., Murphree, R.L., Edmiston, L., Kaspar, C.W., Portier, K.M., and Tamplin., M.L., 1997, Association of multiple antibiotic resistance profiles with point and nonpoint sources of E. coli in Apalachicola Bay. Applied and Environmental Microbiology, v. 63, p. 2607-2612.

 

Plumb, R.H., Jr., 1981, Procedures for handling and chemical analysis of sediment and water samples. Environmental Laboratory, US Army Waterways Experiment Station, Vicksburg, MS.

 

Rantz, S.E, and others, 1992, Measurements and computation of stream flow. US Geological Survey Water-Supply Paper 2175, 631 p.

 

Ray, S.M., 1952, A culture technique for the diagnosis of infection with Dermocystidium marinum, Mackin, Owen, Collier, in oysters. Science, v. 166, p. 360-361.

 

Ringwood, A. H., Conners, D., and Hoguet, J., 1998, The effects of natural and anthropogenic stressors on lysosomal destabilization in oysters, Crassostrea virginica. Marine Ecological Progress Series, v. 166, p. 163-171.

 

Ringwood, A.H., and Keppler, C., 1998, Seed clam growth: A sediment bioassay developed in the EMAP Carolinian Province. Environmental Monitoring and Assessment, v. 51, p. 247-257.

 

Ringwood, A.H., DeLorenzo, M.E., Ross, P.E., and Holland, A.F., 1997, Interpretation of Microtox® solid-phase toxicity tests: the effects of sediment composition. Environmental Toxicology and Chemistry, v. 16, p. 1135-1140.

 

Sanger, D.M., Holland, A.F., and Scott, G.I., 1999a, Tidal creek and salt marsh sediments in South Carolina coastal estuaries: I. Distribution of trace metals. Archives of Environmental Contamination and Toxicology, v. 37, p. 445-457.

 

Sanger, D.M., Holland, A.F., and Scott, G.I., 1999b, Tidal creek and salt marsh sediments in South Carolina coastal estuaries: II. Distribution of organic contaminants. Archives of Environmental Contamination and Toxicology, v. 37, p. 458-471.

 

Simpson, M.R., and Bland, R., 1999, Techniques for accurate estimation of net discharge in a tidal channel. In: Proceedings of the IEEE Sixth Working Conference on Current Measurement, San Diego, CA. March 11-13, 1999. p 125-130.

 

South Carolina Department of Health and Environmental Control, 1997, Environmental Investigations Standard Operating Procedures and Quality Assurance Manual. Office of Environmental Quality Control. Columbia, SC.

 

Strickland, J.D.H., and Parsons, T.R., 1972, A practical handbook of sea water analysis. Fisheries Research Board of Canada. Ottawa, Canada. 309 p.

 

USGS, 1999, Field measurements, Chapter A6, in National field manual for the collection of water-quality data, Wilde, F.D., and Radtke, D. B., eds, U. S. Geological Survey TWRI Book 9, variously paged.

 

Van Dolah, R.F., Chestnut, D.E., and Scott, G.I., 2000, A baseline assessment of environmental and biological conditions in Broad Creek and the Okatee River, Beaufort County, South Carolina. Bureau of Water, South Carolina Department of Health and Environmental Control. Columbia, SC.

 

Van Heukelem, L., Lewitus, A., Kana, T., and Craft, N., 1994, Improved separations of phytoplankton pigments using temperature-controlled high performance liquid chromotography. Marine Ecology Progress Series, v. 114, p. 303-313.

 

Van Heukelem, L., and Thomas, C.S., 2001, Computer-assisted high performance liquid chromotography method development with applications to the isolation and analysis of phytoplankton pigments. Journal of Chromotography A, v. 910, p. 31-49.

 

Wagner, R.J., Mattraw, H.C., Ritz, G.F., and Smith, B.A., 2000, Guideline and Standard Procedures for Continuous Water-Quality Monitors: Site Selection, Field Operation, Calibration, Record Computation, and Reporting. US Geological Survey Water-Resources Investigations Report 00-4252, 53 p.

 

Weinstein, M.P., 1979, Shallow marsh habitats as primary nurseries for fishes and shellfish, Cape Fear River, North Carolina. Fishery Bulletin, v. 77, p. 339-357.

 

Wenner, E.L., and Beatty, H.R., 1993, Utilization of shallow estuarine habitats in South Carolina, U.S.A., by postlarval and juvenile stages of Penaeus spp. (Decapoda: Penaeidae). Journal of Crustacean Biology, v. 13, p. 280-295.

 

 

Table 1.        Methods and detection limits for contaminants in sediments

Chemical

Class

Analyte

CAS Number