Buzzards Bay caged mussel pilot biomonitoring study, 1987-1988

"October 1990." Includes bibliographical references (p. 46-48)...

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w PILOT BIOMONITORING

STUDY

1987-1988

I

XL?

Massachusetts Executive Office of Environmental

John

P. DeVillars,

Secretary

Department of Environmental Protection Daniel S.

Greenbaum, Commissioner

Water Pollution Control Brian M. Donahoe, Director

Division of

315DbbDltit DS4fi l

i:

]

Affairs

11

*** ^^.

BUZZARDS BAY CAGED MUSSEL PILOT BIOMONITORING STUDY 1987 - 1988

Prepared by

Christine L. Duerring Environmental Analyst

MASSACHUSETTS DEPARTMENT OF ENVIRONMENTAL PROTECTION DIVISION OF WATER POLLUTION CONTROL TECHNICAL SERVICES BRANCH WESTBOROUGH, MASSACHUSETTS FOR U.S. ENVIRONMENTAL PROTECTION AGENCY

REGION I WATER MANAGEMENT DIVISION BOSTON, MASSACHUSETTS

Massachusetts Executive Office of Environmental Affairs John P. DeVillars, Secretary Department of Environmental Protection Daniel S. Greenbaum, Commissioner

Division of Water Pollution Control Brian M. Donahoe, Director

OCTOBER 1990

Publication No. 1 6 500- 1 07-37- 1 1-90-CR Approved by: Ric Murphy, State Purchasing Agent ,

:

TITLE:

Buzzards Bay Caged Mussel Pilot Biomonitoring Study

DATE:

October, 1990

AUTHOR

Christine L. Duerring, Environmental Analyst III

REVIEWED BY:

APPROVED BY: ,/

u? Steven G. Halterman Environmental Engineer

y (^Yj^ br^4*

Alan N. Coopermapr, P.E. Supervisor, Technical Services Branch

ii

FOREWORD

The Division of Water Pollution Control in 1987 proposed and received funding from the U.S. Environmental Protection Agency (EPA) to conduct a pilot biomonitoring study in Buzzards Bay using caged mussels Mytilus edulis The study is part of a national estuarine management program developed by the U.S. EPA Office of Marine and Estuarine Protection and Region I of the EPA for Buzzards Bay. The program was initiated to promote and develop coordinated efforts between federal, state, local authorities, research institutions and the public to identify, correct, and monitor environmental problems affecting this nation's estuaries. (

111

)

.

ACKNOWLEDGMENTS

The following people and groups are gratefully acknowledged for their assistance in conducting various aspects of this study:

Patricia Austin, Lawrence Gil, Steve Halterman, Jay Loubris, Steve McGee, Cathy O'Riordan, crew and captain of the fishing boat Phalarope, and Gerry Szal for assisting in the collection and/or deployment of the mussels while maintaining a sense of humor in the face of (usually) adverse weather conditions; Brad Burke, New Bedford Shellfish Constable and his assistant David Goulart, who provided boat transportation and muscle in deploying and collecting the cages throughout the entire study. Their reliable assistance and knowledge of the area eased the field work effort considerably; Staff of the Lawrence Experiment Station, notably Ken Hulme, Rosario Grasso and Alba Flaherty for performing the special water quality and tissue analyses;

Ken Hulme together with Nina Dustin and Jack Schwartz of the Division of Marine Fisheries, Cat Cove Laboratory also offered helpful comments concerning interlaboratory calibration exercises; Bruce Tripp who offered encouragement and technical advice during the study;

Judy Capuzzo and Donald Phelps for helpful review of the data;

And lastly, but most important, this study would not have been possible without the use of the laboratory facilities at the EPA Environmental Research Laboratory in Narragansett, RI and the generous assistance and advice of Skip Nelson in designing and implementing the study.

TABLE OF CONTENTS

PAGE

TITLE

Acknowledgments

v

Abstract

ix

List of Tables

xi

List of Figures

xiii

Introduction

1

Methods and Description of Study Site

5

Results

14

Discussion

37

Summary

45

Bibliography

46

Appendix A:

Field and LES Laboratory Methodology

49

Appendix B:

Field and Laboratory Data

73

Appendix

Sample Statistical Calculations

85

Division of Marine Fisheries Project Plan and Laboratory Methodology

89

C:

Appendix D:

VII

Digitized by the Internet Archive in

2013 with funding from

Boston Library Consortium

Member

Libraries

http://archive.org/details/buzzardsbaycageOOduer

ABSTRACT

Buzzards Bay Caged Mussel Pilot Biomonitoring Study 1987 - 1988

A caged mussel pilot biomonitoring study was conducted in Clarks Cove, New Bedford/Dartmouth, Massachusetts from October 1987 to September 1988. Mussels were deployed at three stations for five consecutive, 60-day exposure periods. Mussel tissue was analyzed for the trace elements: As, Cd, Cr, Cu, Hg, Ni, Pb, Zn, as well as total and fecal coliform bacteria and polychlorinated biphenyls (PCBs), and percent lipid content before and after the exposure periods.

Trace element tissue concentration was extremely variable at all of the stations. Within station (replicate) variability was usually high and masked between station differences in trace element concentration for many of the deployments. However, significant differences were detected between baseline and one or more of the Clarks Cove Stations for tissue concentrations of arsenic, zinc, and lead None of the Clarks Cove Stations (A, B, for several of the exposure periods. or C) exhibited significant differences in trace element tissue concentration from each other, indicating bio-available trace element concentration was not spatially different in Clarks Cove.

Bacteria concentration in the mussel tissue was variable and showed no consistent Based on these results this technique is not pattern throughout the study. recommended for long-term monitoring of coliform densities in coastal areas. PCB tissue concentration between baseline and Clarks Cove Stations showed a consistent pattern of low baseline values, highest concentration at Station A, next highest at Station B, and low at Station C, indicating that this method may be effective for monitoring PCB concentration in coastal areas.

Inter-laboratory calibration exercises performed between the Lawrence Experiment Station and the Division of Marine Fisheries, Cat Cove Laboratory showed large inter-laboratory differences in results from mussel tissue analyzed for trace element concentration from the Clarks Cove study sites. However, results from similar analyses of EPA prepared standard "mega mussel" samples showed good inter-laboratory agreement.

ix

LIST OF TABLES

TABLE

TITLE

PAGE

1

Clarks Cove Combined Sewer Overflows

12

2

Clarks Cove Sediment Data: Trace Elements, PCBs, PAHs, Percent Total Volatile Solids

16

3

Percent Mortality of Mussels and Cage Loss

18

4

Mussel Tissue Total and Fecal Coliform Densities

20

5

Summary of Kruskal-Wallis Nonparametric Analysis of Variance

30

6

Summary of Tukey-type Nonparametric Multiple Comparison Tests

31

7

Inter-laboratory Calibration Results - Trace Element Concentrations

34

8

Inter-laboratory Calibration Results - Standard "Mega Mussel" Tissue

36

9

Tissue Trace Element Concentration - Comparison to Other Studies

41

A-l

Common Sample Treatment Methods

50

A-2

Parameter and Collection Methods Employed at Sediment Stations

51

A-3

Summary of Rated Accuracy of Field Meters and Unit of Measure

52

A-4

Parameters and Analytical Methods for Water Samples

53

A-5

Parameters and Analytical Methods for Sediment Samples

55

A-6

Parameters and Analytical Methods for Tissue Samples

57

A-7

Method for Chlorophyll a Analysis

59

B-l

Clarks Cove Water Quality Data - Field Measurements

74

B-2

Clarks Cove Water Quality Data - Chemical Parameters

77

B-3

Results of Mussel Tissue Trace Element Analysis

80

B-4

Mussel Tissue PCB Concentrations

83

B-5

Percent Lipid Concentration in Mussel Tissue

84

XI

LIST OF FIGURES

TITLE

FIGURE

PAGE

1

Clarks Cove Station Locations

6

2

Station Description

7

3

Clarks Cove Combined Sewer Overflows and Storm Drain Locations

11

4

Clarks Cove Temperature

15

5

Clarks Cove Dissolved Oxygen

15

6

Clarks Cove Salinity

15

7

Mean Mussel Shell Growth by Station and Deployment

19

8

Mussel Tissue Mercury Concentration

22

9

Mussel Tissue Zinc Concentration

23

10

Mussel Tissue Nickel Concentration

24

11

Mussel Tissue Lead Concentration

25

12

Mussel Tissue Copper Concentration

26

13

Mussel Tissue Cadmium Concentration

27

14

Mussel Tissue Chromium Concentration

28

15

Mussel Tissue Arsenic Concentration

29

16

Comparison of PCB Tissue Concentration

33

17

Lawrence Experiment Station vs. Division of Marine Fisheries Values as a Percent of EPA "Mega Mussel" Values

42

Xlll

.

INTRODUCTION

In 1987 the Massachusetts Division of Water Pollution Control (DWPC), Department of Environmental Protection (DEP) applied for and received funding from the U.S. Environmental Protection Agency (EPA) Buzzards Bay Project to conduct a pilot biomonitoring program in Clarks Cove, New Bedford, Massachusetts using caged This study is one of several being conducted in Mytilus ejdulis) mussels Bay Project over the past two years. the EPA Buzzards These Buzzards Bay for research projects are diverse and address water quality issues identified as being priority concerns in Buzzards Bay, mainly; bacterial contamination, nutrient enrichment, and toxic contaminants in fish and shellfish. Information gathered during this study phase will be used by the Buzzards Bay Project staff to develop a Comprehensive Conservation and Management Plan (CCMP) for Buzzards Bay. (

.

The CCMP will provide strategies for pollution abatement and prevention In addition, the CCMP will include throughout the watershed of the bay. recommendations for long-term monitoring to assess the effectiveness of the water quality clean-up and management techniques that are employed. The goals of this project were primarily to address questions relating to water quality monitoring techniques. In general, the "pilot" portion of the study was to design and implement a simple biomonitoring technique that could be performed by local, state, and/or regional agencies that would enable detection of longterm spatial and temporal trends in contaminant concentrations. More specifically, the study was to provide information that could be used to assess trace element and bacterial contamination in the water column of Clarks Cove, an area that receives discharges from as many as nine (9) combined sewer overflows from the City of New Bedford and flows from seven (7) storm drains from Dartmouth and New Bedford watersheds. In addition, the DWPC saw this as an opportunity to expand its water quality monitoring capabilities by examining this methodology for use as a tool to assess trace element contamination in sea water. The Massachusetts state analytical laboratory, the Lawrence Experiment Station (LES), does not have a "Clean bench" facility that is necessary to directly measure the trace concentrations of heavy metals and metalloids present in sea water.

Historically, the basic goal of water quality monitoring programs was to collect chemical and physical data which was used to characterize the general water quality of an area (Perry et. a. 1987). The design of many monitoring programs today still reflect this often random data gathering "objective", despite the fact that the intent and expectations of monitoring programs have matured. Monitoring programs are now relied upon to provide sound information on which to base management decisions. According to Segar, et. al. (1987) most marine monitoring programs have been inefficient or ineffective in providing specific information that can be used by the manager. These researchers recommend the use of transplanted bioindicator organisms to monitor temporal changes of bioavailable contaminants in an area. The test animals, suspended in the water column, ingest, filter and/or absorb what is biologically available to them, providing a time integrated measure of the abundance of specific bio-available contaminants

Within approximately the past fifteen years, the use of indicator organisms to These studies have monitor coastal water quality has become widely accepted. test animals. The most ideal used both transplanted (i.e., caged) or indigenous Capuzzo et. al. organisms for these types of studies appears to be bivalves. (1987) attribute the use of shellfish for these types of studies, particularly in monitoring heavy metals, to their metals bioaccumulation ability, sensitivity to metals concentration gradients, and importance to large programs such as the National Shellfish Sanitation Program and the Mussel Watch Program. They also point out, however, that there is no standard methodology for collecting these data sets.

Farrington et. al. (1987) and Tripp and Farrington (1984) presented the following comprehensive list of reasons why bivalves are considered the most useful organisms for this approach: 1.

This characteristic Bivalves are widely distributed geographically. minimizes the problems inherent in comparing data for markedly different species.

2.

They are sedentary and are thus better than mobile species as integrators of chemical pollution in a given area.

3.

They have reasonably high tolerances to many types of pollution, comparison to fish and Crustacea.

4.

in

2 5 They concentrate many chemicals by factors of 10 to 10 compared to seawater in their habitat making trace constituent measurements easier to accomplish in their tissues than in seawater.

5.

An assessment of biological availability of chemicals is obtained.

6.

In comparison to fish and Crustacea, bivalves exhibit low or undetectable activity of those enzyme systems which metabolize many xenobiotics such as aromatic hydrocarbons and PCBs. Thus, a more accurate assessment of the magnitude of xenobiotic contamination in the habitat of the bivalves can be made.

7.

They have many relatively stable, local populations that are extensive enough to be sampled repeatedly, providing data on short and long-term temporal changes in the concentrations of pollution chemicals.

8.

They survive under conditions of pollution that often severely reduce or eliminate other species.

9.

They can be successfully transplanted and maintained on subtidal moorings or on intertidal shore areas where populations normally do not grow thereby allowing expansion of areas to be investigated.

10.

