6.0 Description of Methods

Contents

6.1 Water Sampling

One of the objectives of the RMP is to evaluate if water quality objectives are met at sampled stations. Therefore, the sampling and analysis methods must be able to detect, and wherever possible quantify, substances below these levels. In order to attain the low detection levels used in the RMP, ultra-clean sampling methods are used in all sampling procedures (Flegal and Stukas, 1987; EPA Method 1669, 1995).

Water samples are collected approximately one meter below the water surface using peristaltic and gear-driven pumps. The sampling ports for both the organic chemistry and trace element samplers are attached to aluminum poles that are oriented up-current from the vessel and upwind from equipment and personnel. The vessel is anchored and the engines turned off. Total (or near-total) and dissolved fractions of Estuary water are measured for trace elements. Particulate and dissolved fractions are measured for trace organics, and totals are calculated.

The RMP used the polyurethane foam plug sampler to collect water for trace organics analyses during the first four years of the Program (Risebrough et al., 1976; de Lappe et al., 1980; 1983) and began to phase in a new, modified, commercially available resin extraction sampler in 1996, beginning with side-by-side comparisons of both sampling systems. XAD (cross-linked amberlite dininyl benzene) resins have been used throughout the world to measure synthetic organic contaminants in both water and air (Infante et al., 1993). The sampler comparisons were continued in 1997, and results from both years are presented in 1997 Annual Report (SFEI, 1999). Beginning with the 1997 monitoring year, the custom-manufactured Axys system (Axys Environmental Systems, Ltd., Sidney, B.C.) has been used to collect all RMP water samples for analysis of trace organic pollutants. It consists of a constant-flow, gear-driven positive displacement pump, ¹/₂ inch Teflon^® tubing, 1 µm glass fiber cartridge particulate filter, and two parallel Teflon^® columns filled with XAD-2 resin with a particle size range of 300-900 µm. Amberlite XAD-2 resin is a macroreticular, styrene-divinyl benzene copolymer, nonionic bead. Each bead is an agglomeration of microspheres. This sponge-like structure offers excellent physical and chemical stability. The discrete pores allow rapid mass transfer of analytes, and the mesh size ensures very little, if any, back pressure during use. The hydrophobic chemical nature of the resin leads to excellent capability of concentrating hydrophobic contaminants.

The sample water is first passed through a coarse screen as it moves into the Teflon^® intake line to remove large particles that may interfere with sample collection; particles greater than 140 Âµm are removed as the sample water passes through the inline pre-filter. The water then passes through the pump head and through a pressure gauge, before it goes through one of two parallel four-inch diameter wound glass fiber filters (1 µm). Using two filters allows a quick change to the second filter if the first filter becomes clogged, without interrupting sample collection. Material retained on the glass fiber filter (or filters) becomes the particulate fraction. After passing through the filter, the water is split and routed through two Teflon^® columns, packed with 85 mL of XAD-2 resin. Two filters are used simultaneously to increase the flow to approximately 1.3 L/min. The compounds which are adsorbed to the XAD resin are classified as the dissolved fraction. Lastly, the water passes through a flow meter and out the exit tube where the extracted water volume is verified with 20 L carbouys.

Equipment blanks are taken for both the resin columns and the glass fiber filters. The two column blanks are collected by leaving both ends of a column open while the filled sample columns are being loaded into the sampler. Similarly, the two glass fiber filter blanks are collected by exposing a filter to the air while loading the sample filters into the cartridges. The blanks receive the same analytical treatment in the laboratory as the field samples.

For trace metals, water samples are collected using a peristaltic pump system equipped with C-Flex tubing in the pump head. Sample aliquoting is conducted on deck on the windward side of the ship to minimize contamination from shipboard sources (Flegal and Stukas, 1987). Filtered water samples are obtained by placing an acid-cleaned polypropylene filter cartridge (Micron Separations, Inc., 0.45 µm pore size) on the outlet of the pumping system. Unfiltered water samples are pumped directly into acid-cleaned containers. Prior to collecting water, several liters of water are pumped through the system, and sample bottles are rinsed five times before filling. The bottles are always handled with polyethylene-gloved "clean hands". The sample tubing and fittings are acid-cleaned polyethylene or Teflon^®, and the inlets and outlets are kept covered except during actual sampling. Samples are acidified within two weeks in a class 100 trace metal laboratory, except for chromium samples, which are acidified and extracted within a hour of collection.

