SIMoN
  Sanctuary Integrated Monitoring Network
Monitoring Project

Analysis of mussels collected near the Moss Landing Power Plant thermal outfall

Principal Investigator(s)

  • Gary Ichikawa
    Moss Landing Marine Laboratories, California State University
  • John Negrey
    Moss Landing Marine Laboratories, California State University
  • Steve Choy
    Monterey Bay National Marine Sanctuary

Funding

  • SIMoN
  • MBSF
Start Date: June 01, 2006
End Date: December 31, 2007

The Mussel Watch Program was established in 1986 by NOAA with the goal of monitoring contaminants in U.S. Coastal waters. The spatial and temporal scale, which has covered the West Coast, East Coast, Gulf of Mexico and Great Lakes annually for the last two decades, makes it one of the most ambitious coastal monitoring efforts in the country. In order to monitor the nearly 140 chemical contaminants targeted by the Mussel Watch program, mussel and oyster specimens are collected every year from approximately 300 sites around the U.S. Mussels and oysters were chosen as indicators of contaminants because they are sessile filter feeders and accumulate contaminants in their tissues, making them good proxies for measuring heavy metals and chemicals. As a part of the local Mussel Watch Program, sampling locations include Point Santa Cruz, Elkhorn Slough, Moss Landing Harbor, and Lovers Point (Figure 1).

In 2006 only, additional sites were sampled at 1) the Highway 1 bridge pilings crossing Elkhorn Slough, 2) Moss Landing Harbor breakwaters, and 3) the outfall pipes at Moss Landing Power Plant (Figure 1). These sampling locations were added to compare measurements from areas influenced by the Moss Landing Power Plant thermal outfall plume to those beyond its influence. The contaminant loads from mussels at the 2006 sites were compared to each other and to those from the long-term sampling sites around Monterey Bay to investigate any differences that may result from the thermal plume.

There has been ongoing concern by some that the thermal plume generated by the power plant is altering water quality conditions in the vicinity of the outfall (beyond those due to elevated temperature). The outfall plume is the result of cooling operations at the power plant. Harbor water is taken in by the plant, used to cool equipment, then discharged back in to the ocean. Water in the harbor is a mix of 1) oceanic water brought in with incoming tides, 2) brackish water from Elkhorn Slough, and 3) freshwater (with agricultural runoff) from Moro Cojo Slough and the old Salinas River channel.

Summary to Date

Mussels collected from bridge pilings and rip-rap from both the breakwater and outfall were in the genus Mytilus. Most were likely the California mussel Mytilus californianus (a native species), but there are also foolish mussels Mytilus trossulus (also a native, but more common north of CA), and the invasive Mediterranean mussel Mytilus galloprovincialis (more common in southern CA).

In 2007 the results of the sampling of thermal outfall sites were reported. The initial results show that the thermal outfall sampling sites were similar to the 20 year averages of the long-term Mussel Watch Program sites for Aluminum (Al), Arsenic (As), Chromium (Cr), Copper (Cu), Lead (Pb), Manganese (Mn), Nickel (Ni), Selenium (Se), Silver (Ag), and Zinc (Zn). Only concentrations of Cadmium (Cd) were found to be higher at the thermal outfall sites compared to the 20 year averages of long-term sites outside of the influence of the thermal plume.

A report highlighting the findings of the Mussel Watch Program from 1986 through 2005 was released in 2008.

Monitoring Trends

  • The Mussel Watch Program reported no significant trends for Arsenic, Cadmium, Copper, Lead, Nickel, or Zinc for the Northern California Region. Since only one year of sampling for the thermal outfall sites has been conducted, no temporal trends could be reported.
  • Mussels collected at the thermal outfall diffusers themselves exhibited higher concentrations of Cd when compared with other sites around Monterey Bay, while concentrations of all other heavy metals tested were similar to other sampling sites. To date, test of statistical significance has not been calculated for the results.

Discussion

The purpose of this analysis was to compare contaminants (mostly heavy metals) in mussels collected from three areas with differing levels of influence from the power plant thermal outfall. Mussels, although uncommon, were sampled from the outfall structure, then the breakwater (moderate distance, about 500 m) rip-rap, and finally from the bridge pilings (about 1 km). As one might expect, given the circulation patterns and mixing of waters in in the Moss Landing Harbor area, there were no differences among the three sites.

When comparing the three sites near the thermal outfall to other sites in Monterey Bay, only cadmium (Cd) levels were higher. This analysis is just a snapshot, comparing spring sampling in 2006 to data collected over a 20+ year period. Ideally these measurements would be repeated periodically, however at a cost of over $10,000, such a study will require external funding to analyze future mussel samples.

Study Parameters

  • Manganese (Mn)
  • Lead (Pb)
  • Aluminum (Al)
  • Chromium (Cr)
  • Copper (Cu)
  • Cadmium (Cd)
  • Nickel (Ni)
  • Arsenic (As)
  • Selenium (Se)
  • Silver (Ag)
  • Zinc (Zn)

Study Methods

Total Mercury 70:30 Nitric/Sulfuric Digest of Tissues
FGS-011.2

Frontier Geosciences Inc.
414 Pontius Avenue North
Seattle, WA 98109

Originated by: Nicolas S Bloom and Efronsini Tsalkitizis
Revised by: Paul Laskowski
January 3, 2000


1.0 SCOPE AND APPLICATION

1.1. By taking of smaller aliquot sizes of the digestate (L range), contaminated solids up to 10,000 ng/g can be directly measured. All samples must be subjected to an appropriate dissolution or leaching step prior to analysis. In general, using clean handling and reagents, the typical detection limit for the method is less than 0.001 g /g as Hg (ie. 0.001 ppm).

1.2. Total mercury as defined by this method means all HNO3/H2SO4+BrCl-oxidizable mercury forms and species found in solid matrix. This includes but is not limited to Hg(II), Hgo, HgS, strongly organocomplexed Hg(II) compounds, adsorbed particulate Hg, and several tested covalently bound organomercurials (i.e., CH3HgCl, (CH3)2Hg, and C6H5HgOOCCH3).

2.0 SUMMARY OF METHOD

2.1 Samples are collected using clean sample handling protocols (Bloom, 1995) into acid-cleaned Teflon vials, plastic or borosilicate glass. A subsample of the homogenized sample is digested with 10 mL 70:30 HNO3/H2SO4. The digested sample is diluted up to 40 mL with 5% BrCl.

3.0 INTERFERENCES

3.1. Due to the high levels of halogens (i.e. iodine) in the digests, it is recommended that aliquots of no more then 1.0 mL of the digestates are analyzed. Otherwise soda-lime traps may be overloaded and the sample analysis gold traps may lose the ability to hold mercury.

3.2. Due to high acidity and halogens in the digests, not changing bubbler water after every 10 mL of digestate analyzed can lead to lower recoveries that would be reflected in the analysis QC.

4.0 SAFETY

4.1. Personnel will don appropriate laboratory attire according to the Chemical Hygeine Plan. This includes, but is not limited to, laboratory coat, safety goggles, and latex gloves under clean gloves.

4.2. The toxicity or carcinogenicity of reagents used in this method has not been fully established. Each chemical should be regarded as a potential health hazard and exposure to these compounds should be a low as reasonably achievable. Chemists should refer to the MSDS for each chemical they are working with.

4.3. All personnel handling environmental samples known to contain or to have been in contact with human waste should be immunized against known disease-causative agents. Frontier will reimburse the expense of Hepatitis A and B immunizations for any laboratory staff member who desires this protection.

5.0 EQUIPMENT

5.1. 40mL or 20mL I-Chem Vials: Borosilicate glass, series 300 vials with Teflon lined septum in lid. The size depends on which size of IC traps are being digested. The vials are volumetrically accurate to 0.5mL when filled such that the meniscus is just to the bottom of the vial neck. The person performing the preparation should verify this.

6.0 REAGENTS

6.1. Water: 18 megohm ultra-pure deionized water starting from a pre purified (distilled, R.O., etc.) source. As a final mercury and organic removal step, the activated carbon cartridge on the 18 megohm system is placed between the final ion exchange bed and the 0.2 M filter. Water should be routinely monitored for Hg--especially after ion exchange beds are changed. Water should typically contain less than 0.2 ng/L total Hg, and remedial action should be taken if the total Hg exceeds 1 ng/L.

6.2. 70:30 Nitric/sulfuric acid: Carefully add 300 mL of pre-analyzed low mercury (<10 ng/L Hg) concentrated sulfuric acid to 750 mL pre-analyzed, low mercury concentrated nitric acid, in a Teflon bottle, with constant stirring. CAUTION: This mixture becomes exothermic and emits caustic fumes.