They are commercially valuable seafood species on a worldwide basis. Therefore, measurement of chemical contamination is of interest for public health considerations.

Another advantage of using mussels and oysters that is relative to this particular study is that these animals can integrate pollutant levels over space and time, an advantage over sampling seawater and sediment for pollution assessment that can provide only very short-term (via seawater) or long-term (via sediments) contaminant integration (Goldberg, 1986). Specific advantages of using transplanted animals taken from a relatively unpolluted site and suspended in cages in the test area over sampling indigenous animals for contaminants are (de Kock and van het Groenewoud, 1985): 1) the animals are derived from a common stock, thereby reducing a potential source of variability when comparing geographical locations; 2) the period of exposure to the environment is known and can be controlled; 3) monitoring locations can be chosen, regardless of whether or not the animals occur there naturally. The EPA conducted a study in 1982 to evaluate the use of caged mussels to monitor ocean disposal of municipal sewage sludge in the New York Bidge (Phelps et. al., The study concluded that the use of transplanted caged mussels as a 1982). biomonitoring tool in coastal waters was feasible. Some of the large scale national water quality monitoring programs employing bivalves include the EPA Mussel Watch Program, which was conducted at over 100 sites around the coast during 1976-1978, and the current National Status and Trends Mussel Watch Program being conducted by National Oceanic and Atmospheric Administration (NOAA) on 150 coastal sites. In the United Kingdom, mussel watch programs were conducted from 1977-1979 at over two hundred sites along the coastlines of England, Wales, Scotland, and Ireland.

There are also more localized bioaccumulation studies using indicator organisms designed to monitor a specific point source. For example, the EPA has required bioaccumulation assessment plans to be included in several recent NPDES permits. These plans call for the use of Mvtvlis edulis (blue mussel) and Crassostrea virginica (eastern oyster) to monitor survivability and contaminant bioaccumulation at sites within the zone of initial dilution of the sewage outfalls. Massachusetts sewage treatment facilities that are currently developing a plan or are already conducting bioaccumulation studies as part of their NPDES permit requirement include the Lynn Water and Sewer Commission, Swampscott Wastewater Treatment Plant, South Essex Sewerage District (SESD), and the Massachusetts Water Resources Authority (MWRA) The EPA provides a guidance document entitled, "Methods for Use of Caged Mussels for In Situ Biomonitoring of Marine Sewage Discharges" (1983) that they recommend for use when designing bioaccumulation studies for these permits. Also in Massachusetts, caged mussel studies conducted by the New England Aquarium (1986, 1988) have been included as part of environmental impact studies to aid in the design and siting of ocean outfalls for SESD and MWRA. .

It is evident from the literature that this methodology has become widely used and accepted by researchers as well as environmental regulators. The Buzzards Bay Technical Advisory Committee (TAC) has recognized the importance of this technique in the development of a coastal monitoring program that would be capable of detecting water quality trends in space and time. The monitoring effort in Buzzards Bay requires efficient techniques that will enable scientists to characterize long-term temporal and spatial water quality changes that result from point and/or nonpoint pollution abatement strategies and/or deleterious activities that may occur within the watershed. Although biomonitoring guidance documents do exist (U.S. EPA, 1983), there still is no single, widely accepted

standard operating procedure for conducting these types of bioaccumulation studies. More over, there appears to be even less agreement on how to interpret With these problems and the needs of DWPC and the Buzzards Bay the results. this pilot study was designed to address the following mind, Project in objectives: 1.

To evaluate the impact of urban point sources of contamination into Buzzards Bay by assessing concentrations of selected trace elements and coliform bacteria in the tissues of the blue mussel (M. edulis) that have been suspended in cages at three sites located along a transect originating in Clarks Cove, New Bedford.

2.

To compare shell growth between stations in a percentage of the test animals.

3.

To examine the feasibility of this type of bio-indicator study as a water quality monitoring technique for the Division of Water Pollution Control.

4.

To conduct an inter-laboratory calibration exercise with the Division of Marine Fisheries to demonstrate the degree of variability between laboratories that may be encountered in a study of this kind.

This report also contains the results of mussel tissue PCB analysis, although this task was not included in the biomonitoring study funded by EPA. Results are reported and briefly discussed in this report mainly because the task was an integral part of this pilot study and the information it provides will be used by DWPC to assess the usefulness of this technique for monitoring PCB contamination in other coastal areas of Massachusetts.

METHODS AND DESCRIPTION OF STUDY SITE

Study Design

Arrays cages were deployed at three stations oriented along a north-south transect originating in Clarks Cove, New Bedford and extending approximately 5.6 km (3.5 mi) in a south-south easterly direction out into Buzzards Bay (Figure Station B was Station A was located near the head of Clarks Cove. 1). established at the mouth of the cove midway between the eastern and western shorelines. Station C was located in Buzzards Bay near Nun #4 LR approximately Water depth at Station 1.7 miles northeast of Round Hill Point in Dartmouth. A and B at low tide was approximately 5 meters and low tide depth at Station C was approximately 9 meters. By establishing stations in a land to seaward direction a contamination gradient was expected to be observed, with highest levels of metals and bacteria predicted in tissues collected from Station A at the head of the cove nearest the urban sources (i.e., combined sewer overflows and storm drains), and lowest levels anticipated in tissues from reference Station C located over 1 mile (1.6 km) (See pages 10 - 13 for a located away from land based pollution sources. Before establishing these station complete description of the study site. ) locations it was important to consider the influence of currents in the study area. Although little information has been published on the hydrodynamics of Clarks Cove, available research results supported the selection of a north-south transect on which to locate stations. In the main body of Buzzards Bay the currents are complex. Net displacement of a particle over a tidal cycle is about 102 km (EG&G for CDM) Signell (1987) characterized the circulation pattern in the bay as tidally dominated. Wind is also an important mechanism determining subtidal circulation especially in shallow embayments and estuaries. EG&G's survey described tidal currents in the New Bedford Clarks Cove area as running generally south to north-northeast into Clarks Cove on the flood tide and north to south-southwest on the ebb tide. .

Each station was located in an area of soft bottom sediments indicating that deposition, rather than scouring was taking place. This also enabled sediments to be collected for analyses from each site and helped maintain similarity between stations. The cage assembly was anchored by one or two 8"xl6" cinder blocks and attached to floating lobster buoys to mark their location. This design was identical to that used by the EPA, Environmental Research Laboratory (ERL) in Narragansett, RI for similar caged mussel biomonitoring studies they have conducted in New Bedford Harbor (Don Phelps and William Nelson, EPA, ERL, Narragansett, RI, personal communication). With this design, field personnel were able to set out and retrieve the cages from a boat rather than rely on scuba divers to access the cages. Each cage contained twenty-five (25) mussels Mvtilus edulis) with total shell lengths all between 5-7 cm. Figure 2 illustrates the design of the cage array for one station. For the growth study ten of the twenty-five animals in one cage of each replicate were marked with an individual number etched in the shell surface (methods employed for the growth study are described below). Each station was made up of four replicates. Each replicate consisted of 50 animals divided equally into two cages for a total of 200 animals per station. A typical exposure period, from deployment to collection lasted about sixty days with a new group of mussels set out each time. (

\

FIGURE

1

BUZZARDS BAY CAGED MUSSEL PILOT BIOMONITORING STUDY Oct.1987-Sept.l988 Clarks Cove Station Locations

FIGURE

2

BUZZARDS BAY CAGED MUSSEL PILOT BIOMONITORING STUDY Oct.l987-Sept.l988 Station Description: 4 Replicates per Station, 2 Cages per Replicate, 25 Mussels per Cage, 200 Mussels per Station

The EPA (1983) recommends a 30 day exposure time for metals bioaccumulation studies whereas de Koch and van het Groenewoud (1985) state that some metals may require over 150 days to bioaccumulate in mussels. After discussions with the Buzzards Bay Technical Advisory Committee and personnel from Woods Hole and EPA, This allowed for ERL, Narragansett the 60 day exposure period was selected. twice the EPA recommended exposure time. Longer periods were rejected to avoid or minimize the degree of fouling that may occur on the cages and to reduce cage The one year loss due to wear and tear from extended periods of weathering. study period that began in October thus allowed for five, 60 day exposure periods, or deployments, that occurred on the following dates:

First deployment Second deployment Third deployment Fourth deployment Fifth deployment

-

-

October 29, 1987 - January 13, 1988 January 13, 1988 - March 16, 1988 March 16, 1988 - May 11, 1988 May 11, 1988 - July 13, 1988 July 13, 1988 - September 21, 1988

Field and Laboratory Procedures The same field procedures were followed for each deployment period. Blue mussels (M. edulis were collected by hand by Division of Water Pollution Control (DWPC) personnel at low tide from a tidal creek near the town beach in Sandwich, MA. Immediately after collection, a subset of these animals were sent, on ice, to These the Lawrence Experiment Station (LES) for baseline tissue analysis. baseline samples consisted of the following: four replicates of 15 animals each were placed in labeled, sterile plastic bags for trace element tissue analysis (As, Cd, Cr, Cu, Hg, Ni, Pb, Zn) Twenty-five mussels were placed in a labeled sterile plastic bag for total and fecal coliform bacteria tissue analyses. Although not funded as part of this study, four replicates of 15 animals each were wrapped in aluminum foil and labeled for PCB and PAH analysis. The samples for the organics analysis were taken to the DWPC laboratory in Westborough and frozen for later analysis at the LES. The remaining mussels were transported in clean, plastic-lined coolers to the EPA Environmental Research Laboratory in Narragansett, RI. At this lab the animals were placed in flow-through seawater tables and left overnight. The following morning the mussels were sorted by size and 120 animals in the 5-6 cm range were selected for the growth study. These mussels were consecutively numbered from 1 to 120 using a dremel drill to etch the surface of the shell. The longest portion of the shell was measured to the nearest 0.1 mm using a Manostat (model 5921) caliper. The same individual performed all of the shell measurements with same caliper throughout the study. This technique was similar to that followed by personnel at the EPA, ERL, Narragansett, RI (William Nelson, EPA, Narragansett, RI, personal communication). )

.

Twenty-five mussels were placed in each cage, which was appropriately labeled by station and replicate. One cage per replicate contained 10 numbered animals among the twenty-five. Lids were secured with small plastic tie wraps. Cages were secured to the trawler float with heavy duty tie wraps for easy removal. The cages were left in the flow through seawater tables overnight. The following morning the mussel cages were transported in coolers to Clarks Cove, New Bedford. All stations were accessed by boat. At each station the cages from the previous deployment were retrieved and the new replicates were deployed. The replicates were spaced approximately 25 meters apart.

8

)

At each station water samples were collected with a van Dorn grab sampler below the surface and 1 meter from the bottom.

1

meter

These samples were collected to assess whether basic environmental conditions were similar at each site, as well as to make sure these conditions were suitable for mussel survival. Water samples to be analyzed for total solids, suspended chlorides, and turbidity were collected in clean polypropylene solids, Samples to be analyzed for total phosphorus, orthophosphate, containers. total Kjeldahl-nitrogen were collected in clean, acid rinsed bottles ammonia, and Samples for chlorophyll and acidified to pH 2.0 with 2 ml of 50 percent H 2 S04 All samples were a analysis were collected in clean polypropylene containers. tagged for identification and stored on ice in coolers for transport to the LES laboratory. .

Temperature, dissolved oxygen, salinity, and conductivity in the water column Data and field at each station were measured with a Hydrolab Surveyor II. observations pertaining to weather, sea conditions, and test animal and cage conditions after retrieval were recorded in a bound field notebook. On several occasions a YS1 Model #33 SCT meter was used to measure temperature, conductivity and salinity; dissolved oxygen was measured according to the Winkler technique. (Refer to Appendix A for details of meter accuracy, and sample treatment methods. On September 21, 1988 one sediment grab was collected at each station with a petite ponar grab dredge (Karlisco International Corp., El Cajon, Ca 92002). Prior to sampling the dredge was rinsed in seawater to remove any residual sediment. The inside of the dredge was then rinsed with reagent grade acetone, followed by a rinse with reagent grade hexane, followed by a final rinse with seawater. All chemical rinse waste was collected and transported back to the laboratory for proper disposal. The dredge contents were emptied into a plastic tray and subsamples of the sediment were scooped into separate specially cleaned 16 oz. glass, screw-top, wide-mouth jars prepared for metals and organics. Care was taken to prevent the collection of sediments in direct contact with the tray and/or sides of the dredge. All samples were identified with tags and stored on ice in coolers for delivery to LES. See Appendix A for details of sample bottle preparation. The sediments were analyzed at the LES for the following parameters: Trace elements (as total metals or metalloids): As, Cd, Cr, Cu, Hg, Ni, Pb, Zn; percent total volatile solids; PCBs and PAHs.

Appendix A presents the methodology employed at the LES for the analysis of the various water and sediment quality parameters. After collection the cages were left unopen and placed on ice in coolers for transport back to the DWPC laboratory in Westborough. The following day the cages were opened and the numbered animals were measured and individual shell length was recorded. The number of animals that were dead were noted along with the degree of fouling on the cages and on the animals themselves. Dead animals were identified by empty shells or by a strong odor of decay. Fifteen mussels were randomly selected from each replicate group and were placed in sterile plastic bags identified for trace element analysis. Fifteen animals were wrapped in aluminum foil and labeled and stored in the freezer (at 4°C) for later PCB and PAH analysis, and the remaining mussels (depending on how many were lost due to

mortality) were placed in labeled sterile plastic bags for total and fecal coliform bacteria analysis. The samples for trace element and bacteria analysis were then immediately transported on ice to the LES.