Samples for conventional water quality parameters are collected using the same apparatus as for trace metals; however, containers are only rinsed three times, and the "clean hands" procedure is unnecessary.

Water samples are collected for toxicity tests using the same pumping apparatus as for the collection of the trace organic samples, but are not filtered. Five gallons of water are collected and placed in ice chests for transfer at the end of each cruise day to the testing laboratory. Two field blanks are collected each cruise by filtering (0.45 Âµm) water known to be non-toxic from the Bodega Marine Laboratory.

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6.2 Sediment Sampling

Sediment sampling is conducted using a Young-modified van Veen grab with a surface area of 0.1 m². The grab is made of stainless steel, and the jaws and doors are coated with Dykon^® (formerly known as Kynar^®) to achieve chemical inertness. All scoops, buckets, and stirrers used to collect and homogenize sediments are also constructed of Teflon^® or stainless steel coated with Dykon^®. Sediment sampling equipment is thoroughly cleaned prior to each sampling event. In order to further minimize sample contamination, personnel handling the sample wear gloves.

A sub-core of sediment is removed for measurement of porewater ammonia. Then, the top 5 cm of sediment is scooped from each of two replicate grabs and mixed in a Dykon^®-coated bucket to provide a single composite sample for each station. Between sample grabs, the compositing bucket is covered with aluminum foil to prevent airborne contamination. After two sediment samples have been placed into the compositing bucket, the bucket is taken into the ship’s cabin and thoroughly mixed to obtain a uniform, homogeneous mixture. Aliquots are subsequently split for each analytical laboratory, for archive samples, and for sediment toxicity tests. The quality of grab samples is ensured by requiring each sample to satisfy criteria concerning depth of penetration and disturbance of the sediment within the grab.

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6.3 Benthic Infauna

Benthic infauna samples are comprised of primarily sedentary invertebrate organisms that burrow in or live on the surface of sediments. One sample is taken at each of the nine RMP sediment stations with a Ponar grab sampler. Lead weights are added to or removed from the outside of the grab as appropriate to the sediment type in order to control depth of penetration. Incomplete closure of the grab results in rejection of the sample. The retrieved grab is placed on a stand designed with a stainless steel funnel directed to a sample bucket. Once the grab has passed acceptance criteria (complete closure, no evidence of sediment washout through the doors, even distribution of sediment in the grab, minimum disturbance of the sediment surface, and minimum overall sediment depth appropriate for the sediment type), the grab jaws are opened, and the sediment is dumped into a five-gallon plastic bucket. The sample is then moved to a wash table for sieving through two screens stacked on top of each other. The top screen has a 1 mm mesh size, and the smaller screen retains animals in its 0.5 mm mesh. The material retained in each screen is gently washed into separate, labeled sample jars. A wash bottle with seawater is used to rinse any material on the inside screen frame and canning funnel into the sample jar. Any organisms remaining on the screens are carefully picked off with forceps and placed in the appropriate sample jars. Jars are taken to the formalin station where seawater is decanted from the sample jars with 0.25 mm Nitex mesh. Relcant (isotonic MgCl₂) is added to the sample through the mesh to a level approximately one third higher than the sample level. The sample is allowed to sit in the relaxant for 15 to 30 minutes, the relaxant is decanted, and 10% buffered formalin is added to the sample through the screen lid. As a final step, two to three drops of stain (rose bengal solution) are added to the sample for ease of organism identification.