6.3. 0.2N Bromine Monochloride: 37.5 g of KBr are added to a 2.5 L bottle of concentrated HCl analyzed and found to be low in Hg (<5 ng/L). A clean magnetic stir bar is placed in the bottle, and stirred for 1 hour, in a fume hood. Next, 27.5 g Hg KBrO3 is slowly added to the acid while stirring. When all of the KBrO3 has been added, the solution should have gone from yellow to red to orange. Loosely cap the bottle, and allow to stir another hour before tightening the lid. CAUTION: This process generates copious quantities of free halogens (Cl2, Br2, BrCl) which are released from the bottle. Add the KBrO3 SLOWLY and in a well operating fume hood!

7.0 PROCEDURE

7.1. Approximately 0.5-1.0 gram of the homogenized sample is accurately weighed into a 40 mL I-Chem vial.

7.2. 10.0 mL of 70:30 v/v HNO3/H2SO4 solution is pipetted in, and the sample is swirled.

7.3. The vial is placed on a hot plate with a Teflon reflux cap in the place of the vials lid.

7.4. The Samples are heated to 125 C for 2 hours after the onset of refluxing or until all organic matter is dissolved.

7.5. The samples are allowed to cool and are diluted to 40 mL with a 5% (v/v) solution of 0.2N BrCl in milli-Q water, capped with their respective lids, and are thoroughly shaken before analysis.

8.0 QUALITY CONTROL

8.1. Maximum Sample Batch Size: 25 samples for standard level QC; 20 samples for high level QC.

8.2. Preparation Blanks: 3 per batch, standard deviation used to determine estimated MDL for each batch.

8.3. SRM: 1 per batch, 75-125% recovery limit.

8.4. MD: 1 per batch, 25 RPD limit.

8.5. MS/MSD: 1 per batch, 75-125% recovery limit, 25 RPD limit.

9.0 CORRECTIVE ACTIONS

9.1. Corrective action to follow if quality assurance measure is out.

10.0 REFERENCES

10.1. Bloom, N.S. (1995) "Ultra-Clean Sample Handling." Env. Lab March/April: 20.

10.2. Bloom N.S (1989) "Determination of Picograms Levels of Methylmercury by aqueous phase ethylation, followed by Cryogenic Gas Chromatography with Cold Vapor Atomic Fluorescence Detection". Can J. Fish. Aquat. Sc. 7: 1131.

10.3. Bloom, N.S and Crecelius, E.A. (1983) "Determination of Mercury in Seawater at Subnanogram per Litre Levels". Mar. Chem. 14: 49.

10.4. Bloom, N.S. and Crecelius, E.A. (1987) "Distribution of Silver, Lead, Mercury, Copper, and Cadmium in Central Puget Sound Sediments," Mar. Chem. 21: 377.

10.5. Bloom, N.S and Fitzgerald, W.F. (1988) "Determination of Volatile Mercury Species at the Picogram Level by Low-Temperature Gas Chromatography with Cold-Vapor Atomic Fluorescence Detection". Anal. Chim. Acta. 208: 151.

10.6. Cossa, D. and Couran, P. (1990) "An International Intercomparison Exercise for Total Mercury in Seawater." App. Organomet. Chem. 4: 49.

10.7. Fitzgerald, W.F. and Gill, G.A. (1979) "Sub-Nanogram Determination of Mercury by Two-Stage Gold Amalgamation and Gas Phase Detection Applied to Atmospheric Analysis". Anal. Chem. 15: 1714.



Method # DFG 102a

SAMPLING MARINE AND FRESHWATER BIVALVES, FISH AND CRABS FOR TRACE METAL AND SYNTHETIC ORGANIC ANALYSIS

1.0 Scope and Application

1.1 The following procedures describe techniques of sampling marine mussels and crabs, freshwater clams, marine and freshwater fish for trace metal (TM) and synthetic organic (SO) analyses.

2.0 Summary of Method

2.1 Collect mussels, clams, crabs, or fish. Mussels or clams to be transplanted are placed in polypropylene mesh bags and deployed. Mussels and clams to be analyzed for metals are double-bagged in plastic zipper-closure bags. Bivalves to be analyzed for organics are wrapped in PE cleaned aluminum foil prior to placement in the zipper-closure bags. Fish are wrapped whole or proportioned where necessary in cleaned Teflon sheets or aluminum foil and subsequently placed into zipper-closure bags. Crabs for TM and/or SO are double-bagged in plastic zipper-closure bags.

2.2 Each sample should be labeled with Date, Station Name, and any other information available to help identify the sample once in the lab.

2.3 After collection, samples are transported back to the laboratory in coolers with ice or dry ice. If ice is used, care must be taken to ensure that ice melt does not come into direct contact with samples.

3.0 Interferences

3.1 In the field, sources of contamination include sampling gear, grease from ship winches or cables, ship and truck engine exhaust, dust, and ice used for cooling. Efforts should be made to minimize handling and to avoid sources of contamination.

3.2 Solvents, reagents, glassware, and other sample processing hardware may yield artifacts and/or elevated baselines, causing inaccurate analytical results. All materials should be demonstrated to be free from interferences under the conditions of the analysis by running method blanks initially and with each sample lot.

3.3 Polypropylene and polyethylene surfaces are a potential source of contamination for SO specimens and should not be used whenever possible.

4.0 Apparatus and Materials

Procedures for equipment preparation can be found in Method # {_______}

4.1 Anchor Chains

4.2 Backpack Shocker (electro-fishing)

4.3 Boats (electro-fishing and/or for setting nets)

4.4 Bone Saw

4.5 Camera, digital

4.6 Cast Nets (10 and 12)

4.7 Data Sheets (see Appendix .)

4.8 Daypacks

4.9 Depth Finder

4.10 Dip Nets

4.11 Dry Ice or Ice

4.12 Gill Nets (various sizes)

4.13 GPS

4.14 Heavy Duty Aluminum Foil, prepared

4.15 Heavy Duty plastic bags, Clear 30 gallon

4.16 Inflatable Buoy

4.17 Labels, gummed waterproof: Diversified Biotech Part #: LCRY-1258

4.18 Nylon Cable Ties, 7/16 wide x 7 long

4.19 Other (minnow traps, set lines, throw nets, etc)

4.20 Otter Trawl (various widths as appropriate)

4.21 Permanent Marking Pen

4.22 Plastic bucket, 30 gallon

4.23 Plastic Ice Chests

4.24 Polyethylene Gloves: VWR Part # 32915-166, 32915-188, and 32915-202

4.25 Polypropylene Mesh, 76mm wide with 13mm mesh

4.26 Polypropylene Mesh, 50mm wide with 7mm mesh

4.27 Polypropylene Line, 16mm

4.28 Rods and Reels

4.29 Screw in Earth Anchor, 4-6 diameter

4.30 Scuba Gear

4.31 Seines (various size mesh and lengths as appropriate)

4.32 Stainless Steel Dive Knives

4.33 Trap Nets (hoop or fyke nets)

4.34 Teflon Forceps

4.35 Teflon Sheet, prepared

4.36 Teflon Wash Bottle, 500 mL

4.37 Wading Gear

4.38 Zipper-closure Polyethylene Bags, 4milx13x18: Packaging Store Part # zl401318redline

5.0 Reagents

5.1 Tap water (Tap)

5.2 Deionized water (DI)

5.3 Type II water (ASTM D1193): Use Type II water, also known as MilliQ, for the preparation of all reagents and as dilution water.

5.4 Micro Detergent: ColeParmer Part # 18100-20

5.5 Methanol: VWR Part # JT9263-3

5.6 Petroleum Ether: VWR Part # JT9265-3

6.0 Sample Collection, Preservation and Handling

6.1 All sampling equipment will be made of non-contaminating materials and will be inspected prior to entering the field. Nets will be inspected for holes and repaired prior to being used. Boats (including the electroshocking boat) will be visually checked for safety equipment and damage prior to being taken into the field for sample collection.

6.2 To avoid cross-contamination, all equipment used in sample collection should be thoroughly cleaned before each sample is processed. Ideally, instruments are made of a material that can be easily cleaned (e.g. Stainless steel, anodized aluminum, or borosilicate glass). Before the next sample is processed, instruments should be washed with a detergent solution, rinsed with ambient water, rinsed with a high-purity solvent (methanol or petroleum ether), and finally rinsed with MilliQ. Waste detergent and solvent solutions must be collected and taken back to the laboratory

6.3 Samples are handled with polyethylene-gloved hands only. The samples should be sealed in appropriate containers immediately.

6.4 Mussels and clams to be analyzed for metals are double-bagged in zipper-closure bags. Bivalves to be analyzed for organics are wrapped in prepared aluminum foil prior to placement in zipper-closure bags.

6.5 Fish are wrapped in part or whole in prepared Teflon sheets and subsequently placed into zipper-closure bags.

6.6 Crabs analyzed for metals and/or organics are double-bagged in plastic zipper-closure bags.

6.7 Data is recorded for each site samples are transplanted to or collected from. Data includes, but is not limited to station name, sample identification number, site location (GPS), date collected or transplanted, collectors names, water depth, photo number, ocean/atmospheric conditions (if appropriate), description of site, and drawing if necessary.