Methods of tissue preparation and analysis for trace element and bacteria in shellfish employed at the LES are outlined in Appendix A.

Inter-Laboratory Calibration An inter-laboratory calibration exercise was carried out between the Massachusetts Division of Marine Fisheries (DMF) and the Lawrence Experiment Station. The DMF proposed to analyze mussel tissue homogenate samples for trace elements: As, Cd, Cr, Cu, Hg, Ni, Pb, Zn at their Cat Cove Marine Laboratory A portion of the same tissue homogenate prepared by the LES for in Salem, MA. trace element analysis was frozen and stored at the laboratory for later delivery In September of 1988 the DMF notified DWPC that by DWPC personnel to the DMF. it would tissue homogenate samples from LES.

Appendix D contains a Ten samples were delivered to DMF on October 3, 1988. complete description of the DMF project plan and analytical procedures followed at the Cat Cove DMF laboratory. On March 28, 1989 the LES and DMF were each given 3 replicate frozen samples of Both a standard mussel tissue homogenate ("mega mussel") prepared by the EPA. laboratories were requested to analyze the tissue homogenate for the same suite of eight heavy metals and metalloids using the same methodologies employed during the caged mussel study.

Description of Study Site Clarks Cove is small, with a surface area of 5.18 km2 (2 mi 2 and an average water depth of 5 meters at MLW. The drainage area for the cove is comparatively large with the majority (approximately 8.1 km or 2,000 acres) lying within the City of New Bedford. The remaining watershed (approximately 2.4 km or 500 acres) is located within the boundaries of the Town of Dartmouth. Almost 94 percent of the total New Bedford drainage area is served by combined sewers (CDM, )

1983).

Along the shoreline of the cove there are nine combined sewer overflow (CSO) outfalls and seven storm drain pipes (Figure 3). Table 1 lists each CSO and its location and description. CDM estimated that 961 million gallons of storm and untreated wastewater were discharged to Clarks Cove in 1983. Forty-three percent (or 413 million gallons) of this was from CSO discharges, 6 percent (58 million gallons) was from dry weather discharges and 51 percent was from storm water runoff. They estimated that CSO discharges occur on an average of 75 times a year and they come from two major active outlets at the head of the cove (CSO #003 and #004).

10

FIGURE

3

BUZZARDS BAY CAGED MUSSEL PILOT BIOMONITORING STUDY Oct.1987-Sept.1988 Clarks Cove Combined Sewer Overflow Outfalls ICDM,1983) and Storm Drain Locations


TABLE

1

1 CLARKS COVE COMBINED SEWER OVERFLOWS

CSO OUTLET NUMBER

DIAMETER LOCATION

(Inches) 54"

003*

Cove Road and Padanaram Ave.

004

Hurrican Barrier Pumping Station

005*

Dudley Street and West Rodney French Blvd. (W.R.F. Blvd.)

18"

006

Lucas Street and W.R.F. Blvd.

24"

007

Capitol Street and W.R.F. Blvd.

24"

008

Calamet Street and W.R.F. Blvd.

18"

009

Aquidneck St. and W.R.F. Blvd.

18"

010

Bellevue St. and W.R.F. Blvd.

12"

0101

Hudson St. and W.R.F. Blvd.

18"

*

96" x 84"

Contaminated by dry weather sanitary flow from storm drains connected to the outfall, as observed by CDM (1983).

CDM Interim Summary Report on CSO Phase

12

I,

December 1983

The dry weather discharges occur as a result of structural or maintenance related problems of the existing sewer system. For example, the dry weather flow at CSO #004, estimated at over 0.16 MGD, is caused by a plugged dry weather connection. Historically, the highest coliform densities in Clarks Cove have been in the northern sector of the cove, presumably because of CSO dry weather discharges. The waterbody is classified as SA in accordance with the Massachusetts Water Quality Standards, but these standards are violated frequently. Clarks Cove receives heavy recreational use in the form of swimming, fishing, and boating. There are two public beaches and one private beach, and several boat ramps The cove is closed to commercial fishing and located around the cove. shellf ishing. Beach closings are reportedly rare.

13

RESULTS

Water Quality The physical and chemical water quality data collected during the study year are presented by station and date in Appendix B. Figures 4-6 illustrate the seasonal trend of temperature, dissolved oxygen, and salinity measured at one meter above the bottom at the three station locations. As shown, these parameters fluctuated similarly at each station throughout the survey year.

Salinity at all stations ranged between 27 - 32.2 parts per thousand during the year. Dissolved oxygen values ranged from a low of 5.0 mg/1 measured at Station A in July to high of 12.8 mg/1 measured at Station C in March. The July dissolved oxygen values exhibited the greatest between station differences (5.0 mg/1 at Station A and 7.2 mg/1 at Station C). Temperature, salinity and dissolved oxygen concentrations were within ranges necessary for mussel growth and survival at all of the stations.

Nutrient concentrations measured at the stations during the study were low to moderate and fell within ranges reported in the Buzzards Bay water quality surveys (MDWPC, 1985, 1986a), with the exception of Station B during March. This station exhibited elevated suspended solids and turbidity as well as high total Kjeldahl-nitrogen, total phosphorus and orthophosphate concentrations in the bottom water column sample. It is possible that the sediments were disturbed during sampling and this contaminated the sample. Suspended solids and turbidity were otherwise low and within expected ranges. These parameters followed similar trends between stations throughout the year. Sediment Quality

Table 2 presents the sediment trace element, PCB, PAH, and percent total volatile solids data for each station. All sediment samples were collected on September A rigorous assessment of the sediment quality was beyond the scope 21, 1988. of this study. Since the results cannot be normalized, and only one sediment grab per station was collected, an in-depth comparison and evaluation of sediment quality cannot be made from these data.

Station A sediments contained the highest concentrations of all trace elements, and organics measured, with the exception of nickel, which was slightly higher at Station C (6.5 mg/km versus 5.5 mg/km at Station A). Zinc and PCB 1254 concentrations were above category III dredge spoils criteria (MDWPC, 1983) at Station A. Arsenic was also elevated at this site. Station B and C sediments contained similar concentrations of most of the trace elements, and results were within ranges reported in the Buzzards Bay sediment survey (MDWPC, 1985-86). PCB 1254 concentration was higher at Station B (exceeded Category III criteria) than at Station C (Category II). PAH concentrations were relatively low at all of the stations, but the greatest number of compounds (and concentrations) were found at Station A and the least at Station C.

Percent total volatile solids were similar at all stations.

14

FIGURE 4

CLARKS COVE TEMPERATURE Stations A, B, and

C

May

March

Oct

Jan

1987

1988

Sept

Sampling Month

FIGURE 5

CLARKS COVE DISSOLVED OXYGEN Stations A, B, and

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TABLE

2

CLARKS COVE

SEDIMENT DATA TRACE ELEMENTS, POLYCHLORINATED BIPHENYLS, POLYCYCLIC AROMATIC HYDROCARBONS AND PERCENT TOTAL VOLATILE SOLIDS 1

September 21, 1988

CATAGORY III DREDGE SPOILS CRITERIA 2

STATIONS

PARAMETER

B

Trace Elements (mg/kg dry weight)

Arsenic Cadmium Chromium Copper Mercury Nickel Lead Zinc

2.4 6.5 41 60 0.335 5.5 90 500

0.170 2.5

2.0 1.0 29 24 0.105 6.5

33 85

25 90

>20 >10 >300 >400 >1.5 >100 >200 >400

>1.0

1.4

<1.0 23 60

PCB 1254 (pg/g)

2.3

1.3

0.91

Percent Total Volatile Solids

5.9

5.4

6.7

0.80 0.57 0.96 0.56 1.10 0.55 1.10

0.20

0.13 0.41 0.18 0.32

0.20 0.10 0.20

5.64

1.24

0.50

_

PAH (pg/g dry weight)

Benzo a anthracene Benzo a pyrene Bnezo(k) f luoranthene Chrysene Fluoranthene Phenanthrene Pyrene (

(

)

)

Total PAHs reported by LES

-

-

-

See Appendix A, Table A-5 for methods of analysis and limits of detection. DWPC,

1983

16

.

Cage Loss and Mussel Mortality

Percent mortality that occurred at each station during each deployment is presented in Table 3. The number of cages (replicates) lost during each exposure The percent mortality was calculated by period is also listed in this table. dividing the total number of dead animals found at a station by 200 (the total number of animals deployed at each station) and multiplying by 100. Mortality was usually very low, generally only 0-4 animals per station were lost. However, during the last exposure period of 7/13-9/21 mortality was very high (25-53 percent). An extreme degree of fouling by barnacles and algae was observed on the cages and animals themselves from this period. Also, several small starfish Cages collected from all other deployments were found in many of the cages. exhibited very little fouling and no starfish were observed inside them.

Other periods Four cages were lost during two of the deployment periods. replicate (C4) lost during the 1 or 2 cages. One experienced only a loss of first deployment period was recovered on 9/21/88 at the same site after almost Out of the original 50 animals, 27 survived from this one year of exposure. group. Shell Growth

Mean shell growth and standard deviation for animals at each station and for each deployment are shown in Figure 7. The average shell growth over a 60-day period of 120 mussels is highly variable as illustrated by the standard deviation bars (one S.D.) on the graph. This variability masks any statistical differences that may exist between Stations A, B, and C for any one deployment period. However, from the graph it appears that mean shell growth at these stations exhibit fairly similar trends during each period. The largest differences in shell growth are seen between exposure periods, although these are not statistically significant due to the large standard deviations. As expected, in general, the spring and early summer exposure groups show the largest increase in average shell growth, and the fall and winter periods produce the least amount of growth.

Tissue Bacteria Concentrations Tissue total and fecal coliform bacteria concentrations are presented in Table 4. Tissue samples from the last deployment period were not analyzed for bacteria concentration due to high mortality resulting in an insufficient number of live mussels available for the analysis. It was felt that the bacteria analysis was the most expendable of the parameters, because tissue bacteria data obtained from the last four deployments were erratic and did not supply any more useful information for monitoring long-term trends in bacteria contamination than could be obtained from direct water column sampling techniques (see discussion section)

Baseline tissue bacteria concentrations were generally much higher than tissue concentrations measured in animals after exposure, indicating that the Sandwich, MA site may not be appropriate for collecting "clean" mussels if bacterial contamination is a concern. A large number of birds were often observed near the area where the mussels were collected.

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In addition, bacteria concentrations between stations for each deployment did not exhibit a discernable pattern. It was expected that animals nearest the head of the cove would accumulate the highest bacteria concentrations. This was not On several occasions, Station C, the reference site located out in the case. In general, if total coliform Buzzards Bay, had the highest bacteria counts. was high (>1,000 colonies per 100 ml), fecal coliform was also elevated.

Tissue Trace Element Concentrations Figures 8 through 15 illustrate the results from the tissue analysis for trace Concentration is reported in mg/kg (wet weight) for mercury (Hg) elements. chromium (Cr) cadmium (Cd), arsenic (As), lead (Pb), nickel (Ni), copper (Cu), and zinc (Zn) ,

Each graph illustrates the tissue concentration of one trace element over all of the deployment periods. The bars represent the mean tissue concentration of the metalloid of all the replicates for each station, grouped by deployment period. One standard deviation is depicted on the graph to illustrate the Appendix B contains the tissue trace variability of the data about the mean. element concentration data as reported by the LES. Results from each deployment were examined separately. Comparison of contaminant concentration throughout the year is not possible since a new set of animals was used for each 60 day deployment period. Inter-exposure period comparisons of this nature would only have been possible if all of the animals had been deployed at the beginning of the study and subsampled throughout the year.

Statistical analysis using the nonparametric Kruskal-Wallis test (Zar, 1984) was performed on the tissue trace element concentration data. Nonparametric statistics were chosen because the variances of the groups of data being compared were not homogeneous. Under these conditions this nonparametric ANOVA test is more powerful than the one-way ANOVA (Zar, 1984). The Kruskal-Wallis statistic tested the null hypothesis that trace element concentration in tissue from the baseline station and Stations A, B, and C were the same. (H Q [metalloid] is the same at all stations.) :

Appendix C contains sample statistical calculations. of the results of the nonparametric ANOVA tests.

Table

5

presents a summary

A significant difference between mean trace element concentration was detected at the 95% confidence level for only 13 of the 35 groups of data tested. (During deployments four and five, detection limits of Cd, Cr and Pb were increased as a result of a change in laboratory procedure. As a consequence almost all values were reported as less than detection limits for these periods, thus limiting further analysis and comparison of these data sets.) Since the Kruskal-Wallis multiple comparison test does not indicate where the significant differences occur in the data set, a nonparametric Tukey-type multiple comparison test was applied to locate where the differences existed (Zar, 1984) for these 13 data sets. (See Appendix C for sample calculations.) Table 6 summarizes the results of these calculations. Due to high standard error values in several of the data sets only 8 of the 13 Tukey tests detected significant differences between the means.