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6.4 Bivalve Bioaccumulation Sampling

Generally, bivalves are collected from uncontaminated sites and transplanted to fifteen stations in the Estuary during the wet season (February through May) and the dry season (June through September). Contaminant concentrations in the animals’ tissues and the animals’ biological condition (expressed as the ratio of dry weight and shell cavity volume) are measured before deployment (referred to as time zero or background samples) and at the end of the 90-100 day deployment period. Since the RMP sites encompass a range of salinities, three species of bivalves are used, according to the expected salinities in each area and the known tolerances of the organisms. The mussel (Mytilus californianus) is collected from Bodega Head and stored in running seawater at the Bodega Marine Laboratory until deployment at the stations west of Carquinez Strait, which are expected to have the highest salinities. Mytilus californianus will survive short-term exposure to salinities as low as 5 ppt (Bayne, 1976). Oysters (Crassostrea gigas) are obtained from Tomales Bay Oyster Company (Marshall, California) and deployed at moderate-salinity sites closest to Carquinez Strait and in the extreme South Bay. Crassostrea gigas tolerates salinities as low as 2 ppt.

In 1998, the freshwater clam Corbicula fluminea was collected from Lake Chabot in Alameda County and held in the lake four weeks prior to deployment to sites with the lowest salinities. The limited quantity of C. fluminea obtained from Lake Chabot was transplanted to Grizzly Bay during Bivalve Deployment Cruise #16. Consequently, additional specimens were collected for analysis on this cruise from the native population in the Sacramento and the San Joaquin rivers. The native population is scarce and requires a lot of time to collect. Resident clams were also collected from the native populations in the Sacramento and the San Joaquin rivers for dry-season deployment. The transplantation approach with the freshwater clam Corbicula fluminea has been discontinued on an interim basis, since reference population from "clean" locations could no longer be found in sufficient numbers. Corbicula fluminea tolerates salinities from 0 ppt to perhaps 10 ppt (Foe and Knight, 1986). The effects of high, short-term flows of freshwater on the transplanted bivalves west of Carquinez Strait are minimized by deploying the bivalves near the bottom where density gradients tend to maintain higher salinities. All bivalves are kept on ice after collection and deployed within 24-48 hours.

The condition of animals from the control sites at Lake Chabot (Corbicula fluminea), Bodega Head (Mytilus californianus), and Tomales Bay (Crassostrea gigas) was determined at the end of each deployment period in order to sort out Estuary effects from natural factors affecting bivalve condition. Survival during deployment was also measured. Composites of tissue were made from 40-60 individual bivalves from each site before and after deployment for analyses of trace contaminants.

Within each species, animals of approximately the same size are used. Mussels are between 49-81 mm shell length, oysters are between 71-149 mm, and clams are 25-36 mm. One-hundred-fifty oysters and 160 mussels and clams are randomly allocated for deployment at the appropriate sites, with the same number being used as travel blank (time zero) samples for analysis of tissue and condition before deployment. At each site, oysters are divided among five nylon mesh bags, and mussels and clams are divided among four nylon mesh bags.

Moorings are associated with pilings or other permanent structures. Mooring installation, bivalve deployment, maintenance, and retrieval are all accomplished by SCUBA divers.

At each site, a line runs from the bottom of the fixed structure out to the bivalve mooring, which consists of a large screw (earth anchor) that is threaded into the bottom. A large subsurface buoy is attached to the earth anchor by a 1-2 meter-long line. The bivalves (in mesh bags) are attached to the buoy line, which keeps the bivalves off the bottom so they are not smothered. In one hundred and fifty individual deployments, loss of a mooring has occurred on only two occasions, probably due to being ripped out by a vessel anchor.

The deployed samples are checked approximately half-way through the 90-day deployment period to ensure consistent exposure. Moorings and nylon bags are checked for damage and repaired, and fouling organisms are removed.