6.8 A chain of custody form (Appendix A-1) will accompany all samples that are brought to the lab. All samples that are processed in the lab MUST be checked in according to Method {_____}.

6.9 Tissue and sediment samples are maintained at -20c and extracted or digested as soon as possible. Complete analyses are performed within 40 days of digestion or extraction.

7.0 Procedure

7.1 Sample collection - mussels and clams

7.1.1 The mussels to be transplanted (Mytilus californianus) are collected from Trinidad Head (Humboldt Bay Intensive Survey), Montana de Oro (Diablo Canyon Intensive Survey), and Bodega Head (all other statewide transplants). The freshwater clam (Corbicula fluminea) source is Lake Isabella or the Sacramento River. Analyze mussel and clam samples for background contaminates prior to transplanting.

7.1.2 Polyethylene gloves are worn while prying mussels off rocks with dive knives. Note: polyethylene gloves should always be worn when handling samples. Mussels of 55mm to 65mm in length are recommended. Fifty mussels are collected for each TM and each SO sample.

7.1.3 Collected mussels are carried out of collection site in zipper-closure bags placed in cleaned nylon daypacks. For the collection of resident samples where only one or two samples are being collected the mussels are double bagged directly into a labeled zipper-closure bag. Samples for SO are wrapped first in prepared aluminum foil.

7.1.4 Clams (Corbicula fluminea) measuring 20 to 30mm are collected by dragging the clam dredge along the bottom of the lake or river. The clams are poured out of the dredge into a 30 gallon plastic bag. Clams can also be collected by gloved hands in shallow waters and placed in labeled zipper-closure bags. 25-200 clams are collected depending on availability and necessity for analyses.

7.1.5 Data is recorded for each site samples are collected from. Data includes, but is not limited to station name, date collected, collectors names, water depth, GPS readings, photo, ocean/atmospheric conditions (if appropriate), description of site, and drawing if necessary.

7.2 Transplanted sample deployment

7.2.1 With polyethylene gloves, fifty transplant mussels are placed in each 76mm X 13mm polypropylene mesh bag. Each bag represents one TM or one SO sample. A knot is tied at each end of mesh bag and reinforced with a cable tie. On one end another cable tie is placed under the cable tie which will be used to secure the bag to the line for transplant deployment. The mussels in the mesh bag are divided into three groups of approximately equal size and sectioned with two more cable ties.

7.2.2 Once bagged, the mussels are placed in a 30 gallon plastic bag and stored in a cooler (cooled with ice) for no more than 48 hours. The ice is placed in zipper-closure bags to avoid contamination.

7.2.3 If marine samples are held for longer than 48 hours they are placed in holding tanks with running seawater at the lab. Control samples for both SO and TM are also held in the tank.

7.2.4 For freshwater clams: clams (25-200) are placed in 50mm X 7mm polypropylene mesh bags using identical procedures to those used with mussels (section 7.2.1). If clams need to be stored for more than 48 hours, the mesh bags are deployed either in a clean source or in holding tanks with running freshwater at the lab until actual sample deployment.

7.2.5 The mussels are attached to an open water transplant system that consists of a buoy system constructed with a heavy weight anchor (about 100lbs) or screw-in earth anchor, 13mm polypropylene line, and a 30cm diameter subsurface buoy. The sample bags are attached with cable ties to the buoy line about 15 feet below the water surface. In some cases the sample is hung on suspended polypropylene lines about 15 feet below the water surface between pier pilings or other surface structures. Creosote-coated wooden piers are avoided because they are a potential source of contamination. In some cases the mussels are hung below a floating dock. In shallow waters a wooden or PVC stake is hammered into the substrate and the mussel bags are attached by cable ties to the stake.

7.2.6 The clams are deployed by attaching the mesh bag with cable ties to wooden or PVC stakes hammered into substrate or screw in earth anchors. The bags containing clams are typically deployed 15cm or more off the bottom. In areas of swift water, polypropylene line is also attached to the staked bags and a permanent object (piling, tree or rock).

7.2.7 Transplants are usually deployed for 1-4 months. Ideally mussels are transplanted in early September and retrieved in late December and early January. Clams are usually transplanted in March or April and retrieved in May or June.

7.2.8 Data is recorded for each site samples are transplanted to or collected from. Data includes, but is not limited to station name, date collected or transplanted, collectors names, water depth, GPS readings, photo, ocean/atmospheric conditions (if appropriate), description of site, and drawing if necessary.

7.3 Sample Retrieval

7.3.1 The transplanted or resident and control mussels analyzed for TM are double bagged in appropriately sized and labeled zipper-closure bags.

7.3.2 All mussels to be analyzed for SO are wrapped in prepared aluminum foil ({Method # }). The foil packet is double bagged in appropriately sized and labeled zipper-closure bags. Note: samples should only contact the dull side of the foil.

7.3.3 The bags containing samples are clearly and uniquely identified using a water-proof marking pen or pre-made label. Information items include ID number, station name, depth (if from a multiple sample buoy), program identification, date of collection, species and type of analysis to be performed.

7.3.4 The samples are placed in non-metallic ice chests and frozen using dry ice or regular ice. (Dry ice is used when the collecting trip takes more than two days.) At the lab, samples should be stored at or below -20c until processed.

7.4 Sample Collection Fish

7.4.1 Fish are collected using the appropriate gear for the desired species and existing water conditions.

7.4.1.1 Electro-fisher boat- The electro-fisher boat is run by a trained operator, making sure that all on board follow appropriate safety rules. Once on site, adjustment of the voltage, amps, and pulse for the ambient water is made and recorded. The stainless steel fish well is rinsed with ambient water, drained and refilled. The shocked target fish are placed with a nylon net in the well with circulating ambient water. The nylon net is washed with a detergent and rinsed with ambient water prior to use. Electro-fishing will continue until the appropriate number and size of fish are collected.

7.4.1.2 Backpack electro-fisher- The backpack shocker is operated by a trained person, making sure that all others helping follow appropriate safety rules. The backpack shocker is used in freshwater areas where an electro-fisher boat can not access. Once on site, adjustment of the voltage, amps, and pulse for the ambient water is made and recorded. The shocked target fish are captured with a nylon net and placed in a 30 gallon plastic bag. The nylon net is washed with a detergent and rinsed with ambient water prior to use. Electro-fishing will continue until the appropriate number and size of fish are collected.

7.4.1.3 Fyke or hoop net- Six-36 inch diameter hoops connected with 1 inch square mesh net is used to collect fish, primarily catfish. The net is placed parallel to shore with the open hoop end facing downstream. The net is placed in areas of slow moving water. A partially opened can of cat food is placed in the upstream end of the net. Between 2-6 nets are placed at a site overnight. Upon retrieval a grappling hook is used to pull up the downstream anchor. The hoops and net are pulled together and placed on a 30 gallon plastic bag in the boat. With polyethylene gloves the desired fish are placed in a 30 gallon plastic bag and kept in an ice chest with ice until the appropriate number and size of fish are collected.

7.4.1.4 Otter-trawl- A 14 foot otter trawl with 24 inch wooden doors or a 20 foot otter trawl with 30 inch doors and 80 feet of line is towed behind a boat for water depths less than 25 feet. For water depths greater than 25 feet another 80 feet of line is added to capture fish on or near the substrate. Fifteen minute tows at 2-3 knots speed are made. The beginning and ending times are noted on data sheets. The trawl is pulled over the side of the boat to avoid engine exhaust. The captured fish are emptied into a 30 gallon plastic bag for sorting. Desired fish are placed with polyethylene gloves into another 30 gallon plastic bag and kept in an ice chest with ice.

7.4.1.5 Gill nets- A 100 yard monofilament gill net of the appropriate mesh size for the desired fish is set out over the bow of the boat parallel to shore. The net is retrieved after being set for 1-4 hours. The boat engine is turned off and the net is pulled over the side or bow of the boat. The net is retrieved starting from the down-current end. If the current is too strong to pull in by hand, then the boat is slowly motored forward and the net is pulled over the bow. Before the net is brought into the boat, the fish are picked out of the net and placed in a 30 gallon plastic bag and kept in an ice chest with ice.

7.4.1.6 Beach seines- In areas of shallow water, beach seines of the appropriate length, height, and mesh size are used. One sampler in a wetsuit or waders pulls the beach seine out from shore. The weighted side of the seine must drag on the bottom while the float side is on the surface. The offshore sampler pulls the seine out as far as necessary and then pulls the seine parallel to shore and then back to shore, forming a half circle. Another sampler is holding the other end on shore while this is occurring. When the offshore sampler reaches shore the two samplers come together with the seine. The seine is pulled onto shore making sure the weighted side drags the bottom. When the seine is completely pulled onshore, the target fish are collected with polyethylene gloves and placed in a 30 gallon plastic bag and kept in an ice chest with ice. The beach seine is rinsed off in the ambient water and placed in the rinsed 30 gallon plastic bucket.