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TABLE

5

SUMMARY OF KRUSKAL-WALLIS NONPARAMETRIC ANALYSIS OF VARIANCE

TRACE ELEMENT

DEPLOYMENT

DEPLOYMENT

1

DEPLOYMENT

2

3

DEPLOYMENT

DEPLOYMENT

4

5

Zinc

accept H

reject H

accept H

reject H

reject H

Mercury

accept HQ

accept HQ

accept H Q

accept H Q

accept H c

Nickel

accept HQ

accept H Q

reject H Q

accept H Q

accept H Q

Cadmium

accept H

accept HQ

accept H Q

can not analyze

can not analyze

Chromium

accept H

accept H Q

accept H Q

can not analyze

can not analyze

Arsenic

reject H

reject HQ

reject H

reject H

reject H

Lead

reject H

accept H

reject H

can not analyze

accept H

Copper

reject H

accept H Q

accept H Q

reject H Q

accept H Q

Hypothesis being tested: HQ

:

The mean trace element concentration of baseline = Station A = Station B = Station C

30

TABLE 6

SUMMARY OF TUKEY-TYPE NONPARAMETRIC MULTIPLE COMPARISON TEST

DATA SET Zinc deployment Zinc deployment Zinc deployment

5:

Nickel deployment

3:

No significant differences detectable due to large standard error

Arsenic deployment

1:

No significant differences error Baseline different (lower) Baseline different (lower) Baseline different (lower) Baseline different (higher)

Arsenic Arsenic Arsenic Arsenic

2:

4:

deployment 2: deployment 3: deployment 4: deployment 5:

Lead deployment

1:

Lead deployment

3:

Copper deployment

1:

Copper deployment

4:

RESULTS Baseline different (lower) than Sta. A,B,C; but A,B,C same Baseline different (lower) than Sta. B; but all others same Baseline different (higher) than Sta. C; but all others same

detectable due to large standard than than than than

Sta. Sta. Sta. Sta.

but but A, but A, but C, B,

all all all all

others others others others

same same same same

No significant differences detectable due to large standard error Baseline different (higher) than Sta. C, but all others same

No significant differences detectable due to large standard error No significant differences detectable due to large standard error

31

In every case, the significant differences in the means were due to the baseline mean tissue trace element concentration being different from one or more of the Usually, but not always, baseline concentrations other stations (A, B, or C) in these cases were lower. .

five of the exposure periods arsenic baseline tissue In four out of concentrations were significantly different from either Station A, B, or C. In three of these data sets arsenic was lowest in the baseline samples. However, since neither Station A, B, or C were consistently highest (or lowest) throughout the study, spatial patterns of arsenic distribution in this area are not evident.

Baseline concentration of zinc for three out of five exposure periods was However, as with arsenic, significantly different from Station A, B, or C. distribution of zinc at these stations cannot be spatial patterns of consistent detected nor can further speculation as to what may be causing these differences be made.

For lead, the baseline concentration was significantly higher than Station C for one exposure period.

Tissue concentration of cadmium, chromium, mercury, nickel and copper were not significantly different for any of the exposure periods. None of the test Stations (A, B, or C) exhibited significant differences in trace element tissue concentration indicating differences in bioaccumulation of these elements were not spatially significant for these stations. This suggests that trace element concentration available for uptake in the water column at these stations was not significantly different between Station A, B, or C. PCB Tissue Concentrations

The results of the PCB analysis of tissue from deployments 1, 3, 4, and 5 are illustrated in Figure 16. Mean values of the data normalized with percent lipids are shown on the graph. Appendix B lists the PCB tissue concentrations as reported by LES. Only arochlor 1254 was detected in any of the tissue samples. Percent lipid concentration for each sample is also reported in Appendix B. The lowest PCB concentrations were consistently tissue and the highest PCB concentrations were The next highest PCB concentrations were found concentrations of PCB were usually detected in

measured in the baseline mussel found in tissue from Station A. at Station B and relatively low tissue from Station C.

Interlaboratorv Calibration Exercise The results from the interlaboratory calibration exercise between the Department of Environmental Protection's laboratory (LES) and the Division of Marine Fisher ie's laboratory (DMF) at Cat Cove, Salem, MA are presented in Table 7.

32

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Values for cadmium and chromium are not comparable because the detection limits The DMF of the LES analyses were much higher than the DMF's detection limits. mercury concentrations of with the values reported "not quantifiable" falling This range is less than or near the between 0.006 mg/kg and 0.020 mg/kg. detection limit reported by the LES for mercury analysis. For copper, nickel, zinc and lead the values reported by the LES were approximately five times higher than that reported by the DMF for the same tissue homogenate samples.

On March 28, 1989 standard mussel tissue samples prepared by the EPA laboratory in Narragansett, RI were hand delivered to the Lawrence Experiment Station and the Division of Marine Fisheries laboratory at Cat Cove, Salem, MA. Results of each laboratory's analyses are presented in Table 8.

35

TABLE 8

INTERLABORATORY CALIBRATION RESULTS 1 U.S. EPA STANDARD "MEGA MUSSEL" TISSUE (mg/kg dry weight

)

METAL AGENCY

Cd

Cr

Cu

Ni

Pb

2.08

2.15

12.8

6.84

9.11

135

Lawrence Experiment Station

1.9

1.9

11.7

6.2

8.3

119

U.S. EPA range of values'

1.992.18

1.912.36

12.213.8

6.377.24

2.17

1.98

11.7

U.S. EPA, Narragansett, RI

Zn

(Average tissue metals concentration)

Division of Marine Fisheries

LES obtained dry weight of sample by drying homogenate for and weighing entire sample. J

3

2

7.9410.25

126-

8.06

90

days at 90 °C

LES results are reported as an average of 2AA analyses (except for Pb only enough sample for one analysis). U.S. EPA analyzed 50 samples to obtain range and average tissue metals concentration of the standard homogenate.

Division of Marine Fisheries results were converted to dry weight by multiplying wet weight values by 6.83 (EPA's reported wet/dry weight ratio for the "mega mussel" homogenate).

36

142

DISCUSSION

Study Design Studies that involve comparisons of selected variables over space and time ideally require that all environmental conditions that may affect test results be similar either through controlled laboratory conditions or, in field studies, However, too much control placed on the as a function of study design. artificial situation which may obscure create an design may experimental interpretation of the relationship of the data to actual field conditions. For this study it was important that the stations selected exhibit very similar The three stations chosen were oriented measurable environmental conditions. on a north-south transect from the head of shallow Clarks Cove to open water in Buzzards Bay; consequently depth was not the same at each location. (5 meters Despite depth differences however, at Station A and B vs 9 meters at Station C. were essentially the same at each dissolved oxygen and salinity temperature, station supporting the assumption that all of the animals were most likely exposed to similar environmental conditions during each deployment period. )

Growth (as measured by average shell length increases over the exposure period) and mortality were not significantly different between stations which also indicates that the environmental conditions necessary for mussel growth and survivorship at each site were the same. If growth differences between sites were evident in this study, then differences in tissue trace element bioaccumulation between sites (if present) would be more

difficult to interpret and could not necessarily be attributed available contaminant concentrations in the water column.

solely

to

Enseco, Inc. (1990) reported that mussels deployed near sewage treatment outfalls in Boston Harbor that survived appeared to be generally healthier than reference site organisms. Based on these findings, assumptions that more polluted sites would negatively affect the health (and growth) of test animals cannot be made.

Although not performed in this study other methods of growth or condition assessment may be more effective than simple shell length measurements. A practical method of determining a body condition index should be investigated and, if at all possible, applied in future caged mussel studies of this kind.

Mortality was usually very low except for the last exposure period where predation by starfish was suspected to have caused the 25-53 percent mortality observed in the cages. Although it is not known if starfish predation on bivalves occurs seasonally in Buzzards Bay it may be wise to avoid deploying the mussels in cages during this time of year in this particular area. For all but the last deployment, the low mussel mortality assured sufficient numbers of animals for tissue analysis. In addition, similar mussel growth, mortality and environmental conditions found at each station reduces sources of variation that may influence spatial differences in contaminant uptake by the mussels.

37

Coliform Contamination The use of caged mussels to monitor coliform contamination over time and space was ineffective. Since the animals clear their gut in approximately 24-48 hours, any assumptions regarding temporal changes in bacteria concentration in the water Furthermore, the potential for column are limited to one day time periods. encountering variability within the stations is high due to the fact that the animals are filter feeders, and may each be filtering different volumes of water over a 24-hour periods and thus ingesting highly variable amounts of bacteria For this reason, monitoring of coliform over this relatively short time. bacteria to detect long-term changes in water column bacteria densities should not be performed via tissue concentrations.

Monitoring whole mussel tissue bacteria concentration is potentially a valid technique for making spatial comparisons of bacteria concentrations in the water column at discrete time periods. However, the method is much more labor intensive, results are highly variable, and it offers only slight advantage (i.e, from a temporally non-integrated grab sample of water versus a 24-48 hour time integrated tissue sample) over simple, direct water column bacteria sampling methods. Trace Element Concentration As is evident from the data, tissue trace element concentration was extremely variable, not only statistically between replicates at each station, but spatially and temporally as well. Due to variation within the data set significant differences in trace element concentrations, if they existed in the water column at any of these stations over time, were not usually detectable. The magnitude of trace element bioaccumulation in the mussel tissue was small in comparison to this variability. It is important to examine the major factors that may influence the variability of the data and its resulting usefulness.

The often large variances of the station replicates as well as the differences in average tissue trace element concentration between Stations A, B and C (spatial differences) and between baseline mussel tissue and Stations A, B, and C (temporal differences) may be the result of any one, or a combination of the following factors: 1) natural seasonal variability; 2) data bias or errors resulting from field study design and implementation; 3) data bias or errors resulting from laboratory procedures; 4) actual temporal or spatial differences in water column trace element concentrations.

Natural seasonal variation can account for as much as 15-60 percent of the variability in observed values (Capuzzo, et. al., 1987). Seasonal variation may be a result of the physiological state of the animals, environmental conditions, and metal speciation and bioavilability (Capuzzo, et. al., 1987). Seasonal variability would not influence the between station (spatial) differences of the data because comparisons of these results were made between mussel tissue from the same exposure period. As previously discussed, results indicated that these mussels were experiencing similar seasonal environmental and physiological conditions as measured by temperature, dissolved oxygen, salinity, chlorophyll a concentrations, and shell growth at each site. Tissue trace element concentrations were not compared at each station over several exposure periods. With this study design, comparisons of this type would

38

be weak because discrete groups of animals were set out and measured each exposure time, rather than subsampled periodically from a large group that had been exposed for the entire study year. However, seasonal variability may have caused differences between baseline tissue concentrations and Stations A, B, and C since baseline animals were collected in Sandwich at the beginning of the exposure period approximately 60 days earlier than the animals they were compared to from Clarks Cove.

Percent lipids were not measured in the tissues homogenized for trace element analyses; except for growth, no other parameters were measured to assess the physiological condition of the mussels. Percent lipids were measured in tissue Although not homogenate prepared for organics analysis (see Appendix B) assessed during this study, spawning condition of the animals is known to be Lipid directly related to whole animal percent lipid concentration. concentration increases as animals prepare to spawn and drops sharply after Spawning reportedly leads to loss in tissue weight, increase in spawning. Prior to spawning lipid-rich percent water and decline in condition indices. gametes may contain higher concentrations of lipophilic organic contaminants and lower concentrations of heavy metals than somatic tissues. After spawning a drop in organic concentrations and an increase in metal concentrations may result Therefore, to greatly enhance tissue data (Robinson and Ryan, 1988). interpretation future caged mussel studies should include an assessment of the spawning condition of the animals. This should be made at the time of deployment, when baseline trace element tissue concentrations are measured, and when the animals are retrieved after the exposure period. Inferences about adverse impacts of toxic trace elements on the health of the mussel cannot be made, although this factor may have also been responsible for some variability of the data. Animals showed an average increase in shell length for each exposure period. Average shell increases were the largest during the third and fourth deployments (March 16 - May 11 and May 11 - June 16, respectively) No correlation between growth and trace element concentrations can be made. The goals of this study did not include an attempt to relate contamination concentration with indications of stress in the organism. .

.

As previously discussed, the field study design attempted to equalize as many between station environmental variables as possible. The study design may benefit from including at least one more replicate at each station since the variances between the four replicates were often high. In addition, it has been suggested that not all of the animals receive equal exposure time bunched-up in the square cages. Flat cages that spread the animals into one-layer would allow all to have more of a chance to filter equal amounts of water. Cages of this design were not available for this study. To reduce the likelihood of this type of bias animals were selected randomly from the bunches in the cages when preparing the sample bags for the laboratory. Other studies performed with square cages did not report evidence of this type of bias (Robinson and Ryan, 1986, 1988 and Nelson, personal communication).

Besides the systematic or random variability introduced via seasonality and field study design, data variability introduced through laboratory procedures must be considered an important factor when interpreting the results. The Lawrence Experiment Station analyzes samples in "batches." QA/QC tests are performed on

39

The QA/QC results during this study variation between stations and/or replicate suggesting that were acceptable, samples was due to other factors (i.e., the effects of seasonality, or differences in contaminant concentrations). a percentage of samples from each batch.