Upon retrieval, the bags of bivalves are placed into polyethylene bags and taken to the surface. On the vessel, the number of dead organisms are noted. Twenty percent of the live organisms are allocated for condition measurement, and the remainder are equally split for analyses of trace metal and organic compounds. Bivalves used for trace organic analyses are rinsed with reagent grade water to remove extraneous material, shucked using a stainless steel knife (acid-rinsed), and homogenized (until liquefied) in a combusted mason jar using a Tissumizer or Polytron blender. Bivalves used in trace element analyses are shucked with stainless steel knives, gonads are removed, and remaining tissue is rinsed with ultrapure water and placed in acid cleaned, plastic coated, glass jars. The sample is then homogenized (until liquefied) using a Brinkmann homogenizer equipped with a titanium blade.

Based on findings by Stephenson (1992) during the RMP Pilot Program, bivalve guts are not depurated before homogenization for tissue analyses, although gonads are removed from organisms for trace metal analyses. Stephenson (1992) found that, with the exception of lead and selenium, no significant differences exist in trace metal concentrations between mussels depurated for 48 hours in clean Granite Canyon seawater before homogenization and undepurated mussels. However, sediment in bivalve guts may contribute to the total tissue contaminant concentration.

6.5 Analytical Methods

6.5.1 Conventional Water Quality Parameters

Samples for dissolved nutrients are analyzed using the Lachat QuikChem 800 System Nutrient Autoanalyzer (Ranger and Diamond, Lachat Instruments, 1994). The QuickChem methods used are: 31-114-27-1 for silicates, 31-107-06-1 for ammonia, 31-107-04-1 for nitrate/nitrite, and 31-115-01-3 for phosphate. Chlorophyll and phaeophytin are measured using a fluorometric technique with filtered material from 200 mL samples (Parsons et al., 1984). Shipboard measurements for temperature, salinity, pH, and dissolved oxygen content are made using a hand-held Solomat 520 C multi-functional chemistry and water quality monitor. Dissolved organic carbon (DOC) is measured using high-temperature catalytic oxidation with a platinum catalyst (Fitzwater and Martin, 1993). Total suspended solids (TSS) are determined using method 2540D in Standard Methods for the Examination of Water and Wastewater (Greenberg et al., 1992).

A Sea-Bird SBE19 conductivity, temperature, and depth probe (CTD) is used to measure water quality parameters at depths throughout the water column. CTD casts are taken at each site during water and sediment sampling. At each site, the CTD is lowered to approximately one meter below the water surface and allowed to equilibrate to ambient temperature for 3 minutes. The CTD is then lowered to the bottom at approximately 0.15 meters per second, and raised. Only data from the down cast are kept. Data are downloaded onboard the ship, and processed in the laboratory using software supplied by Sea-Bird.

The CTD measures temperature, conductivity, pressure, dissolved oxygen, and backscatter at a sampling rate of two scans per second. These data are edited and averaged into 0.25 m depth bins during processing. Also during processing, salinity (based on conductivity measurements), oxygen, time, and depth (based on pressure) are calculated. Although the CTD data are not detailed in this report, SFEI maintains these data in its database.

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6.5.2 Trace Elements

In water, total and dissolved (0.45 µm filtered) concentrations of mercury, arsenic, selenium, chromium, copper, nickel, lead, silver, and zinc are measured. Mercury, arsenic, and selenium samples are obtained from the same field sample. The mercury sub-samples are photo-oxidated with the addition of bromium chloride, and quantified using a cold-vapor atomic fluorescence technique. Arsenic and selenium are analyzed by hydride-generation atomic absorption with cryogenic trap preconcentration based on a method described in Liang et al. (1994) and Cercelius et al. (1986).

Chromium samples are collected separately. The suspended particulates undergo hydrofluoric acid digestion, and the dissolved chromium is co-precipitated with a ferrous hydroxide scavenger (Cranston and Murray, 1978). Chromium is quantified by graphite furnace atomic absorption spectrometry (GFAAS).

The remaining trace elements in water are measured using the APDC/DDDC organic extraction and preconcentration method (Bruland et al., 1985; Flegal et al., 1991) and then quantified by GFAAS.