7.4.1.7 Cast net- A 10 or 12 foot cast net is used to collect fish off a pier, boat, or shallow water. The cast net is rinsed in ambient water prior to use and stored in a covered plastic bucket. The target fish are sampled with polyethylene gloves and placed in a 30 gallon plastic bag and kept in an ice chest with ice.

7.4.1.8 Hook and line- Fish are caught off a pier, boat, or shore by hook and line. Hooked fish are taken off with polyethylene gloves and placed in a Ziploc bag or a 30 gallon plastic bag and kept in an ice chest with ice.

7.4.1.9 Spear fishing- Certain species of fish are captured more easily by SCUBA divers spearing the fish. Only appropriately trained divers following the dive safety program guidelines are used for this method of collection. Generally, fish in the kelp beds are more easily captured by spearing. The fish are shot in the head area to prevent the fillets from being damaged or contaminated. Spear tips are washed with a detergent and rinsed with ambient water prior to use.

7.4.2 As a general rule, five fish of medium size or three fish of larger size are collected as composites for analysis. The smallest fish length cannot be any smaller than 75% of the largest fish length. Five fish usually provides sufficient quantities of tissue for the dissection of 150 grams of fish flesh for organic and inorganic analysis. The medium size is more desirable to enable similar samples to be collected in succeeding collections.

7.4.3 When only small fish are available, sufficient numbers are collected to provide 150 grams of fish flesh for analysis. If the fish are too small to excise flesh, the whole fish, minus the head, tail, and guts are analyzed as composites.

7.4.4 Species of fish collected are chosen for their importance as indicator species, availability or the type of analysis desired. For example, livers are generally analyzed for heavy metals. Fish without well-defined livers, such as carp or goldfish, are not collected when heavy metal analyses are desired.

7.4.5 Fish collected, too large to fit in clean bags (>500 mm) are initially dissected in the field. At the dock, the fish are laid out on a clean plastic bag and a large cross section from behind the pectoral fins to the gut is cut with a cleaned bone saw or meat cleaver. The bone saw is cleaned (micro, DI, methanol) between fish and a new plastic bag is used. The internal organs are not cut into, to prevent contamination. For bat rays, a section of the wing is cut and saved. These sections are wrapped in prepared Teflon sheets, double bagged and packed in dry ice before transfer to the freezer. During lab dissection, a subsection of the cross section is removed, discarding any tissue exposed by field dissection.

7.4.6 Field data (Appendix A-2) recorded include, but are not limited to site name, sample identification number, site location (GPS), date of collection, time of collection, names of collectors, method of collection, type of sample, water depth, water and atmospheric conditions, fish total lengths (fork lengths where appropriate), photo number and a note of other fish caught.

7.4.7 The fish are then wrapped in aluminum foil or Teflon sheets if thylates are analyzed. The wrapped fish are then double-bagged in zipper-closure bags with the inner bag labeled. The fish are put on dry ice and transported to the laboratory where they are kept frozen until they are processed for chemical analysis.

7.5 Sample Collection- Crabs

7.5.1 Crab/lobster traps- Polyethylene traps are baited to collect crabs or lobsters. Traps are left for 1-2 hours. The crabs are placed in a zipper-closure bag or a 30 gallon plastic bag and kept in an ice chest with ice.

8.0 Analytical Procedure

8.1 Tissue Preparation procedures can be found in Method #______.

8.2 Trace Metal and Mercury Only digestion procedures can be found in Method #______ and Method #______, respectively.

8.3 Trace Metals are analyzed with ICP-MS according to EPA 200.7

8.4 Mercury samples are analyzed by FIMS according to Method #______ or by DMA and Method #______.

9.0 Quality Control

9.1 Field Replicates: One field replicate is taken for every twenty stations and is analyzed for both TM and SO.

9.2 A record of sample transport, receipt and storage is maintained and available for easy reference.

10.0 References

10.1 Flegal, R.A. 1982. In: Wastes in the Ocean, Vol VI: Near Shore Waste Disposal. B.H. Ketchum (ed.). John Wiley and Sons Inc. Publishers, New York, 1982.

10.2 Goldberg, E.D., ed. 1980. The International Mussel Watch. National Academy of Sciences Publ., Washington, D.C.

10.3 Gordon, R.M., G.A. Knauer and J.H. Martin. 1980a. Mytilus californianus as a bioindicator of trace metal pollution: variability and statistical considerations. Mar. Poll. Bull. 9:195-198.

10.4 Hayes, S. P. and P. T. Phillips. 1986. California State Mussel Watch: Marine water quality monitoring program 1984-85. State Water Resources Control Board Water Quality Monitoring Report No. 86-3WQ.

10.5 EPA. 1995. Guidance for Assessing Chemical Contaminant Data for Use in Fish Advisories. Volume 1: Fish Sampling and Analysis. EPA 823-R-95-007.



ANALYSIS OF
EXTRACTABLE SYNTHETIC ORGANIC COMPOUNDS IN TISSUE

1.0 Scope and Application

1.1 This method describes the sample preparation using an automated extraction system for the determination of trace residue levels of a selected list of organochlorine pesticides and poly-chlorinated biphenyls (PCBs) in fish and shellfish tissues by high resolution gas chromatography using electron capture detection. Table 1 lists the target pesticide compounds currently analyzed with their method detection limits and reporting limits for tissues. Table 2 lists the PCB congeners and Aroclor mixtures analyzed with their reporting limits.

1.2 These procedures are applicable when low part per billion analyses are required to monitor differences between burdens in organisms from relatively uncontaminated reference areas and contaminated areas. In addition, the procedures are applicable when low detection limits are required for the estimation of potential health effects of bioaccumulated substances.

2.0 Summary of Method

2.1 Sets of 12-16 homogenized tissue samples are scheduled for extraction by the project lead chemist. Extraction methods employed were developed and validated by the Water Pollution Control Laboratory. Extract cleanup and partitioning methods are modifications of the multi-residue methods for fatty and non-fatty foods described in the U.S. Food and Drug Administration, Pesticide Analytical Manual, Vol. 1, 3rd Edition 1994, Chapter 3, Multi-residue Methods, Section 303-C1.

Homogenized tissue samples are removed from the freezer and allowed to thaw. A separate extraction bench sheet is initiated for each project, sample matrix type, and analysis type.

2.2 A 1-5 g (tissue homogenate) sample is weighed into a pre-weighed aluminum planchet and placed in a 70oC oven for 48 hours to determine moisture content. A 10 g sample is mixed using a clean glass stirring rod with

Table 1. Organochlorine Compounds Analyzed and Their Minimum Detection Limits (MDL) and Reporting Limits (RL) in Tissue.

MDL ng/g RL ng/g
wet wt. wet wt.

Aldrin 0.26 1.0
Chlordane, cis 0.68 2.0
Chlordane, trans 0.40 2.0
Chlordene, alpha 0.26 1.0
Chlordene, gamma 0.25 1.0
Chlorpyrifos 0.81 2.0
Dacthal 0.58 2.0
DDD, o,p' 0.71 2.0
DDD, p,p' 0.84 2.0
DDE, o,p' 0.53 2.0
DDE, p,p' 0.56 2.0
DDMU, p,p' 1.1 3.0
DDT, o,p' 1.0 3.0
DDT, p,p' 2.0 5.0
Diazinon 6.4 20
Dichlorobenzophenone, p,p' TBD 10
Dicofol (Kelthane) NR NR
Dieldrin 0.40 2.0
Endosulfan I 0.74 2.0
Endosulfan II TBD 10
Endosulfan sulfate TBD 10
Endrin 0.71 2.0
Ethion 1.9 6.0
HCH, alpha 0.36 1.0
HCH, beta 0.56 2.0
HCH, gamma 0.27 1.0
Heptachlor 0.51 2.0
Heptachlor epoxide 0.37 1.0
Hexachlorobenzene 0.10 0.3
Methoxychlor 1.3 5.0
Mirex 0.93 3.0
Nonachlor, cis 0.96 2.4
Nonachlor, trans 0.35 1.0
Oxadiazon 0.88 3.0
Oxychlordane 0.29 1.0
Parathion, ethyl 0.64 2.0
Parathion, methyl 1.2 4.0
Tetradifon (Tedion) 0.54 2.0
Toxaphene TBD 20

Table 2. PCB Congeners and Aroclor mixtures Analyzed and Their Detection Limits in Tissue, ng/g wet weight.


