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 (see Appendix B),
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 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 Appendix
D. 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 300900
µm. Aberlite XAD-2 resin is a macroreticular, styrene-divinylbenzene
copolymer, nonionic bead. Each bead is an agglomeration of microspheres.
This spong-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
140µm 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.
Field
blanks are taken for both the resin columns and the glass fiber
filters. The two column field 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 field
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 (0.45µm) water known to be non-toxic from the Bodega
Marine Laboratory.
Sediment
Sampling
Sediment
sampling is conducted using a Young-modified van Veen grab with
a surface area of 0.1m2. 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, gloves are worn by personnel
handling the sample.
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.
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.
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 90100 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
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 1997, the freshwater clam Corbicula fluminea
was collected from Putah Creek for the wet-season deployment and
moved to the University of California, Davis (UCD) for depuration
and deployed at sites with the lowest salinities. Corbicula fluminea
tolerates salinities from 0 ppt to perhaps 10 ppt (Foe and Knight,
1986). Clams were collected from the San Joaquin River for dry-season
deployment. 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 2448 hours.
Because
of the unavailability of clams at Lake Isabella, the RMP's traditional
reference site, clams were collected from Putah Creek and the San
Joaquin River and conditioned at a pond fed by Davis well water
and located at the UCD Institute of Ecology. Additionally, the condition
of animals from the control sites at UCD (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 4060 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 4981 mm shell length, oysters are between 71149
mm, and clams are 2536 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. 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.
For
a more detailed description of field methods, see RMP News, Volume
4, Issue 2 (Gold and Bell, 1998).
Analytical
Methods
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 sediments (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.
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.
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 1997, electron
capture detector (ECD) surrogates consisted of PCB 103 and PCB 207
for the first fraction, and pentachloronitorobenzene for fractions
2 and 3. The mass spectral detector (MSD) surrogate consisted of
deutereated 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 fractions
1 and 3, and DOB or PCB 209 for Fraction 2. 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 0C) and pressure (15002000 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 compounds of interest
to the RMP. Surrogate standards are added prior to extraction to
account for methodological analyte losses. ECD surrogates consist
of DOB, 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 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 consisted of naphthalene-d8, acenaphthene-d10, phenanthrene-d10,
chrysene-d12, and perylene-d12. In 1997, PCBs in tissue were quantified
according to EPA Method 1668 (isotope dilution techniques) using
high-resolution gas GC/MS. Pesticides in tissue were 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 was conducted
on tissue samples also.
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
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.
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).
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.
Bivalve
Condition and Survival References
The
condition of bivalves is a measure of their general health following
exposure to Estuary water for 90100 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 60oC
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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