Results for cadmium, chromium, copper, nickel, lead, silver, and zinc are reported by the laboratory in weight/weight units (µg/kg). For use in this report, those values are reported as µg/L, without taking account of the difference in density between Estuary water and distilled water. This difference was not taken into account because it is much less than the precision of the data, which was on the order of 10%. In some instances, dissolved metal concentrations are reported as higher than total (dissolved + particulate) metal concentrations. This is due to expected analytical variation in the methods of analysis, particularly at concentrations near the detection limits. Such results should be interpreted as no difference between dissolved and total concentrations, or that the total fraction of metals is in the dissolved phase.

Sediments are digested with aqua regia to obtain "near-total" concentrations of aluminum, silver, cadmium, chromium, copper, iron, manganese, nickel, lead, and zinc (Flegal et al., 1981). The metals are quantified by inductively coupled plasma atomic emission spectrometry (ICP-AES) or by ICP-MS. The method chosen for RMP sediment analysis is comparable to standard EPA procedures (Tetra Tech, 1986), but does not decompose the silicate matrix of the sediment. Because of this, any element tightly bound as a naturally occurring silicate may not be fully recovered.

Bivalve tissue samples are digested with aqua regia to obtain near-total concentrations of trace elements similar to techniques used in the California State Mussel Watch Program (e.g., Flegal et al., 1981; Smith et al., 1986) and consistent with the RMP Pilot Program (Stephenson, 1992). The trace metals are quantified by ICP-AES or ICP-MS. Hydride generation coupled with atomic absorption spectroscopy is used to quantify arsenic. Mercury is quantified using a cold-vapor atomic fluorescence technique, and selenium using the methods of Cutter (1986). Butyltins are measured following NOAA Status and Trends Mussel Watch Project methods described in NOAA Technical Memorandum NOS/ORCA/CMBAD71 vol. IV (NOAA, 1993). This technique involves extracting the sample with hexane and the chelating agent tropolone and measuring the butyltin residues by capillary gas chromatography. Concentrations are expressed in total tin per gram of tissue dry weight.

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6.5.3 Trace Organics

For water samples, each of the two resin columns (each sample is contained in two parallel resin columns) and filters containing the particulate fraction are spiked with extraction surrogates. In 1998, electron capture detector (ECD) surrogates consisted of PCB 207 for the first fraction, and Polychloronitorobenzene (PCNB) for fractions 2 and 3. The mass spectral detector (MSD) surrogate consisted of deuterated acenaphthalene. The XAD columns are eluted in reverse with methanol and methylene chloride in a method similar to the filter cartridges. The separate extracts are then combined and separated into three fractions. Extraction methods are based upon standard EPA and Axys extraction protocols.

The extracts are subjected to Florisil column chromatography resulting in three fractions, a PCB/aliphatic, a pesticide/aromatic fraction, and a polar third fraction, which contains diazinon and other polar pesticides. Chlorinated hydrocarbons (CH) are analyzed on a Hewlett Packard 6890 capillary gas chromatograph utilizing electron capture detectors (GC/ECD). A single 2 µL splitless injection is directed onto two 60 m x 0.25 mm columns of different polarity (DB-17 and DB-5) using a y-splitter to provide two-dimensional confirmation of each analyte. The quantitation internal standards utilized for the CH analysis are dibromo-octafluorobiphenyl (DOB) for fraction 1, and PCB 209 for fractions 2 and 3. Analyte concentrations are corrected for surrogate losses prior to reporting. PAHs are quantified in the F-2 fraction by analysis on a Hewlett-Packard 6890 capillary gas chromatograph equipped with a 5971A mass spectral detector (GC/MS). A 2 µL splitless injection is chromatographed on a DB-5 column and analyzed in a selected ion monitoring (SIM) mode. The quantitation internal standard utilized for the PAH analysis when samples are at 100 µL is hexamethyl benzene (HMB). Dibromo-octafluorobiphenyl is used as an internal standard for diazinon.