NIST Congeners:

PCB Congener 8 PCB Congener 128
PCB Congener 18 PCB Congener 138
PCB Congener 28 PCB Congener 153
PCB Congener 44 PCB Congener 170
PCB Congener 52 PCB Congener 180
PCB Congener 66 PCB Congener 187
PCB Congener 87 PCB Congener 195
PCB Congener 101 PCB Congener 206
PCB Congener 105 PCB Congener 209
PCB Congener 118

Additional Congeners:

PCB Congener 5 PCB Congener 137
PCB Congener 15 PCB Congener 149
PCB Congener 27 PCB Congener 151
PCB Congener 29 PCB Congener 156
PCB Congener 31 PCB Congener 157
PCB Congener 49 PCB Congener 158
PCB Congener 70 PCB Congener 174
PCB Congener 74 PCB Congener 177
PCB Congener 95 PCB Congener 183
PCB Congener 97 PCB Congener 189
PCB Congener 99 PCB Congener 194
PCB Congener 110 PCB Congener 201
PCB Congener 132 PCB Congener 203

All individual PCB Congener reporting limits are 0.2 ng/g wet weight.

Aroclors: Detection Limits ng/g wet wt.

Aroclor 1248 25
Aroclor 1254 10
Aroclor 1260 10
Aroclor 5460 (polychlorinated terphenyl) 100

approximately 7 g of pre-extracted Hydromatrix7 in a glass beaker until the mixture is free flowing. The mixture is then poured into a 33 mL stainless steel Dionex Accelerated Solvent Extractor (ASE 200) extractor cell and packed by tamping the mixture. A solution containing pesticide and PCB surrogate compounds is added to the cell and the cap is screwed onto the cell. The extractor cells (maximum of 24) are placed on the ASE 200 autosampler rack and the samples are extracted with a 50/50 mixture of acetone/dichloromethane (DCM) using heat and pressure.
The extracts are automatically collected in 60 mL VOA vials.

2.3 The extracts are dried using sodium sulfate, evaporated to approximately 0.5 mL using Kuderna-Danish (K-D) glassware equipped with 3-ball Snyder columns and micro-Snyder apparatus and diluted to 5 mL using DCM. The extracts are then filtered through a 0.45 μm syringe filter into ABC Autoprep 2000 Gel Permeation Chromatograph (GPC) autosampler tubes. One milliliter each of the filtered extracts is removed and placed in a pre-weighed aluminum planchet for percent lipid determination.

2.4 The GPC autosampler tubes are then placed on the GPC autosampler for initial sample cleanup. Samples containing less than 10% lipid are cleaned up using the Asmall@ GPC column. Samples containing greater than 10% lipid are cleaned up using the Alarge@ GPC column.

2.5 The cleaned-up extracts are evaporated using K-D apparatus and solvent exchanged into petroluem ether. The extracts are then fractionated using a standard 4 inch x 22 mm Florisil7 column. The Florisil7 columns are eluted with petroleum ether (PE) (Fraction 1), 6% diethyl ether/PE (Fraction 2), 15% diethyl ether/PE (Fraction 3), and 50% diethyl ether/PE (Fraction 4). The fractions are concentrated to an appropriate volume using K-D/micro K-D apparatus prior to analysis by dual column high resolution gas chromatography. A mixture of synthetic organic standards is eluted through the Florisil7 column to determine the recovery and separation characteristics of the column. The distribution of synthetic organic compounds in the four fractions is listed in Table 3.

Table 3. Distribution of Synthetic Organic Compounds Among the Four Fractions of a Standard Florisil7 Column.


(0%) Fraction 1/ (6%) Fraction 2/ (15%) Fraction 3/
HCH alpha 5/ HCH alpha 5/ dacthal
aldrin HCH beta diazinon
cis chlordane 5/ HCH gamma p,p' dichloroben
chlordane, alpha HCH delta zophenone
chlordane, gamma chlorbenside dieldrin
op' DDE cis chlordane 5/ endosulfan I 6/
pp' DDE 5/ trans chlordane endrin
pp' DDMU Chlorpyrifos endosulfan II 7/
op' DDT 5/ op' DDD oxadiazon
pp' DDT 5/ pp' DDD parathion, ethyl
heptachlor pp' DDE 5/ parathion, methyl
hexachlorobenzene 5/ pp' DDMS tetradifon
trans nonachlor 5/ op' DDT
PCB 1248 pp' DDT 5/ (50%) Fraction 4/
PCB 1254 ethion endosulfan II 7/
PCB 1260 heptachlor epoxide endosulfan sulfate
PCT 5460 hexachlorobenzene (HCB) 5/
methoxychlor
cis nonachlor 5/
oxychlordane
toxaphene


____________________

1/ 0% ethyl ether in petroleum ether.
2/ 6% ethyl ether in petroleum ether.
3/ 15% ethyl ether in petroleum ether.
4/ 50% ethyl ether in petroleum ether.
5/ In both 0% and 6% fractions.
6/ In both 6% and 15% fractions.
7/ In both 15% and 50% fractions.

3.0 Interferences

3.1 Solvents, reagents, glassware, and other sample processing hardware may cause GC artifacts and/or elevated baselines, resulting in the misinterpretation of chromatograms. All materials should be demonstrated to be free from interferences under the conditions of the analysis by running method blanks initially and with each sample lot. Specific selection of reagents and purification of solvents by distillation in all glass systems are required. High purity, distilled in glass solvents are commercially available.

An effective way of cleaning laboratory glassware is by rinsing with polar and non polar solvents before use. The cleaning procedure used must be tested by analyzing procedural blanks prior to analyzing samples.

3.2 Phthalates are common laboratory contaminants that are used widely as plasticizers. Sources of phthalate contamination include plastic lab-ware, plastic tubing, plastic gloves, plastic coated glassware clamps, and have been found as a contaminant in Na2SO4.
Polytetrafluoroethylene (PTFE) can be used instead of polypropylene or polyethylene to minimize this potential source of contamination. However, use of PTFE lab-ware will not necessarily preclude all phthalate contamination.

3.3 Interferences co extracted from tissue samples limit the method detection and quantitation limits. For this reason, sample extract cleanup is necessary to yield reproducible and reliable analyses of low level contaminants in the tissue sample.

4.0 Apparatus and Materials

4.1 Wide mouth, borosilicate glass, pre-cleaned and certified, 250 mL, Qorpak or equivalent.

4.2 Chromatographic Column 300 cm x 22 cm borosilicate glass chromatography column with Teflon stopcock.

4.3 Glass wool, Pyrex - solvent washed prior to use.

4.4 Kuderna Danish (K D) Apparatus

4.4.1 Concentrator tube 10 mL, graduate (Kontes K0570050 1025, or equivalent). A ground stopper, 19/22 joint, is used to prevent evaporation of extracts.

4.4.2 Evaporation flask 500 mL (Kontes K 570050 0500, or equivalent), attached to concentrator tube with blue clamp (Kontes K 662750 0012).


4.4.3 Snyder column three ball (Kontes K 503000 0121, or equivalent).

4.4.4 Micro-Snyder column - (Kontes VWR KT569261-0319 or equivalent).

4.4.5 Boiling chips, Hengar granules, high purity amphoteric alundum extracted with acetone and petroleum ether. Note that boiling chips can be a significant source of contamination if not properly cleaned.

4.5 Water bath, Organomation Assoc. Inc.(OA-SYS/S-EVAP-KD), 115 V, thermostatically controlled with stainless steel cover to fit 5 K-D apparatus, installed in a fume hood.

4.6 Nitrogen evaporator/water bath, Organomation Assoc. Inc.(N-EVAP 112), 115 V, thermostatically controlled with stainless steel cover for culture tubes, installed in a fume hood.

4.7 Extractor, automated, Dionex Accelerated Solvent Extractor (ASE 200), Dionex P/N 047046.

4.7.1 Extraction Cells, 33 mL, Dionex P/N 049562

4.7.2 Filters, cellulose for ASE extraction cells, Dionex P/N 049458.

4.7.3 VOA Vials, 60 mL, pre-cleaned and certified.

4.8 Sample vials glass, 2 5 mL with PTFE lined screw cap.

4.9 Analytical balance capable of weighing 0.1 mg.

4.10 Drying oven.

4.11 Balance capable of 100 g to the nearest 0.01 g.

4.12 Disposable Pasteur Pipets (rinsed with solvents before use).

4.13 Aluminum dishes for moisture and lipid determination.

4.14 Desiccator with indicating desiccant.

4.15 Glass funnel, 75 mm.

4.16 Graduated cylinder, 250 mL and 100 mL.

4.17 Culture tubes, 16 x 100 mm, with PTFE lined cap.

4.18 Gas chromatographs (2), Hewlett-Packard HP 6890 plus, equipped with two micro ECD detectors with EPC, split-splitless injector with EPC, and autosampler.

4.19 Capillary columns, 60 meter DB5 and 60 meter DB17 (J&W Scientific) (0.25 mm I.D. and 25 μm film thickness) connected to a single injection port using a "Y" press fit connector.

4.20 Data System, Hewlett-Packard, to collect and record GC data, generate reports, and compute and record response factors for multi-level calibrations. Data system should be capable of calibrating a method using a minimum of 5 concentrations of analytical standards.