Determination of dry weight concentrations of the trace element was not requested However, water content is extremely variable in these as part of this study. animals, not only seasonally but individually, and will definitely affect the calculation of the results. Ideally, dry weight should be determined separately for each sample homogenate prepared, rather than using an average dry weight of Robinson and Ryan (1988) state that in mussel tissue to normalize the data. transplant studies it is impossible to determine whether metal body burdens actually changed as a result of exposure if changes in tissue weights were not monitored. They report that changes in mussel tissue weight can be assessed by Future measurements of tissue dry weight, condition index and gonadal index. determination of tissue dry weight should include a tissue biomonitoring studies to reduce data variability. Possible sources of data variability were discussed with LES personnel and they Some included procedures in sample preparation and analytical methodologies. of these sources can be minimized with the use of a more efficient method of tissue homogenation and/or via procedural modifications such as determining the dry weight of samples and using consistent sample sizes for analysis throughout the study.

Results of tissue metals concentration from this study and ranges of values reported for several other metals bioaccumulation studies are compared in Table 9. Arsenic concentration was not measured in the other studies listed here, so Mercury, cadmium, and chromium it is not included in this comparison. Mercury concentrations fall within the ranges reported by other researchers. concentration never approached the US Food and Drug Administration limit of 1 /jg/g wet weight. Cadmium and chromium were also very low, often below the detection limits of the analyses, and concentrations never fluctuated much from site to site, nor did they vary over exposure times. From this study, it appears that mercury, cadmium and chromium either require a longer exposure period to bioaccumulate in the mussel or there were low concentrations of bioavailable metal in the water column at these sites, de Kock and van het Groenewoud (1985) report that cadmium accumulation is a slow process requiring about 150 days to reach equilibrium values. These researchers were also unable to demonstrate differences in mercury concentration from several sites in 60 day transplant studies. Robinson and Ryan (1988) state that transplanting clean mussels to polluted sites to assess seawater contaminant levels is only successful when metals concentrations are high enough to result in appreciable bioaccumulation.

Maximum concentrations of lead, copper, nickel, and zinc greatly exceeded ranges reported from other studies (see Table 9). Station A tissue most often contained the highest metals concentrations; however as previously discussed, between station differences of tissue concentrations of these metals could not be detected due to large within station variances. In general, seasonal peaks in Cu, Ni, Pb and Zn tissue concentration occurred more in the late spring and early summer (also the period when the greatest shell length increases were measured). Possible reasons for these extremely high values of Zn, Cu, Ni, and Pb includes laboratory sources of variability discussed previously as well as the natural

40

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Lawrence Experiment Station (LES) v* D.v.s.on

of Marine Fisheries as a Percent of EPA

(DMR

Values

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42

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deviat on.

variability of actual metals concentration and varying rates of bioac cumulation and regulation of each metal by Mvtilus edulis during different times of the year.

No range of arsenic tissue concentrations were available for comparison. Ranges of concentration for this element were not included in Table 9 for this reason. For arsenic the within station variances were usually lower than for other Arsenic concentrations from July and September samples were metals. significantly higher at all stations than at any other time of year.

Interlaboratorv Calibration Exercise The interlaboratory calibration exercise with the DMF yielded differences in tissue wet weight concentrations of copper, nickel, and zinc from aliquot s of the same sample homogenate. Values of cadmium, chromium, and lead reported by DMF were lower than the detection limits established by LES in their analysis DMF obtained unquantif iable therefore these metals data were not comparable. concentrations of mercury (0.006 ppm
attempt to assess whether these differences were actually due to differences in laboratory techniques and not in the samples themselves, each laboratory was requested to analyze standard EPA mega mussel tissue homogenate. Results show concentrations obtained by the LES are within 9-12 percent of the values reported by EPA (Figure 17). It is interesting to note that the LES values are all slightly less than EPA average concentrations. The LES results for Pb and Cr fall within the range of values reported by EPA. Cd, Cu, Ni, and Zn values were only slightly less than the minimum EPA values. DMF results for Cd, Cr, and Pb fall within EPA's reported ranges. As illustrated in Figure 17, values for Cd, Cr, Cu, and Pb reported by DMF are within 4-11 percent of EPA averages. DMF did not report results for nickel. Zinc concentration obtained by DMF was comparatively low however, differing by more than 30 percent from the EPA average. In

an

Considering that the EPA averages were based on a sample size of 50 and the LES and DMF values were derived from an average of two (or less) analysis, results for these trace element analyses appear to be in very good agreement among the laboratories. Both laboratories tended to have a bias toward lower values as compared to EPA results but the reasons for this trend are unknown.

43

Based on the results of the "mega mussel" interlaboratory calibration exercise, no evidence for why the results of the study mussel tissue interlaboratory analyses were so different between LES and DMF can be found.

Tissue Concentrations of PCBs and PAHs This project only required that a portion of the animals from each exposure However, since the period be archived (frozen) for future organics analysis. organics laboratory at the LES was able to perform the analysis on many of the archived samples during the study period, the results are presented and briefly discussed as part of this report. No PAHs were detected in the tissue samples from either Clarks Cove or Sandwich, MA. In contrast, Capuzzo, et. al. (1987) report mussel tissue collected from a variety of sites in New England, including Cape Cod, contained detectable levels of PAHs. Eisler (1987) however, found in general that PAHs show little tendency to biomagnify in food chains. He attributed this to the fact that most Specific reasons for PAHs not being detected in PAHs are rapidly metabolized. this study cannot be offered. An interlaboratory comparison between the LES and EPA, Narragansett or Woods Hole Oceanography Institute organics laboratories may provide some insight as to what is happening here.

PCB tissue concentrations were normalized by the percent lipid concentration of the sample to account for differences in PCB concentration created by differences in lipid content of the tissue. As seen from Figure 16 the results show a consistent pattern of higher PCB concentrations in the tissues from Station A to decreasing amounts in Station B and even lesser amounts in Station C. Not between only are spatial differences evident, but differences can be seen baseline and test site concentrations for each deployment period. From this consistent pattern it appears that 60 day exposure periods allow sufficient time for bioaccumulation of measurable amounts of PCBs in mussels. EPA recommends at least 30 days (U.S. EPA, 1983), although differences in PCB concentration of test animals have been detected after just 2 weeks of exposure in New Bedford Harbor (W. Nelson, personnel communication)

Based on the well documented PCB contamination in New Bedford Harbor it is not surprising that PCB concentrations at Station B as well as Station A, were relatively high. The area that encompasses both stations has been closed to bottom fishing and lobstering by the Department of Public Health due to PCB contamination. None of the tissues from this study contained PCBs in excess of the FDA action level of 2.0 \iqlq. Concentrations ranged from <0.04-1.2 p/g/g. This range falls within that observed in US Mussel Watch data from Cape Cod and Buzzards Bay. In New Bedford Harbor the range of PCB tissue concentration from Mussel Watch data was much higher (3.08-6.86 pg/g) (Capuzzo, et. al., 1987). In this study the use of Mytilus edulis as a sentinel organism to monitor PCB contamination in the water column appears more successful than for monitoring metals contamination. The standard deviations of the station replicates were low and concentration averages followed an expected pattern for every deployment. Sediment PCBs also followed a similar, relative concentration gradient. Results

appear to be more straightforward to interpret both spatially and temporally. Also by normalizing with percent lipids much of the variability that may have been introduced as a result of differences in reproductive condition of the animals was eliminated.

44

.

SUMMARY The use of caged mussels for coastal biomonitoring proved to be a very feasible field technique from the standpoint of available resources at the Technical Questions that remain should be addressed through Services Branch of DWPC. increased communication with the analytical laboratory, continued interlaboratory Based on the calibration exercises, and modification of the study design. results and suggestions from other researchers, several modifications of the 1) trace elements that study design and analytical procedure are recommended: from be eliminated the study (Cd, Cr, and exhibited low bioconcentration should Hg); 2) tissue dry weight should be determined for each sample homogenate; 3) the sample should be thoroughly homogenized; 4) interlaboratory calibrations should continue with sample tissues from the study sites as well as with a standard tissue homogenate (EPA mega mussel); 5) increase focus on using this technique to monitor PCB contamination; 6) examine the effect of longer exposure periods by subsampling from a large group of transplanted mussels over a one year period; and 7) the method should not be used to monitor coliform bacteria contamination In most of Buzzards Bay, metals contamination is most likely not high enough to significant amounts. If definitive bioaccumulate to statistically bioaccumulation was not measured at Clarks Cove, other less impacted areas would be less expected to show significant bioaccumulation of tissue in trace element concentrations. From this study it is evident that actual differences, either spatial or temporal would have to be very large to be significant. However, this study as well as others indicate that temporal and spatial characterization of changes in PCB contamination are possible using caged mussels. Serious consideration should be given to using this technique as part of a LONG-TERM monitoring program in Buzzards Bay, especially in the New Bedford area. It is important that biomonitoring studies such as this continue to be developed

and performed by agencies responsible for water quality monitoring. Of the three basic methods used to assess pollutants in the coastal environment; water sampling, sediment sampling, and sampling of biota, the later has received the least attention by the Massachusetts Division of Water Pollution Control. The bioavailability of contaminants however, should be a major concern, not only because it can provide a means of determining time-integrated pollutant concentrations but because of the long-term implications to human health, and more important, the overall health of the ecosystem. Although water pollution standards today are based on measurements of water and sediment, a contaminant can only be considered a threat to the environment if it can be taken up by the biota.

45

BIBLIOGRAPHY

American Public Health Association Q { »at er „, ,..^.>„. _

1985

ys; :^;

^

" " SU™ ar*

R **° rt ° n

..i^n


Camp Dresser and McKee. 1983. city of New Bedford combined Sewer Overflow. Phase I. TJ*J>° Capuzzo,

J.M.,

A.

218 » Street.

Filling in

McElroy,

and

G.

Wallace

^

^ „

t "I t '



1

1987

»-

v,

..,^£££H£ "^Bpp. ** """"" R^°"< """^ "s^*

^^^^^x^SST'^rS!

Commonwealth of Massachusetts, Division of Water Pn ,i Bay Mater Quality survey Da

-

-«*.

y

1

9

Procedures -

W " er Biomomtoring 011

""

1'

<-

j^^s^rs-

——

ourvey uata, Part A.

^^^^e^r^:^ ^ 0X58X.

f

Westborough, MA

Program. "

85

01581

-as.

Westborough,

MA

de Kock, w. Chr. and H. van het Groenewoud ,,. loft* v, Modelling and Elimination Dynamics of s™» Bioaocumulation t'- -." P° Uutants cd Based on "in Situ-^e"atfonsTi ' ». FCB, HCB, th 1 ,or society, Heport

Z

,,

^^ XOS^if^^f ^X ""^ „^

<

,'

Report

.

85(1.11).

June 1990.

" eView *

U S '

Fish Wildlife Serv ice

'

81pp.

Marblehead, MA

M ° nit ° rin 10945

9

*«*"». "89

Rim

Deployment.

^

Farrington, J.W., A.C. Davis, B.W. Trior, D y »>,., "Mussel watch - Measurement, p* 11 of f" indicator of Coastal Environmental v Bivalves As One tal Quality » He „ a "*"_;teBtaasiisa to Mnnii-n.-i ,,,, Aouati. ssaaa ASTM ftmerican s ° ci <*y «« Testing and Materials, Phila., ' ppl25-X 3V

Cb^aXpo^tanL

^l^',

Farrington, j.w. and G.c. Medeiros ISA* Analysis for Petroleum Hvdt^V k Proceedings f erence On^reve'nl coastal Research Center woods! Lie

£^£

* Evaluation of Some Methods of ° r9anisms Gained in C°° tr ° 1 f Poll ^°»° ° ,

""^ hT

46

-

U

-

oxsix

,

BIBLIOGRAPHY (CONTINUED)

Galloway, W.B., J.L. Lake, D.K. Phelps, P.F. Rogerson, V.T. Bowen, J.W. Farrington, E.D. Goldberg, J.L. Laseter, G.C. Lawler, J.H. Martin, and R.W. Intercomparison of Trace Level The Mussel Watch: 1983. Risebrough. Toxicology and Chemistry Environmental Vol. Constituent Determinations. 2. pp395-410. .

1986. The Mussel Watch Concept. Goldberg, E.D. Vol. 7 (1986) 91-103. Assessment

Environmental Monitoring and

.

1987. "Innovative Designs for Perry, J. A., D.J. Schaeffer and E.E. Herricks. Water Quality Monitoring: Are We Asking the Questions Before the Data Are ASTM STP 940 Collected?", New Approaches to Monitoring Aguatic Ecosystems T.P. Boyle, Ed. American Society for Testing and Materials. Phila. pp28.

39.

Phelps, D.K., W.B. Galloway, B.H. Reynolds, W.G. Nelson, G. Hoffman, J. Lake, 1982. Evaluation C. Barsycz, F.P. Thurberg, J. Graikowski and K. Jenkins. Report: Use of Caged Mussel Transplants for Monitoring Fate and Effects of Ocean Disposal in the New York Bight Apex. US EPA Environmental Research Laboratory, Narragansett, ERIN No. 586. 35pp.