Sediment samples are analyzed based on the methods followed by NOAA’s Status and Trends Program. Samples are extracted according to EPA Method 3545 (acclerated solvent extraction) using elevated temperature (100 ºC) and pressure (1500-2000 pso) to achieve analyte recoveries equivalent to those from Soxhlet extraction, using less solvent and taking significantly less time. This extraction procedure is applicable to the extraction of all trace organic compounds of interest to the RMP. Surrogate standards are added prior to extraction to account for methodological analyte losses. ECD surrogates consist of DOBFB (Dibromoctatfluorbiphenyl), PCB 103, and PCB 198. The extract is concentrated and purified using a combined silica/alumina column purification to remove matrix interferences. Internal standard solutions are tetrachloro-m-xylene (TCMX) and dibutyl chlorendate (DBC). Chlorinated hydrocarbons are quantified in sediment extracts via high-resolution capillary gas chromatography using GC/ECD. Dual-column confirmation on 30-m long, 0.25-mm internal diameter fused silica capillary columns with DB-5 and DB-17 bonded phase is conducted.

Tissue samples are homogenized and macerated, and the eluate is dried with sodium sulfate, concentrated, and purified using a combination of EPA Method 3611 alumina column purification and EPA Method 3630 silica gel purification to remove matrix interferences. PAHs and their alkylated homologues in both sediment and tissue extracts are quantified by GC/MS in the SIM (Selected Ion Monitoring) with a temperature-programmable gas chromatograph with a 30-m long, 0.32-mm internal diameter fused silica capillary column with DB-5MS bonded phase. Surrogates for PAHs consist of naphthalene-d8, acenaphthene-d10, phenanthrene-d10, chrysene-d12, and perylene-d12. In 1998, PCBs in tissue are quantified according to EPA Method 1668 (isotope dilution techniques) using high-resolution gas GC/MS. Pesticides in tissue are quantified via high-resolution capillary gas chromatography using GC/ECD. Dual-column confirmation on 30-m long, 0.25-mm internal diameter fused silica capillary columns with DB-5 and DB-17 bonded phase is conducted on tissue samples also.

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6.5.4 Aquatic Bioassays

Water column toxicity is evaluated using a 48-hour bivalve embryo development test and a seven-day growth test using the estuarine mysid Mysidopsis bahia. The bivalve embryo development test is performed according to ASTM standard method E 724-89 (ASTM, 1991). The mysid test is based on EPA test method 1007. Larval Mytilus spp. are used in both wet- and dry-season sampling periods. The mysid growth and survival test consists of an exposure of 7-day old Mysidopsis bahia juveniles to different concentrations of Estuary water in a static system during the period of egg development and is used during both sampling periods. Appropriate salinity adjustments are made for Estuary water from sampling stations with salinities below the test species’ optimal ranges. Reference toxicant tests with copper chloride and potassium dichromate are performed for the bivalve and mysid tests, respectively. These tests are used to determine if the responses of the test organisms are relatively consistent over time.

The salinities of the ambient samples and the control/diluent (Evian spring water) are adjusted to 5 ppt using artificial sea salts (Tropic Marin). The test concentrations are 100%, 50%, and control, each with eight replicates, and with 20 larvae per replicate. Waste, dead larvae, excess food, and 80% of the test water are siphoned from the test chambers daily, and general water chemistry parameters of dissolved oxygen, pH, and salinity are recorded before and after each water change.

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6.5.5 Sediment Quality Characteristics

Sediment size fractions are determined with a grain-size analyzer based on x-ray transmission (Sedigraph 5100). Total organic carbon is analyzed according to the standard method for the Coulometrics CM 150 Analyzer made by UIC, Inc. This method involves measurements of transmitted light through a cell. The amount of transmitted light is related to the amount of carbon dioxide evolved from a combusted sample. Spectrophotometric analyses of sulfides in sediment porewater are performed using a method adapted from Fonselius (1985) with variations from Standard Methods (APHA, 1985).