4.21 Homogenizer, Bucchi Model B-400 (Brinkman P/N 16-07-200-1) or equivalent equipped with titanium knife assembly (Brinkman P/N 16-07-222-2) and glass sample vessel (Brinkman P/N 16-07-245-1).

4.22 Homogenizer, Brinkman Polytron or equivalent equipped Teflon and titanium generator assembly (for homogenization of small sample amounts).

4.23 Gel Permeation (size exclusion) Chromatograph, automated, J2 Scientific AccuPrep 170, equipped with
70 g S-X3 BioBeads J2 Scientific P/N C0100 (100% DCM).

5.0 Reagents

5.1 Petroleum ether (PE), Burdick and Jackson, distilled in glass and pesticide residue or HRGC grade or equivalent.

5.2 Acetone. (Same as above).

5.3 Iso Octane. (Same as above).

5.4 Diethyl ether preserved with 2% ethanol.(Same as above).

5.5 Dichloromethane (DCM). (Same as above).

5.6 Chem Elut-Hydromatrix7, Varian P/N 0019-8003. Pre-extracted on ASE-200 with acetone/DCM prior to use.

5.7 Sodium sulfate. Anhydrous granular reagent grade,
rinsed with PE prior to use.

5.8 Florisil7, 60/100 mesh, PR grade, Floridin Corp.

5.9 Nitrogen, pre-purified grade (99.9999%) or better
(used for ASE, GPC and solvent evaporation).

5.10 Nitrogen, ultra-pure (99.99999%) for ECD makeup.

5.11 Helium, ultra-pure (99.99999%) for GC carrier gas.

5.12 Air, compressed, breathing quality, for ASE pneumatics.

5.13 OC Surrogate Mix containing: 100 ppb of DBOB, dueterated p,p=-DDD, PCB 207, and DBCE.

CAUTION

The toxicity or carcinogenicity of each compound or reagent used in this method has not been precisely determined. However, each chemical compound should be treated as a potential health hazard. Exposure to these compounds should be reduced to the lowest possible level. The laboratory is responsible for maintaining a current awareness file of OSHA regulations regarding the safe handling of the chemicals specified in this method. A reference file of data handling Material Safety Data Sheets should also be made available to all personnel involved in these analyses.


6.0 Sample Collection, Preparation, and Storage

6.1 In the field, sources of contamination include sampling gear, grease from ship winches or cables, ship and/or motor vehicle engine exhaust, dust, and ice used for cooling. Efforts should be made to minimize handling and to avoid sources of contamination. This will usually require that resection (i.e., surgical removal) of tissue be performed in a controlled environment (e.g., a laboratory). The samples should be double wrapped in aluminum foil and immediately frozen with dry ice in a covered ice chest. Ice should be in water tight plastic bags for transporting live shellfish.

6.2 To avoid cross contamination, all equipment used in sample handling should be thoroughly cleaned before each sample is processed. All instruments must be of a material that can be easily cleaned (e.g., stainless steel, anodized aluminum, or borosilicate glass). Before the next sample is processed, instruments should be washed with a detergent solution, rinsed with tap water, rinsed with a high purity acetone, and finally rinsed with Type II water.


6.3 Resection should be carried out by or under the supervision of a competent biologist. Each organism should be handled with clean high carbon steel, titanium, quartz, or Teflon instruments (except for external surfaces). The specimens should come into contact with pre-cleaned glass surfaces only. Polypropylene and polyethylene surfaces are a potential source of contamination and should not be used. To control contamination when resecting tissue, separate sets of utensils should be used for removing outer tissue and for resecting tissue for analysis. For fish samples, special care must be taken to avoid contaminating target tissue (especially muscle) with slime and/or adhering sediment from the fish interior (skin) during resection. The incision "troughs" are subject to such contamination; thus, they should not be included in the sample. In case of muscle, a "core" of tissue is taken from within the area bordered by the incision troughs, without contacting them. Unless specifically sought as a sample, the dark muscle tissue that may exist in the vicinity of the lateral line should not be mixed with the light muscle tissue that constitutes the rest of the muscle tissue mass.

6.4 The resected tissue sample should be placed in a clean glass or PTFE container which has been washed with detergent, rinsed twice with tap water, rinsed once with distilled water, rinsed with acetone, and, finally, rinsed with high purity petroleum ether.

6.5 The U.S. EPA has published a guidance document containing specific recommendations regarding holding times and temperatures for tissue samples to be analyzed for semi volatile organic compounds. The following holding conditions should be observed. Tissue samples should be maintained at < 20o C and analyzed as soon as possible, but within 12 months of sample receipt.

7.0 Sample Extraction

7.1 Remove homogenized tissue samples from freezer and allow to thaw. Prior to extraction, the tissue samples are homogenized using a Bucchi B-400 mixer equipped with a titanium knife assembly or for small samples a Brinkman Polytron7 equipped with a titanium and Teflon generator. Sample sets of 12-16 should be extracted when possible. The ASE-200 extractor will extract 24 cells. Be sure to reserve enough cells for method blanks, matrix spikes, and laboratory control spikes.

7.2 A separate extraction bench sheet is started for each project, sample matrix type, and analysis type. Several bench sheets may be used for an extraction set.

7.3 Prepare a glass rod or Teflon spatula for each sample to be weighed by rinsing 3 times with petroleum ether using a Teflon wash bottle.

7.4 Label 60 mL VOA vials for the collection of the sample extract. The labels must be placed between 1.5" and 3" from the top of the VOA cap, if they are placed outside of this area they will interfere with the ASE optical sensor. Use two VOA vials for each sample to be extracted. Label with Project Number (eg. L#) or Project Name (eg. TSM) and the sample identifier with the second VOA vial for each sample additionally labeled ARE-EXTRACT@. Label and weigh aluminum planchets for lipid and moisture determinations (samples ID can be made on the bottom of planchets using a ball point pen.

7.5 Tare 250 mL glass jar. Using a clean glass rod, stir the tissue and make sure that water has not separated from the tissue. Weigh 10 g of tissue into the jar, record the weight on the bench sheet, and add the Hydromatrix from one ASE cell. Stir the mixture thoroughly and go on to the next sample. After approximately 15 minutes stir the sample again. Repeat this at 15 minute intervals two more times or until the sample mixture is free flowing.
7.6 Weigh 1-5 g of additional sample into a pre-weighed and tared aluminum planchet for % moisture analysis. Place planchets in 70oC oven for 48 hours and re-weigh dry weight.

7.7 Place a pre-rinsed powder funnel on top of a 33 mL ASE cell containing a pre-extracted cellulose filter (the filter is the one that was used to pre-extract the Hydromatrix).

7.8 Pour the tissue/Hydromatrix mixture through the powder funnel back into the extraction cell that the Hydromatrix was poured from. Tap the cell against the counter top to settle the contents. The mixture will fill the cell and it may be necessary to pack it slightly using the glass rod and the end of the powder funnel. The cells used for the method blank and laboratory control spike and its duplicate (if used) will contain only Hydromatrix.

7.9 All of the extraction cells are spiked with the pesticide and PCB surrogate standard. Spike each cell with exactly 1.0 mL of the pesticide surrogate solution labeled P110498A (40 ng/mL). Surrogate spikes must be witnessed, recorded and dated on the extraction bench sheet.

7.10 The extraction cells used for the matrix spike (MS) and duplicate matrix spike (MSD) and laboratory control spike (LCD) and its duplicate (LCSD) (if used) are spiked with exactly 1.0 mL of the pesticide matrix spike solution labeled P110498B (40 ng/mL). Matrix spikes must be witnessed, recorded and dated on the extraction bench sheet.

7.11 The extraction cells are capped (finger tighten only) and placed on the ASE 200 carrousel. The first set of labeled VOA collection vials are placed on the ASE 200 collection carrousel with the position numbers corresponding to the numbers of the extraction cells. Make sure that the solvent reservoir contains enough solvent for the extraction.

7.12 Samples are extracted with acetone/methylene chloride (DCM) 50:50 using the following conditions:

Pre-heat 0 min.
Heat 5 min.
Static 5 min.
Flush 60%
Purge 300 sec.
Cycles 1
Pressure 1500 psi
Temp 100 oC
Sol A Other 100%

7.13 After the initial extraction is complete, remove full VOA vials and place in a Nalgene rack and replace collection VOA vials with the vials labeled RE-EXTRACT. Check each of the extraction cells to make sure that the caps are finger tight as they tend to loosen with the first extraction. Make sure that the replacement vials are in the correct order. Make sure that the solvent reservoir contains enough solvent for the re-extraction. Re-start the ASE-200.

7.14 When extraction is completed place VOA vials in a Nalgene rack with the RE-EXTRACT vials next to the vials from the first extraction. The extracts should be re-capped with solid green caps (Qorpak) and placed in a refrigerator for storage until they are removed for the GPC cleanup procedure.