Robinson W.E. and D.K. Ryan. 1986. Bioaccumulation of Metals, Polychlorinated Biphenyls, Polyaromatic Hydrocarbons and Chlorinated Pesticides in the A Mussel, Mvtilus edulis L., Transplanted to Salem Sound, Massachusetts. final report submitted to Camp Dresser and McKee, Inc. 20 October 1986. Robinson W.E. and D.K. Ryan. 1988. Bioaccumulation of Metal and Organic Contaminants in the Mussel, Mytilus edulis Transplanted to Boston Harbor, Massachusetts. In Project Report submitted to Camp Dresser and McKee, Inc. by the Edgarton Research Laboratory, New England Aquarium, 15 February 1988. .

205pp.

Segar, D.A., D.J.H. Phillips., and E. Stamman. 1987. "Strategies for Long-Term Pollution Monitoring of the Coastal Oceans," New Approaches to Monitoring Aguatic Ecosystems . ASTM STP 940, T.P. Boyle, Ed. American Society for Testing and Materials, Phila. ppl2-27.

Signell, R.P. 1987. Tide and Wind-Forced Currents in Buzzards Bay, MA Hole Oceanographic Institute, Woods Hole, MA WHOI-87-15. 86pp. Tripp,

.

Woods

and J.W. Farrington. 1984. Using Sentinel Organisms to Monitor Chemical Changes in the Coastal Zone: Progress or Paralysis. Proceedings of the Ninth Annual Conference of the Coastal Society. Oct. 14-17, Atlantic City, NJ. B.W.

U.S. Environmental Protection Agency.

Water and Wastes.

1983.

Methods for Chemical Analysis of

EPA-600/4-79-020.

U.S. Environmental Protection Agency. 1983. Methods for Use of Caged Mussels for in situ Biomonitoring of Marine Sewage Discharges. EPA-600/4-83-000.

47

BIBLIOGRAPHY (CONTINUED)

U.S. Food and Drug Administration.

Analytical Manual. Zar,

January.

Food and Drug Procedure. Washington, D.C. 1988.

Biostatistical Analysis 1984. Englewood Cliffs, NJ 01632. 718pp. J.H.

48

,

2nd Edition.

Pesticide

Prentice-Hall Inc.,

APPENDIX A

FIELD METHODOLOGY

AND

LAWRENCE EXPERIMENT STATION LABORATORY METHODOLOGY

49

TABLE A-l

COMMON SAMPLE TREATMENT METHODS

PARAMETER

Dissolved Oxygen

SAMPLE VOLUME 300 ml (2)

Temperature

SAMPLE CONTAINER 1

IMMEDIATE SHIPBOARD PROCESSING & STORAGE

G (1)

MnS04 ; KI: no sunlight/ or (4) "in situ."

"

In situ recorded to

(1)

nearest 0.1°C/F or (3), (4),

(5)

Specific Conductance

1

1

(2)

P/G (1)

"In situ" reading/or cool 4°C (3), (4)

Total Solids

1

1

(2)

P/G (1)

Cool 4°C

Suspended Solids

1

1

(2)

P/G (1)

Cool 4°C

Chloride

1

1

(2)

P/G (1)

Cool 4°C

Total Kjeldahl-Nitrogen

500 ml (2)

G (1)

H 2 S04/ pH <2.0, cool 4°C

Ammonia-Nitrogen

500 ml (2)

G (1)

H 2 S04 , pH <2.0, cool 4°C

Total Phosphorus

500 ml (2)

G (1)

H 2 S04

Orthophosphate

500 ml (2)

G (1)

H 2 S04/ pH <2.0, cool 4°C

Turbidity

1

G (1)

Cool 4°C

Chlorophyll a/ Phytoplankton

200 ml

P/G (1)

Cool 4°C (5)

1

(2)

,

pH <2.0, cool 4°C

G - Glass P/G - Polypropylene or glass (1)

Required containers, preservation techniques, and holding time, per Table II 40 CFR Part 136.

(2)

Massachusetts Division of Water Pollution Control, Technical Services Branch, Engineering Section, Standard Operating Procedures.

(3)

Yellow Springs Instrument, Model 33-S-C-T meter and probe. Instrument Co., Inc., Yellow Springs, Ohio 45387.

(4)

Hydrolab Surveyor II, Model SVR2-SU sonde unit, Model SVR2-DV Digital read out. Hydrolab Corp., P.O. Box 50116, Austin TX 78763.

(5)

Massachusetts Division of Water Pollution Control, Technical Services Branch, Biomonitoring Program 1988, Standard Operating Procedures.

50

Yellow Springs

TABLE A-2

PARAMETER AND COLLECTION METHODS EMPLOYED AT SEDIMENT STATIONS

SAMPLE VOLUME

IMMEDIATE FIELD PROCESSING & STORAGE

(Liters)

SAMPLE CONTAINER

PCB 1016/1242 Sediment

2(25-100 g)

G/Aluminum Foil Septum

Cool to 4°C

PCB 1248 Sediment

2(25-100 g)

G/Aluminum Foil Septum

Cool to 4°C

PCB 1254 Sediment

2(25-100 g)

G/Aluminum Foil Septum

Cool to 4°C

PCB 1260 Sediment

2(25-100 g)

G/Aluminum Foil Septum

Cool to 4°C

PAHs Sediment

2(25-100 g)

G/Aluminum Foil Septum

Cool to 4°C

Metals Sediment

25-100 g

G/Teflon Septum or Plastic Wrap Septum

Cool to 4°C

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7

TABLE A-

METHOD FOR CHLOROPHYLL a ANALYSIS (MDWPC, 1988)

3.7.1

Chlorophyll is a pigment found in plants that allows the DEFINITION organism to use radiant energy for converting carbon dioxide into organic Several types of compounds in a process called photosynthesis. pigments used are to characterize chlorophylls exist and these and other algae. One type, chlorophyll a, is measured for it is found in all algae. A knowledge of chlorophyll a concentrations provides qualitative and quantitative estimations of phytoplanktonic and periphytic biomasses for comparative assessments of geographical, spatial and temporal variations.

3.7.2

EQUIPMENT NEEDS

;

1.

Fluorometer - either Turner 111 or the Turner Design 10-005-R field fluorometer is used. They must be equipped with blue lamp F4T5.

Corning filter -5-60-excitation Corning filter - 2-64-emission Photomultiplier 2.

Tissue grinder and tube - Thomas Tissue Grinder

3.

Side arm vacuum flask and pump

4.

Millipore filter holder

5.

Glass fiber filter:

6.

Centrifuge (Fisher Scientific Safety Centrifuge)

7.

15 ml graduated conical end centrifuge tubes with rubber stoppers

8.

90% aqueous acetone

9.

IN

Reeve angel, grade 934H, 2.1 cm

HCL

10.

Saturated magnesium solution in distilled water

11.

Test tube racks

12.

Borosilicate cuvettes - Turner 111 - 3" cuvettes Turner Design - 8" cuvettes

13.

Aluminum foil

14.

Test tube brushes - conical end

15.

Parafilm

59

TABLE A-7 (CONTINUED)

3.7.3

LOG- IN PROCEDURE The As samples are received they are logged in and assigned a number. samples can be frozen for further analysis, or the filter ground up for analysis the following day.

3.7.4

SAMPLE PREPARATION Samples are generally processed as soon as they come into the laboratory, unless there are extenuating circumstances, such as faulty equipment and/or time constraints. Samples not to be analyzed within 24 hours are frozen for future analysis. The procedure for freezing samples follows: 1.

Label a 2-inch Whatman petri dish with the sample number using an indelible pen.

2.

Using tweezers, take a 2.1 cm Reeve Angel, grade 934AH, glass fiber Do filter and place it on the Millipore filtering flask screen. not touch the filter. Attach the glass tube to the filter flask with the metal clamp.

3.

Shake the sample well.

4.

Measure out 50 mis of sample or less. If an amount other than 50 mis is used it should be recorded in the chlorophyll data book.

5.

Pour the measured sample into the filter tube and turn on the vacuum. The sample should pass quickly through the glass fiber filter; therefore more of the sample should be added. If the sample is not filtering through - either because too much sediment is present or the algal concentration is too high - then less than 50 mis can be filtered. A notation is made in the chlorophyll data book which lists the amount that was filtered.

6.

Unclamp the filter holder and with tweezers transfer the filter to the previously marked petri dish.

7.

Cover the petri dish and wrap it in aluminum foil to keep out the light. The petri dish with the glass fiber filter is then stored in the freezer.

8.

Return the sample bottle to the refrigerator if algal counts or identifications are requested.

9.

Rinse the graduated cylinder and filter holder in distilled water.

60

.

TABLE A-7 (CONTINUED)

3.7.5

ANALYTICAL PROCEDURE 1.

Follow steps 2-6 under "Sample Preparation."

2.

Filter 50 ml (or less if necessary) of sample through a glass fiber filter under vacuum.

3.

Push the filter to the bottom of tissue grinding tube.

4.

Add about

5.

Grind contents for

6.

The contents of the grinding tube are carefully washed into a 15 ml graduated centrifuge tube.

7.

Bring the sample volume to 10 ml with 90% acetone.

8.

Test tubes are wrapped with refrigerator for 24 hours.

9.

Test tubes are taken out of the refrigerator and put centrifuge.

3

ml of 90% acetone and 0.2 ml of the MgC0 3 solution. 3

minutes.

aluminum

foil

and

stored

in

the

into the

10.

Test tubes are then centrifuged for 20 minutes and the supernatant decanted immediately into stoppered test tubes.

11.

Tubes are allowed to come to room temperature. The temperature is recorded and the samples are poured into a cuvette (3" for Turner 111 and 8" for Turner Design)

12.

The Turner 111 requires a warm-up period of at least one-half hour, while the Turner Design 10-005-R does not require a warm-up period.

13.

With Turner 111, use a blank of 90% aqueous solution of acetone to zero the instrument. Open the front door of the fluorometer and put in the cuvette containing the 90% acetone and close the door. Press the start switch. The dial should move back to 0; adjustments can be made with the calibration knob. This process should be repeated as often as necessary, i.e., if the blank is not staying on zero; but no alteration should be made until a series of samples is completed.

14.

The Turner Design must also be zeroed to an acetone blank. The sample holder is located at the top of the Turner Design field fluorometer and should be recovered with the black cap after the sample is put in it.

61

TABLE A- 7 (CONTINUED)

15.

Readings for both the Turner 111 and the Turner Design should be within 20-80% of the scale. This can be achieved by either reducing or increasing the opening to the lamp by moving the knob on the right front of the Turner 111 fluorometer. The sensitivity levels The sensitivity level must be recorded are lx, 3x, lOx, and 30x. in the chlorophyll data book in addition to whether the high After the first reading, 2 intensity or regular door was used. A piece of parafilm is drops of 2N HC1 is added to the cuvette. used to cover the cuvette which is then inverted four times to mix The sample is re-read and the new value the sample thoroughly. recorded.

16.

The procedure for the Turner Design field fluorometer is basically the same as for the Turner 111. The sample is put into the cuvette holder and the manual switch used to go from one sensitivity level to the next without opening the door. A reading of between 20-80% is still required for accuracy. Readings are taken before and after acid is added to the sample. The level of sensitivity (lx, 3x, 6x, lOx, 31. 6x) must be recorded in the chlorophyll data book, as well as whether the levels were set at 1 or 100.

Calculation of Chlorophyll Concentrations Chlorophyll formulas:

concentrations

chlorophyll

(fig/1)

= Fs

are

determined

rs

by

using

the

following

(Rb-RA)

rs-1

pheophytin

(/ug/l)

= Fs

rs

(rsRa-Rb)

rs-1 where, Fs = conversion factor for sensitivity level "s" rs = before and after acidification ratio of sensitivity level "s" Rb = fluorometer reading before acidification Ra = fluorometer reading after acidification

A computer program is used to calculate the chlorophyll concentrations for samples run on the Turner Design fluorometer. This program requires the investigator to type in the sensitivity level and the difference between the before and after acidification values. During the summer of 1986 personnel of the Technical Services Branch (TSB) conducted a laboratory experiment with a Turner Design Fluorometer in order to determine the effect of pheophytin b on freshwater chlorophyll a readings. Pheophytin b is the degradation product of chlorophyll b which is the primary pigment of green algae. The Turner

62

TABLE A- 7 (CONTINUED)

Design instrument measures the fluorescence of chlorophyll a as well as Chlorophyll b is not read at the same that of pheophytin a and b. frequency as chlorophyll a. The emission filter used at the TSB (Corning C/S 2-64) partially rejects pheophytin b (See: "References" - Turner Designs, 1981). It was found and recorded in various unpublished memoranda (See "References") that unless a sample had elevated counts of green algae the readings obtained prior to acidification and 90 seconds thereafter would give a reliable estimate of the concentration In cases with elevated counts of of chlorophyll a in an algal sample. green algae an annotation should be made alongside the chlorophyll a concentration stating that the concentration may reflect the presence of chlorophyll b and is probably lower than as recorded. As a result investigation, the TSB now of this present chlorophyll data as chlorophyll a in mg/m .

3.7.6

INSTRUMENT CALIBRATION

Fluorometers are calibrated using chlorophyll samples provided by the United States Environmental Protection Agency. Calibrations are performed at the start of every field season and redone if any changes are made to the fluorometer such as changing the light bulb. Samples for chlorophyll analysis are periodically split with another laboratory or run on two separate fluorometers.