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6.5.6 Sediment Bioassays

The RMP uses two sediment bioassays: a ten-day acute mortality test using the estuarine amphipod Eohaustorius estuarius exposed to whole sediment using ASTM method E 1367 (ASTM, 1992), and a sediment elutriate test where larval bivalves are exposed to the material dissolved from whole sediment in a water extract using ASTM method E 724-89 (ASTM, 1991). Elutriate solutions are prepared by adding 100 g of sediment to 400 mL of Granite Canyon seawater, shaken for 10 seconds, allowed to settle for 24 hours, and carefully decanted (EPA and COE, 1977; Tetra Tech, 1986). Larval mussels (Mytilus spp.) are used in both sampling periods, with percent normally developed larvae as the measurement endpoint.

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6.5.7 Bivalve Condition and Survival

The condition of bivalves is a measure of their general health following exposure to Estuary water for 90-100 days. Measurements are made on subsamples of specimens before deployment and on the deployed specimens following exposure. Dry weight (without the shell) and the volume of the shell cavity of each bivalve is measured. Bivalve tissue is removed from the specimens and dried at 60 ºC in an oven for 48 hours before weighing. Shell cavity volume is calculated by subtracting shell volume of water displaced by a whole live bivalve less the volume of water displaced by the shell alone. The condition index is calculated by taking the ratio of tissue dry weight and the shell cavity volume.

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6.6 References

APHA. 1985. Standard Methods for the Examination of Water and Wastewater. 16th Edition. American Public Health Association, Washington, DC. 1268 p.

ASTM. 1991. Designation E 724-89: Standard guide for conducting static acute toxicity tests starting with embryos of four species of saltwater bivalve molluscs. Volume 11.04. American Society for Testing and Materials, Philadelphia, PA.

ASTM. 1992. Designation E 1367: Standard guide for conducting 10-day static sediment toxicity tests with marine and estuarine amphipods. Volume 11.04. American Society for Testing and Materials, Philadelphia, PA.

Bayne, B.L. 1976. Marine Mussels: Their Ecology and Physiology. Cambridge University Press, Cambridge. 506p.

Bruland, K.W., K.H. Coale, and L. Mart. 1985. Analysis of seawater for dissolved cadmium, copper, and lead: An intercomparison of voltametric and atomic absorption methods. Marine Chemistry 17:285-300.

Cercelius, E.A., N.S. Bloom, C.E. Cowan, and E.A. Jeane. 1986. Speciation of Selenium and Arsenic in Natural Waters and Sediments, Volume 2: Arsenic Speciation. EPRI.

Cranston, R.E. and J.W. Murray. 1978. The determination of chromium species in natural waters. Analytica Chimica Acta 99:275-282.

Cutter, G.A. 1986. Speciation of selenium and arsenic in natural waters and sediments. Volume 1: Selenium Speciation. EPRI. EA-4641. Research project 2020-1.

de Lappe, B.W., R.W. Risebrough, A.M. Springer, T.T. Schmidt, J.C. Shropshire, E.F. Letterman, and J. Payne. 1980. The sampling and measurement of hydrocarbons in natural waters. In Hydrocarbons and Halogenated Hydrocarbons in the Aquatic Environment, B.K. Afghan and D. Mackay, eds. Plenum Press, NY, pp. 29-68.

de Lappe, B.W., R.W. Risebrough, and W. Walker II. 1983. A large-volume sampling assembly for the determination of synthetic organic and petroleum compounds in the dissolved and particulate phases of seawater. Can. J. Fish Aquat. Sci. 40(2):322-336.

EPA. 1995. Method 1669: Sampling ambient water for trace metals at EPA water quality criteria levels. EPA 821-R-95-034, United States Environmental Protection Agency, Washington, DC.

EPA and COE. 1977. Technical committee on criteria for dredged and fill material, ecological evaluation of proposed discharge of dredge material into ocean waters; implementation manual of Section 103 of Public Law 92-532 (Marine Protection and Sanctuaries Act 1972), July 1977 (2nd printing April 1978). US Environmental Protection Agency; Environmental Effects Laboratory, U.S. Army Corps of Engineers, Waterways Experiment Station, Vicksburg, MI.