8.0 Gel Permeation Chromatography

IMPORTANT: All glassware, glass wool, and sodium sulfate must be triple-rinsed with petroleum ether before they are used for this procedure.

8.1 Remove VOA vials containing the sample extracts from the refrigerator. Make sure the vials are capped with the green Qorpak caps. Add a small amount of pre-cleaned sodium sulfate (~5 g) to the VOA vials and shake immediately. Add sodium sulfate until it is free-flowing in the VOA vial. MAKE SURE TO SWIRL OR SHAKE VIAL IMMEDIATELY AFTER ADDING THE SODIUM SULFATE OR IT WILL CAKE IN THE BOTTOM OF THE VIAL.


8.2 Set up and label pre-cleaned K-D flasks (4-6) with concentrator tubes attached on ring stands in the fume hood. Add a solvent rinsed micro-boiling chip to each K-D concentrator tube. Place a funnel containing a plug of pre-cleaned glass wool in the bottom of the funnel and place the funnel in the top of the K-D flask. Add about two inches of sodium sulfate to the funnel.

8.3 Pour sample extracts from the VOA vials through sodium sulfate into the K-D flask. Add about 10 mL of DCM to the VOA vial cap and shake and add this rinse to the sodium sulfate. Repeat with another 10 mL DCM rinse. Rinse the sodium sulfate with an additional portion of DCM (~10-20 mL). Use a small clean glass beaker to transfer DCM for rinses, use Teflon wash bottle for rinsing glassware only....never for dispensing DCM.

8.4 Place a Snyder column on the K-D flask, clamp with a green clamp and place the flask on the hot water bath set at 80-82oC. Evaporate the solvent until the reflux line falls below the top of the Snyder column. At this point there should be between 1-5 mL visible in the concentrator tube while the K-D apparatus is still on the hot water bath and 10 mL or less of the solvent remaining after the K-D flask is removed from the hot water bath and the solvent drains from the Snyder column.

8.5 After the K-D apparatus has cooled and all of the solvent has drained from the Snyder column, remove the Snyder column, label the concentrator tube and then remove the concentrator tube from the flask and place the tube in a test tube rack and cover with pre-rinsed aluminum foil. Rinse the Snyder column with petroleum ether and place back in the column rack for storage. After all of the flasks have been removed from the hot water bath repeat steps 1-5 for the remaining samples extracted with this set.

8.6 Add a new micro-boiling stone and place a clean micro-Snyder column on the concentrator tube with a blue clamp and place in a 400 mL beaker containing hot water heated to approximately 75oC on a hot plate . If the solvent does not begin to boil, remove the tube from the bath immediately, allow it to cool slightly, add a new micro boiling stone to prevent it from bumping and place it back in the bath. Evaporate the solvent until only 0.5 mL remains in the concentrator tube. Four or five tubes can be evaporated at one time.

8.7 When the solvent has been evaporated to 0.5 mL remove the tube from the bath and allow it to cool in a test tube rack. Remove the micro-Snyder column and add DCM to the concentrator tube to reach a final volume of 5.0 mL.

8.8 Draw the sample up into a clean 10 mL syringe with a 0.45 μm filter attached. Filter the sample into a 12 mL culture tube. Using a disposable pipet remove one mL of the filtered sample and place it in a pre-weighed aluminum planchet for lipid determination. Place the culture tube in the autosampler of the GPC.

8.9 Samples to be analyzed for pesticide (SO) compounds should be cleaned up using GPC method 1. Samples to be analyzed for PCBs only use GPC Method 2. Start the GPC using the following procedure:

8.9.1 Turn on the pump and nitrogen gas, set the column switch to Inline and allow solvent to pump through the system for about 30 minutes to allow column to stabilize and to rinse out any contaminants remaining in the column and/or detector. Make sure the solvent rinse and eluent reservoirs are full of DCM. Check to make sure that no bubbles of air are entering the detector from the column. Set up the GPC for calibration by entering the following:

Dump Time: 30 minutes (small column)
60 minutes (large column)
Collect Time: None

8.10.2 Measure and adjust the flow rate to 5 mL/min. Load calibration solution using a 10 mL syringe and 5 mL of pre-filtered GPC calibration standard solution in a dry GPC sample vial. Check UV detector parameters (0.2 AUFS, 0.3 sec Rise Time) and system pressure (10-15 psi) and make sure that the recorder is turned on with the following settings (50 mV and 20 cm/hr). Run the calibration solution and obtain a UV chromatogram showing a discrete peak for each calibration component.

8.10.3 Set the GPC time for each component of the calibration solution.
SO Compounds (pesticides and PCBs):
Set dump time to remove 85% of the corn oil and collect time to recover 85-90% of the phthalate and stop collection prior to the sulfur.
PCBs Only
Set dump time which removes >85% of the phthalate but recover >95% of methoxychlor and stop the collection after perylene but prior to the sulfur.

8.11 The GPC eluate is collected in 125 mL pre-cleaned Boston round bottles if small GPC column is used or 250 mL Erlenmeyer flask if the large column is used. Pour GPC eluate into K-D flask and add 0.5 mL of iso-octane (2,4,5-trimethyl pentane) to each flask as a Akeeper@. Add a micro boiling chip, attach a Snyder column to the flask and evaporate solvent on the hot water bath. When the apparent volume of solvent in the concentrator tube is 5-10 mL, add 20-30 mL of petroleum ether through the top of the Snyder column. Repeat this procedure when the apparent volume is again at 5-10 mL. Repeat a third time. When the reflux line falls below the top of the Snyder column, the K-D apparatus should be removed from the hot water bath. Remove the concentrator tube from the K-D apparatus.

8.12 Place a clean micro-Snyder column on the concentrator tube with a blue clamp , add a boiling chip and place in a 400 mL beaker containing water heated to approximately 85oC on a hot plate . If the solvent does not begin to boil, remove the tube from the bath immediately, allow it to cool slightly, add a new micro boiling stone to prevent it from bumping and place it back in the bath. Evaporate the solvent until only 0.5 mL remains in the concentrator tube. Four or five tubes can be evaporated at one time. Allow the solvent to Acook@ for about 2 minutes. Remove the evaporation assembly from the water bath and allow it to cool. Add petroleum ether to achieve a final volume of 10 mL and gently mix solution.

8.13 Transfer the solution from the concentrator tube to a culture tube and cap with a Teflon faced cap. Place extracts in a refrigerator for storage until the final Florisil7 column cleanup is done.


9.0 Florisil7 Column Fractionation

IMPORTANT: All glassware, glass wool, and sodium sulfate must be triple-rinsed with petroleum ether (PE) before they are used for this procedure. Florisil7 must be activated in an oven at 130oC for at least 24 hours prior to use.

9.1 This procedure is performed after the GPC cleanup procedure for all tissue samples analyzed for pesticides and PCBs. No more than 30 minutes prior to performing the Florisil7 cleanup, add a small amount (~1 g) of sodium sulfate to the culture tubes to remove residual water from the extract. If the sodium sulfate becomes a solid plug in the bottom of the tube, add more until some of the sodium sulfate is free flowing when the tube is shaken. If extracts are allowed to remain in contact with sodium sulfate for longer that 30 minutes, target analyte loss may result.

9.2 Prepare the reagents to be used for Florisil7 cleanup: 6% ethyl ether in petroleum ether, 15% ethyl ether in PE, and 50% ethyl ether in PE. Make an amount slightly in excess of what is actually needed to allow for any loss which may occur during solvent transfer. The required volume is 200 mL per sample for the 6% (F2), 15%(F3), and 50%(F4) fractions. Fill the 250 mL separatory funnels located above the Florisil7 columns with 250 mL of petroleum ether (0% or F1 fraction). These funnels will be used for eluant reservoirs.


9.3 Prepare the chromatography columns. Place a small piece of PE rinsed glass wool in the bottom of the column and tap into place with a PE rinsed glass rod. Cover with a small portion (0.5 inch) of sodium sulfate. Measure 4 inches from the top of sodium sulfate and mark column outside of the column with a permanent marker. Fill the column with Florisil7 to about 3/4 inch beyond the mark and tap column with rubber "mallet" to firmly settle the Florisil7. Add more Florisil7 as necessary so that it is even with the mark after settling. Top column with 3/4-1 inch of sodium sulfate. This will prevent the column from being disrupted when solvent is added and will remove any residual water. Tie a Kimwipe around the column to catch any condensation or accidental overflow which could roll down the outside of the column and contaminate the sample.

9.4 Place a 600 mL beaker under the column and pre wet the column with about 40 mL of petroleum ether. Filling the column to 1 inch above the "Kimax" label is usually sufficient.

IMPORTANT: From this point and through the elution process, the solvent level should never be allowed to go below the top of the sodium sulfate layer and the column stopcock should never be closed.