63

WET TISSUE DIGESTION FOR METALS ANALYSIS BY ATOMIC ABSORPTION SPECTROSCOPY

AND/OR ICP EMMISSION SPECTROSCOPY (FISH, CLAMS, MUSSELS, ETC.)

CHEMISTRY LAB SOP UPDATED 04/13/88

65

.

TABLE OF CONTENTS

I

II.

III. IV.

V.

VI.

VII.

VIII. IX. X.

XI.

Sample Storage

1

Sample Transport

1

Sample Receipt and Recording

1

Preparation of Glassware

1

Sample Preparation

1

Sample Weighing

2

Digestion Procedures

2

Sample Digest Filtration

3

Q.C. Samples

4

Safety Precautions

4

Glassware, Chemicals, Equipment and Supplies

4

67

I.

II

Ill

.

.

Sample Storage a.

Fish samples should be wrapped with plastic wrap and stored in sealed plastic bags.

b.

Clam and Mussel samples should be stored in sealed plastic bags.

Sample Transport a.

Samples collected and brought to LES the transported in a cooler with ice.

b.

Samples collected and stored for future delivery to LES should be placed in freezer. Samples should be removed from freezer and placed in a cooler with ice for delivery.

same day

should be

Sample Receipt and Recording a.

Samples received by the LES Chemical Lab personnel are immediately numbered on I.D. Tags and recorded into the Chemistry Lab Log Book.

IV.

b.

Chain of Custody Samples must be accompanied with approved forms.

c.

Samples are processing.

in

freezer

until

they

are

readied

for

Preparation of Glassware a.

V.

stored

All glassware is washed in micronox cleaning solution, rinsed with tap water, acid washed with 40% nitric acid solution, and rinsed 2x or 3x with deionized - distilled water.

Sample Preparation a.

Remove samples from freezer and thaw.

b.

(1)

Fish - The total fish fillet is diced into small sections on a nalgene cutting board using a stainless steel knife. Transfer the diced fish sections into a 40 oz. or small size glass blender top (depending on the amount of sample.)

(2)

Shellfish - Scrub outside of shellfish with a stiff nylon bristled hand brush while rinsing under tap water. Shuck total clam or mussel sample collected into glass blender top.

c.

Using a variable speed blender start homogenizing sample on low speed for 1 or 2 minute intervals (shut off blender between intervals to prevent overheating or burning out the blender motor)

68

.

d.

Once blender blades start making a uniform contact with sample, Continue this use higher speeds for 1 or 2 minute intervals. procedure until sample is thoroughly homogenized.

e.

With a teflon spatula transfer homogenized sample to plastic or Label and number. Place in glass container and seal. refrigerator or freeze samples for up to 6 months. Note For some large fillets, it may be necessary to split sample aliquots, homogenize separately, and recombined in a clean into plastic container. Transfer to multi purpose plastic containers, label and number. ;

f

VI

.

Rinse knife with deionized - distilled water and wipe clean with paper towels. Rinse cutting board with tap water, wash with 5% nitric acid solution, rinse 2x or 3x with deionized - distilled water, and wipe dry. Clean inside of glass blender and rotor blades with hard bristled nylon brush and hot tap water. Rinse with deionized - distilled water 2x or 3x. This cleaning procedure must be repeated after every sample.

Sample Weighing a.

Label and tare 400 ml beaker on balance.

b.

Weigh 10.0 gms of homogenized sample into beaker. Note

;

Teflon spatula used to transfer sample.

c.

Cover beaker with watch glass.

d.

Record weight to nearest 0.1 gm into Digestion workbook.

e.

For every 10 samples or less a duplicate and spiked sample is weighed out.

Note The sample is spiked before digestion using Eppendorf pipets and stock 1000 ppm certified standards. Spiked concentrations are determined for each batch of samples. ;

VII.

Digestion Procedure a.

Add 10 ml concentrated nitric acid to the beaker with sample. Note Acid should be added under fume hood, safety glasses and gloves must be worn. ;

b.

Cover with watch glass.

c.

Place on steam bath and reflux for

d.

Remove watch glass and evaporate to near dryness.

69

2

hours.

e.

Add 10 ml concentrated nitric acid and 10 ml of 30% hydrogen peroxide (H 2 C 2 to beaker. )

VIII.

f.

Cover beaker with watch glass and reflux on steam bath for hours.

g.

Remove watch glass and evaporate to near dryness.

h.

Add approximately 50 ml of 1% vol/vol hot nitric acid to beaker and let stand for 15-30 minutes on steam bath.

2

Sample Digest Filtration a.

Set up nessler tube (100 ml graduated) in rack with filter funnel and #42 Whatman filter paper (18.5 cm).

b.

Wash down filter paper with deionized - distilled water. Discard washing from nessler tube. Rinse nessler tube with deionized distilled water. Replace tube in rack with washed filter paper and funnel.

c.

While decanting sample into Remove beaker from steam bath. funnel, wash sidewalls (inside) and bottom of beakers with deionized - distilled water (use a 500 ml side arm wash bottle).

d.

Rinse beaker with two solution, and filter.

e.

Rinse filter with deionized - distilled water.

f.

Q.S. to 100 ml with deionized - distilled water.

g.

Transfer digest to labeled sample container (125 ml rectangular H.D. polypropylene bottle).

10

ml

aliquots

of

hot

1%

nitric

acid

- To

ensure thorough mixing, pour digest back into nessler tube, and transfer back into sample bottle.

Note High density polypropylene sample containers may become porous. Acid washing or acid soaking in some cases doesn't remove 100% of the contaminates adsorbed within the container. Therefore, it is recommended that once samples have been quantitated, reports have been checked and mailed, that the sample containers be discarded. :

IX.

Q.C. Samples a.

A reference standard, duplicate and spiked samples are processed through the digestion and filtration procedure for each set of 10 samples or less. One reagent blank is processed through the digestion and filtration procedure for every set of samples.

70

X.

Safety Precautions a.

Lab safety practices must be strictly followed.

b.

Protective glasses, gloves, and lab coats must be worn.

c.

Fume hoods should be used whenever necessary.

d.

Safety respirator with acid vapor removal cartridge should be worn.

XI.

Glassware, Chemicals, Equipment and Supplies a.

Glassware 1.

2. 3.

4.

b.

Watch glasses 100 ml graduated nessler tubes

Chemicals 1.

2. 3.

c.

400 ml beakers (heavy duty) 50 ml graduated cylinders

Nitric acid 1000 mg/L Standards for Atomic Absorption spectrophotometers (certified ACS grade) 30% Hydrogen Peroxide (certified ACS grade)

Equipment 1.

2. 3. 4.

Nalgene filter funnels (10 cm diameter) Teflon spatulas 125 ml H.D. Polypropylene sample bottles 4 oz. and 16 ox. polypropylene multipurpose containers with lids.

5.

6. 7.

8. 9.

10. 11. 12. 13. 14. 15. 16. 17. 18.

Repeater pipettors, or automatic dilutors. (500 ml base, 10 ml delivery) 500 ml side arm wash bottle Shellfish shucking knife Stainless steel fillet knife Nessler tube rack Stiff bristled nylon brush (wooden handle) Nalgene cutting board Safety glasses Safety gloves Safety respirator with acid vapor removal cartridges Mettler PE1600 Balance Waring Blender #7012, Model 34BL97, 7 speed Eberback 40 oz. glass blender with handle (#8442) Eberback small size glass blender (#8470)

71

d.

Supplies 1.

2. 3. 4. 5. 6.

Micronox cleaning solution Plastic Bags (sealable) Label tape China marker Lab coat Paper towels

72

APPENDIX B FIELD AND LABORATORY DATA

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TABLE B-4

MUSSEL TISSUE POLYCHLORINATED BIPHENYLS (fjg/g wet weight) Aroclors 1254 and 1242

DEPLOYMENTREPLICATE 1-1 1-2 1-3 1-4 ~x

s

3-1 3-2 3-3 3-4

l

BASELINE

A

0.49 ND** 0.15

0.95

s

4-1 4-2 4-3 4-4 ~x

s

5-1 5-2 5-3 5-4 "x

8

B

0.82

C

0.39

-

-

-

-

-

0.68*

0.066 0.077

-

-

-

-

-

0.047 <0.040* <0.040

1.1 1.0 0.81 1.2 1.028

-

0.423 0.004

"x

STATION

0.166

0.058 <0.040 0.040 ND 0.035 0.025

0.58 0.78 0.58 0.71 0.66 0.10

0.041 <0.040 0.070 <0.040 0.048 0.02

0.61 0.55 0.65 -

0.60 0.05

-

0.40 1.1 1.0 -

0.83 0.38 —

-

0.74 -

0.62 0.68 0.09

0.28 -

0.81 0.68 0.62 0.70 0.097

0.29 0.20 0.26 0.05



0.45

0.94 0.41 0.43 0.59 0.30

-

0.66 0.53 0.547 0.110

Sample not analyzed Less than values averaged as reported number ** ND (none detected) values treated as in calculation of average. *** Tissue exposed from 10/27/87 to 9/21/88. *

83

TABLE B-5

PERCENT LIPID CONCENTRATION IN MUSSEL TISSUE

DEPLOYMENTREPLICATE

STATION BASELINE

A

B

C

1.8

2.2

2.7

-

-

-

-

2.1

1.8

2.2

3-1 3-2 3-3 3-4 X

1.4 1.4 2.8

0.87 2.2 2.4

1.9

2.7 3.0 1.4 1.8 2.2

4-1 4-2 4-3 4-4 X

1.9 1.4 1.3 1.1 1.4

1.3 1.2 1.4 1.6 1.38

5-1 5-2 5-3 5-4

1.8 1.1 1.0 1.3 1.3

1.3 1.2 1.1

1-1 1-2 1-3 1-4 X

2.6 1.7 1.9

-

~x

-

1.2

-

Sample not analyzed * Sample lost in analysis ** Tissue exposed from 10/27/87 - 9/21/88

84

-

-

1.4** 2.7 3.0 -

1.8

2.5 2.75



0.80

1.8 1.4 2.7 2.0 —

1.9 0.98 1.0 1.3

-

1.1 0.83 0.91 1.5 *

1.6 1.6

APPENDIX C SAMPLE STATISTICAL CALCULATIONS

85

KRUSKAL - WALLIS TEST FOR ANOVA

DEPLOYMENT #4 Arsenic HQ HA

:

:

As concentration is the same in all groups As concentration is not the same o< =0.05

Arsenic Base 0.45 0.37 0.45 0.45

A 2.2 4.0 4.5 3.9

(2.5) (1) (4)

(2.5)

C

B

3.2 3.0 2.7 2.8

(6.5) (14) (15) (13)

(12) (11)

2.4 (8)

(9)

2.1 (5) 2.2 (6.5)

(10)

nx = 4

n2 = 4

n3 = 4

n4 = 3

R x = 10

Rj = 48.5

R3 = 42

R4 = 19.5

N = 4+4+4+3 = 15 12

s;

H = N(N+1) =

_E;_

2L

12

10

[

2

2

-3

(N+l)

48 .5

+

2

+

42

2

+

2

19.

]-3(16)

15(16) —

12

[

25 + 588.06 + 441 + 126.75J-48

240 = =

H =

.05

[

1,180.81 ]-48

59.04-48 11.04

number of groups of tied ranks =

^T

=

3

^(V

C = 1-

- tj)

2

.ST N^-N

= 12 =

(2

3

- 2)

+

(2

3

-2)

C = 1-

6 + 6

12

3,360 C = 0.9964 H c = _H_ = C

11.04 0.9964

= 11.0799

H Q 0.05,4,4,4,3 = 7.14 .*«

reject H Q because H c > 7.14

86

NON-PARAMETRIC MULTIPLE COMPARISONS

A Nonparametric Kruskal-Wallis test is applied to Deployment 4 Arsenic values and To determine where the the null hypothesis (they are the same) is rejected. significant differences occur use: Nonparametric Tukey-type multiple comparisons:

SE =

T = 12

N (N + 1)

12(N-1)

12

n>

nB

SE for n =4,4 12

15(16}

12(14)

12

SE for n = 3,4

12

12(14)

12

Samples ranked by mean ranks

Comparison

Baseline

(i)

rank sums

(Rj)

sample sizes

(n,)

mean ranks R " % SE

I

,4

+

I

=

/9.9643 = 3.157

4/

=

15(16)

Q =

f

10

R

2.5

/l + 1'

(3

= /11.624 = 3.41

4J

4 (C)

3

2

(B)

(A)

19.5

42

48.5

10.5

12.13

6.5

RA

Difference

SE

-2o.05,4-

Conclusion

2

vs

1

12.13-2.5=9.63

3.157

3.050

2.639

Reject 1^: [As] different in A & Baseline

2

vs 4

12.13-6.5=5.63

3.41

1.651

2.639

Accept H Q same in A

2

vs

3

vs 1

10.5-2.5=8.0

3

vs

1

do not test

4

vs

1

6.5-2.5=4.0

3

:

[As] & C

do not test 3.157

2.534

2.639

Accept H Q [As] same in B & baseline

3.41

1.173

2.639

Accept H Q [As] same in Baseline

:

:

& C

Overall conclusion: Arsenic concentration is different between baseline and Station A but the same in all other comparisons

87

APPENDIX D

DIVISION OF MARINE FISHERIES PROJECT PLAN AND LABORATORY METHODOLOGY

89

,

QUALITY ASSURANCE PROJECT PLAN

QUALITY CONTROL SECTION OF THE PILOT MONITORING PROGRAM. DEPARTMENT OF ENVIRONMENTAL QUALITY AND ENGINEERING, DIVISION OF WATER POLLUTION CONTROL

FF.EFAFED EY COMMONWEALTH OF MASSACHUSETTS

DEPARTMENT OF FISHERIES. WILDLIFE. ANO ENVIRONMENTAL LAW ENFORCEMENT

FOR U.S. ENVIRONMENTAL PROTECTION AGENCY

REGION 1 WATER MANAGEMENT DIVISION MAY 28, 1987 (revised August 19, 1987)

APPROVALS:

Mr. W. Leigrr Br idges, <£V inci.pal

Investigator

Dr. Wenay Wiltse. Buzzards Bay Project Monitor

Mr.