Fitzwater, S.E. and J.M. Martin. 1993. Notes on the JGOFS North Atlantic bloom experiment--dissolved organic carbon intercomparison. Marine Chemistry 41:179-185.

Flegal, A.R. and V.J. Stukas. 1987. Accuracy and precision of lead isotopic composition measurements in sea water. Marine Chemistry 22:163-177.

Flegal, A.R., L.S. Cutter, and J.H. Martin. 1981. A study of the chemistry of marine sediments and wastewater sludge. Final Report to California State Water Resources Control Board.

Flegal, A.R., G.E. Smith, G.A. Gill, S. Sanudo-Wilhelmy, and L.C.D. Anderson. 1991. Dissolved trace element cycles in the San Francisco Bay Estuary. Marine Chemistry 36:329-363.

Foe, C. and A. Knight. 1986. A method for evaluating the sublethal impact of stress employing Corbicula fluminea. American Malacological Bulletin, Special Edition No. 2: 133-142.

Fonselius, S.H. 1985. Determination of hydrogen sulfide. In Methods of Seawater Analysis. Grasshoff, K., M. Ehrhardt, and K. Kremling, eds. 2nd Edition, pp. 73-81.

Greenberg, A.E. (Editor), L.S. Clesceri, and A.D. Eaton. 1992. Standard Methods for the Examination of Water and Wastewater, 18th Edition. Prepared and published jointly by American Public Health Assoc., American Wastewater Assoc., and Water and Environmental Federation. APHA, Washington, DC.

Infante, A.P., N.C. Guajardo, J.S. Alonso, M.C.M. Navascues, M.P.O. Melero, M.S.M. Cortabitarte, and J.L.O. Narvion. 1993. Analysis of organic water pollutants isolated by XAD-2 resins and activated carbon in the gallego river, Spain. Water Res. 7:1167-1176.

Liang, L., R. Danilchik, and Z.R. Huang. 1994. Elimination of dependence on experimental conditions in the determination of Se in water, sediment, coal and biological samples by hydride generation. Atomic Spectroscopy, July/August:151-155.

NOAA. 1993. Sampling and analytical methods of the National Status and Trends Program National benthic surveillance and mussel watch projects 1984-1992, volume IV: Comprehensive descriptions of trace organic analytical methods. G.G. Lauenstein and A.Y. Cantillo (eds.) NOAA Technical Memorandum NOS ORCA 71. National Oceanic and Atmospheric Administration, Silver Spring, MD.

Parsons, T.R., T. Maita, and C.M. Lalli. 1984. A manual of chemical and biological methods for seawater analysis. Pergamon Press, NY. 173 p.

Ranger, C. and D. Diamond. 1994. Lachat Instruments.

Risebrough, R.W., B.W. de Lappe, and W. Walker II. 1976. Transfer of higher-molecular weight chlorinated hydrocarbons to the marine environment. In Marine Pollutant Transfer, H.L. Windom and R.A. Duce, eds. D.C. Heath Company, Lexington, Massachusetts and Toronto, pp. 261-321.

SFEI. 1999. 1997 Annual Report: San Francisco Estuary Regional Monitoring Program for Trace Substances. RMP Report 37. San Francisco Estuary Institute, Oakland, CA pp. A67-A80.

Smith, D.R., M.D. Stephenson, and A.R. Flegal. 1986. Trace metals in mussels transplanted to San Francisco Bay. Environ. Toxicol. and Chem. 5:129-138.

Stephenson, M. 1992. A report on bioaccumulation of trace metals and organics in bivalves in San Francisco Bay. Submitted to California Regional Water Quality Control Board, San Francisco Bay Region. California Department of Fish and Game, Moss Landing Marine Labs, Moss Landing, CA 95039.

Tetra Tech. 1986. Recommended protocols for measuring selected environmental variables in Puget Sound. Prepared for the Puget Sound Estuary Program by Tetra Tech, Inc., Bellevue, WA.