9.5 When approximately 1 inch of PE remains above the surface of the column, place a K-D flask under column and adjust flow rate to about 5 mL/min (32 drop/12 sec). When the meniscus of the rinse PE reaches the column bed surface, pour the sample extract onto the column. Immediately add 5 mL of PE to the tube, shake vigorously, and set aside. When the collected volume reaches 10 mL, pre-wet the next column. If the columns are started in this sequential fashion, the lag time will be adequate to perform the necessary tasks for up to six columns.

9.6 When the sample extract reaches the sodium sulfate layer, add the PE rinse from the culture tube. Add another 5 mL to the culture tube, shake and immediately add this rinse to the top of the column. Repeat rinse a third time. When the final rinse reaches the sodium sulfate layer, fill the column one half full with PE from the reservoir. Adjust the drip rate from the separatory funnel to approximately equal that from the column tip. Try to keep the solvent level in the column constant to avoid variations in flow rate.

9.7 When all of the F1 solvent has been transferred to the column from the solvent reservoir, close the reservoir stopcock and fill the separatory funnel with 200 mL of the 6% diethyl ether/PE mixture. Just before the PE reaches the sodium sulfate layer, change the K-D flask. Add 0.5 mL of PCB 207 surrogate to the flask (this is added only to the F2, F3, and F4 flasks because PCB 207 is used to determine relative retention times during GC analysis). When the solvent reaches the sodium sulfate, add the 6% diethyl ether/PE to the column and elute as before. Add a micro boiling stone and attach a Snyder column with a green clamp to the K-D flask containing the 0% (F1) fraction and place vessel in the hot water bath with the temperature set at 80-82 oC and reduce volume to an apparent volume of 1 mL. Tap the Snyder column to make sure solvent is not trapped between the balls then remove the vessel from the bath and place in the vessel stand to cool.

9.8 Repeat the above using 200 mL of 15% diethyl ether/PE mixture.

9.9 Repeat the above using 200 mL of 50% diethyl ether/PE mixture.

9.10 When the vessels are cool, remove the concentrator tube from the K-D flask add a new micro boiling stone and attach a clean micro-Snyder column to the concentrator tube with a blue clamp and place in a 400 mL beaker containing hot water heated to approximately 85oC on a hot plate. Evaporate the solvent until only 0.5-1 mL remains in the concentrator tube. Four or five tubes can be evaporated at one time.

9.11 When the solvent has been evaporated to 0.5-1 mL remove the tube from the bath and allow it to cool in a test tube rack. Remove the micro-Snyder column and add iso-octane to the concentrator tube to reach a final volume of 2.0 mL. Mix the tube contents by tapping the bottom of the tube causing a vortex which will rinse the sides of the tube. A Vortex Genie mixer may be used for this step. Transfer the extract to a clean labeled culture tube and cap.

9.12 Repeat for 6% (F2), 15% (F3), and 50% (F4) extracts. The extracts are ready for analysis by GC-ECD.

10.0 Analytical Procedure

10.1 Chlorinated hydrocarbon analysis:

10.1.1 Hewlett-Packard 6890plus gas chromatograph equipped with two 63Ni micro-electron capture detectors with EPC and autosampler. Two 60 meter, 0.25 mm ID, 0.25 um (film thickness) fused silica columns (J&W) are used. A 5 meter length of DB-5 column is connected to a press fit "Y" union which splits the column effluent into two 60 m columns, a DB 5 and a DB 17. The injector is a split-splitless injector with EPC.

10.1.2 Chromatograph conditions:
The injector is operated isothermal at 240oC. The oven has an initial temperature of 70oC and is immediately temperature programmed to 210oC at a rate of 15oC/min and held for 10 min. It is then programmed to 280oC at a rate of 2oC/min and is held for 11 min. Helium is used as the carrier gas at a linear velocity of 35 cm/sec. Nitrogen is used for the detector makeup at 30 mL/min.

10.1.3 Sample volume:
Three microliters of samples and standards are injected and split approximately 50/50 onto the 60 m DB 5 and the 60 m DB 17.

10.1.4 Data processing:
Detector signals are acquired and processed with a Hewlett-Packard Model 3365 Series II Chemstation.

11.0 References

Tetra Tech, Inc. 1986. Bio Accumulation monitoring Guidance: 4. Analytical Methods for U.S. Priority Pollutants and 301 (h) Pesticides in tissues from Estuarine and Marine Organisms. TC 3953 03. U.S. EPA Washington, DC.

U.S. Environmental Protection Agency. 1993. Guidance For Assessing Chemical Contaminant Data For Use In Fish Advisories, Volume I, Fish Sampling and Analysis. EPA 823-R-93-002. U.S. EPA, Office of Water, Washington D.C.

U.S. Food and Drug Administration. 1994. Pesticide Analytical Manual. Volume 1, Chapter 3, Multiclass Multiresidue Methods. U.S. Food and Drug Administration, Rockville, MD.



Marine Pollution Studies Laboratories
Department of Fish and Game
Moss Landing Marine Laboratories
7544 Sandholdt Road
Moss Landing, CA 95039

Project Manager: Autumn Bonnema
Phone: 831-771-4175
Fax: 831-633-0805
e-mail: bonnema@mlml.calstate.edu

Project Name: MBNMS Project Number:
Parameter: Trace Metals Matrix: Mussel Tissue
Report Number: TM07-007 Report Date: 04-30-07

QA/QC SUMMARY

SAMPLE CUSTODY

Nine Mussel samples were received in good condition April 19 and June 29, 2006. Samples were dissected in a clean lab in three replicates.


QA/QC DATA QUALITY OBJECTIVES (DQO)


Analyte Reference Method Range of Recovery Relative Precision Detection Limit Reporting Limit

As EPA 200.8 25% 25% 0.02 g/g wet 0.06 g/g wet
Cd EPA 200.8 25% 25% 0.002 g/g wet 0.006 g/g wet
Zn EPA 200.8 25% 25% 0.40 g/g wet 1.2 g/g wet
Pb EPA 200.8 25% 25% 0.006 g/g wet 0.02 g/g wet
Cu EPA 200.8 25% 25% 0.04 g/g wet 0.10 g/g wet
Mn EPA 200.8 25% 25% 0.02 g/g wet 0.06 g/g wet
Ni EPA 200.8 25% 25% 0.02 g/g wet 0.06 g/g wet
Se EPA 200.8 25% 25% 0.04 g/g wet 0.12g/g wet
Ag EPA 200.8 25% 25% 0.006 g/g wet 0.02 g/g wet
Cr EPA 200.8 25% 25% 0.06 g/g wet 0.20 g/g wet
Al EPA 200.8 25% 25% 2.00 g/g wet 6.00 g/g wet


METHOD

Samples were digested using EPA 3052 (Modified): Microwave Assisted Acid Digestion of Determination of Trace Metals in Ambient Waters by ICP-MS.




HOLDING TIME

Samples were dissected by Fish and Game and digested in January 2007. All samples were analyzed within the EPA holding time of 1 year from collection.


CALIBRATION VERIFICATION

Initial Calibration Verification (ICV) and all Continuing Calibration Verification (CCV) were within DQO of 10%.


DETECTION LIMIT

All detection limits listed in the table above were achieved.


METHOD BLANKS

Two method blanks were analyzed with each batch samples. All elements were below detection limits. Samples are all blank corrected with the average blank value for that batch.


REPLICATES

One pair of analytical duplicates selected at random was analyzed with each batch of samples. All RPDs met the DQO of 25%.


MATRIX SPIKES

One matrix spike/matrix spike duplicate (MS/MSD) pair was analyzed with each batch of samples. All recoveries and RPDs met the DQO of 25%


STANDARD REFERENCE MATERIAL

SRM 2976 was analyzed with each batch of samples. All recoveries met the DQO of 25% with the exception of Ag and Ni. The matrix spikes for Ag and Ni, however, were within range.


REFERENCES

US Environmental Protection Agency Method 3052. 1996. Microwave Assisted Acid Digestion of Siliceous and Organically Based Matrices. US Environmental Protection Agency, Washington, DC.

Modifications to EPA 3052

US Environmental Protection Agency Method 200.8. 1996. Determination of Trace Elements in Tissues by Inductively Coupled Plasma- Mass Spectrometry. US Environmental Protection Agency, Washington, DC. Modifications to EPA 1638



Figures and Images

Figure 1. Map of long-term Mussel Watch Program sites in Monterey Bay (red circles) and extra sites sampled in 2006 near the thermal outfall (yellow circles). Moss Landing Power Plant is indicated (orange circle).


Figure 2. Moss Landing Power Plant as seen from the entrance to Salinas River State Beach on February 13, 2009.


Figure 3. Aerial view of the thermal plume (white dot near harbor entrance) generated by the Moss Landing Power Plant. In comparison, the natural tidal plum created by Elkhorn Slough (brown ribbon) is extending well offshore and into the clearer waters of Monterey Bay. Used with permission by Dr. J. Paduan, Navy Postgraduate School.