Charles Porfert, DeDuty Quality Assurance Officer

91

x^Date

Date

Date

TABLE OF CONTENTS

E2Q§ Project Projec: Dare of Date of Project Projec: Quality

!^ame

1

Reauesrea By Reauest Project Initiation Officer__

i

1 1 1

.Monitor

1

Assurance Officer

i

P r o ec : Desc r d t en a. Cc active anc Sccoe i

j

1

i

1

Data Usace C. Design an.G Rationale Mc~ itoring Parameters/Frecuency of Collection D. Protect Fiscal Information E.

Schedule of Tasks ana Procucts Project Organization and Responsibility Data Quality Reauirements ana Assessments Samolm- and Analytical Proceaures Same e Custody Procedures Calibration Proceaures ana Preventive Maintenance Documentation, Data Reduction, and Reporting Da a Va ca 1 on Performance ana System Auatts Correct.ve Action Reoorts .

t

1 1

2 2 2 3 3 3

I

t

'

4 4 4

1

4 -_

4 4

LIST OF TABLES AND FIGURES

Table Laboratory Analyses Table 2. Esttmatea Project Costs Figure 1. Analysis Recues t Form, Cat Cove Marine Laboratory _.___ ".

.

CoDtes sent

to:

Wenav Wilts© (EPA) Char les Porfert (EPA) W. Leigh Br igges (OMF) Jack P. Schwartz (DMF) Nina M. Dustcn (DMF) Chr is Duerr ing (DEQE)

92

1 1

6

.

1.

l

:

i

Project- name: Quality Control

section for DEQE Pilot Monitoring Program

2.

Project recuested by: U.S. EPA. Region 1

3.

Date of reauest

Aon

I

1987

15.

4

Da t e of p r o ec t n 1 a 1 on to be determined bv DEGE

5.

Pro lect Of f icer Rona d Man f r eccn Mr

j

1

l

1

1

\

a

Pro lect Mon tor Dr Wendv W tse :

.

7.

i

:

.

6.

s

:

i

I

Project oescriDticn: A.

Objective ana scope

The Division of Water Pollution Contrc: (Deoartment of Envi ronmmental Quality and Engineering, Commonweal tn of Massacnuset ts) is conducting a monitoring program involving the analysis of the blue mussel, Mvt is eoul s, for trace guantities c: arsenic (As), caamium (Cd). chromium (Cr), coDDer (Cu). lead (Pb), mercury (Hg)'., nickel (Ni), and zinc (Zn) As Dart of this studv, Cat Cove Marine Laboratorv (Division of Marine Fisneries, Department of Fisheries. Wildlife, and Environmental Law Enforcement) has the objective of providing auality control information on a suPset of mussel samples in order to verify trace metal analyses on the larger data Pase and ensure consistency Petween sampling periods for the duration of the monitoring program. i

i

.

B.

Data Usage

(to Pe determined by DEQE.) C. Design and Rationale (to be determined bv DEQE)

D.

Monitoring parameters/f reouency of collection

Mussel swill Pe mom tored for the e gr a foremen t oned metal Samoling will Pe concucted once every two montns ror one year :

i

93

i

E.

TaDle

Parameter Table

Laboratory Analyses

1.

Reference

rax mum \~c Id nc

acid digestion AA/hot vacor

E3 A (1979)

6

ace digestion AA/coid vaocr

Stc. Met noes I6tn ea. 2C2f

acid diaestion AA/flame

ED A (1979)

i

•ameter

Matrix

yn its

eculis tissue

uc/g

M.

.s

<~H

M ethcc

206.5/206.3

213.1

Cr

218.1

_Cu

220.1

Pb

239.1

Mi

249.1 289.1

8.

Project

fiscal

information

Table 2. Est imated "Project Costs

Total

* samDies = 60

Total cos.t for analysis @ Sl70.00/samole

Total

Project Cost

=

$10,200.00

Schedule of Tasks and Projects (to be determined by DECE)

9.

94

i

months

1

1

me

Project Organization and ResDonsibi

10.

I

i

ty

W. Leign Bridges (Massachusetts Division of Marine Fisheries, Mr. Eoston, MA 02202, teleohone [6171 727-3194) will be the principal He will be resDonsible to E?A for the invesriaator for this project. resDons ibi me y comD let ion of the project ana will have over a t ty suomissicn or well ana on as preparation as for data interprerat reoorts to E?A. i

I

I

I

I

i

i

Schwartz (Division of Jack P. Bridges will be assisted by Dr. Marine Fisheries. Cat Cove Marine Laooratory. Salem, MA 01S70, Dr. telecncne [617] 727-3958) as laboratory analysis leader. all will resocnsible for the samoies orocessmc or be Scnwartz rece:vec from DECE including duality control/cuai tv assurance. anal vt ca I- procedures, and ca:a storace and analysis. Mr.

i

i

11.

Data Quality Reduirements and Assessments

Accuracy will be measured as oerdent recovery or an EPA stanaard Corrections will be made reference material analyzed with each batch. Average laooratorv recoveries will be for badkgrouna levels. maintained in the range of 80-120%. Unsptkea blanks will accompany every batch as a further measure of accuracy.

Precision will be measured as the relative standard deviation of triplicate analyses perrormed uoon 10% of the samoLes in eacn batcn. Instrumental preci s ion wi be monitored through the use of triplicate readings on the transition metals oigestate or througn' tr ipl icate readings of oalibration standards7 for arsenic and mercury. Should results vary by more than 10% readinas will be repeated. I

I

Completeness will be measured as the percentage of total samples received that were completely analyzed. We expect to achieve 100% completeness of a analyses. I

12.

-I

Sampling and Analytical Procedures

Elgi.d-Samp.Mng (to be determined by DEQE) Anaj_y_tj_caJ

Pr.ocedu.res

Arsenic analyses will be performed according to U.S. EPA method 206.5/206.3. Mercury analyses will be performed according to Stanoarb Methods (16th ed.) 303F for the Examination of Water and Wastewater. Analysis for cadmium, dhromium, copper, lead, nickel, and zind will be performed using EPA methods 213.1, 218.1, 220.1, 239.1, 249.1, and 289.1, respectively. All methods wi compliment EP A methods muse by DEQE. Concentrations of arsenic will be determined bv atomic aosorption hot vaoor technique. Mercury concentrations will be I

I

95

determined by atomic adsorption cold vaoor techniaue. Transition metal concentrations will be determined by atomic aPsorption flame Analyses will be performed using a Perkin-Elmer Mocel technigues. Arsenic and mercury 3020E atomic absorDtion soect roDnotometer analyses will also use a Perkin-Elmer Mccel PHB 10 mercury hydride Samples will be ccmcared to external stanaarcs suitaoie for system. the metal being analyzed. .

12.

Same e Custodv Procedures. I

Hcmcqen zee mussel samoles will be sniDDea frozen in polyethylene Pags dv DEGE cersonnei accompanied Pv an analysis recues: form (figure "). Laooratory cersonnei wi taxe custccv or ail sample materia: wmen wiil pe assigned laooratory tracking numoers (loggec-in) ana loekee in freezers. Due to a lack of space there are no plans to aremve samples. Any samo e material remaining after the completion of all analyses will Pe made availaPle to DEQE. Mussel samples that are not frozen uDon oelivery will not Pe taken into custoay and returned to DEGE with the shipper. i

I

l

I

14.

CaliPration Procedures and Preventive Maintenance

The atomic aosprption soect roDnotometer will be calibrated through the use of external standards (certified atomic absorption grade standards ootained from Fisher Scientific Company). CaliPration of the instrument will occur at the Peginning of every sampling run and will be checked every ten samples and 'the end of every sampling run. Routine maintenance performed at the time of a run will be noted in the laboratory notebook. The instrument is covered by a maintenance with contract Perkin-Elmer. Any breakdowns will be promptly rerpai red.

i«;

Documentation, Data Reduction, and Reporting

raw data generated during laPoratory analysis will be kept in permanently bound nctepook. A permanently Pound noteoook will be kept of all auality control tests conaucted at the laPoratory. Data printouts will be kept on file and avai ladle for inspection. A. a

16.

All

Data Val idat ion

All data produced Py the laboratory will pe suPiect to a 10055 check for errors in transcription and calculation Py the Senior Chemist. Dr Nina M. Duston, and the Laooratory Analysis Leader, Or. Jack P. Schwartz. The Principal Investigator, Mr. W. Leigh Bridges, will look at all logoooks and notepooks to ensure that reauirements are met. Data wnicn do not meet the SDecified auality reauirements will not Pe mcluaed in the report. Analytical reports will be signed by the Senior Chemist or Laooratory Analysis Leaaer Pefore being

released.

96

17.

Performance and System Audits

Performance will be monitored through EPA water Pollution Laboratory Performance Evaluation Studies which provides for routine intercal ibrat ion with U.S.

18.

EPA every six months.

Corrective Action

Meetincs between all laooratory personnel and Investigator of the stuay will beheld at the Proolems will be identified as samoie batch. When corrective action is recuirea it will oe no zee m the appropriate laboratory ncteocoK.

19.

the Principal comDietion of eacn the stucv progresses. taken immec lately anc

Reports

The reports generated during this study are as follows:

Quality assurance project plan, due May 29. 1987. This report will inciuae the objectives, scope, methods, and products associated A.

wi th

th s study. i

At the completion of analyses of each samoie batch a report will be forwarded to the Principal Investigator for transmittal to appropriate U.S. EPA and DEQE personnel. This report will be completed before the next batch of samples is received.

8.

97

DIVISION OF MARINE FISHERIES

Laboratory Methodology Wet tissue Digestion Procedure for Trace Metals Analysis

Chemicals 1.

HNO3 70.0 - 71.0% n Baker Instra-Analyzed Reagent for Trace Metal Analysis.

2.

H2

2

,

30% Baker Analyzed Reagent.

a.

Weigh approximately 10 grams of blended tissue sample in a preweighed Record sample wet weight to or tared tall form beaker (200 ml). nearest 0.01 grams.

b.

Add 10 ml concentrated HN0 3 to sample in the tall form beaker. Cover with a watch glass and let sit overnight (15 to 16 hours) in ventilated fume hood to cold digest.

c.

Place covered samples on a steam bath until almost all tissue is digested. At this time spike the appropriate quality control samples with a standard spike solution containing concentrations as listed below for the particular species being digested. ialyte

Pb Zn Cu Cr Cd

Lobster ppm

Finf ish ppm 4.0 10.0 10.0 1.0 0.5

4.0 50.0 50.0 1.0 0.5

Shellfish ppm 4.0 20.0 2.0 1.0 0.5

(All standard solutions made in 2% V/V HN0 3

)

Use of these standard spike solutions will result in the enrichment values listed below for the final 50 mL volume of spiked digestate. alyte

Cd Cr Cu Pb Zn

Finf ish ppm

Lobster ppm

0.05 0.10 1.00 0.40 1.00

4.

Reflux the samples for

5.

Remove watch glass after to near dryness.

2

0.05 0.10 5.00 0.40 5.00

Shellfish ppm 0.05 0.10 0.20 0.40 2.00

hours. 2

99

hours of refluxing and evaporate sample

6.

Once all samples are evaporated to near dryness and are at room temperature, add 10 ml concentrated HN0 3 and 10 ml of 30% H 2 2 to each sample. Cover beaker with watch glass and let sit overnight (15 to 16 hours)

7.

Place covered samples on cold stream bath and slowly bring up to (Watch for violent reactions.) Reflux for 2 hours on temperature. steam bath.

8.

Remove watch glass and evaporate to near dryness.

9.

Add approximately 20 ml of a 2% v/v hot HNO3 solution to beaker and let heat for 5 minutes on steam bath.

10.

Remove beaker from steam bath, wipe off any moisture on the outside of beaker and filter the sample using a glass filter funnel with a reeve Angel 802 12.5 cm fluted filter paper or equivalent. Collect filtrate in 50 ml volumetric flask. Rinse beaker with two aliquots of 5-10 ml hot 2% v/v HN03 to remove as much yellow coloring as possible from the filter paper. Remove filter paper and rinse glass funnel with hot 2% HN0 3 taking care not to go over the 50 ml mark.

11.

Q.S. to 50 ml with 2% v/v HN0 3 and transfer to sample containers.

12.

Sample digestate is then analysed for metals on a Perkin Elmer AAS 3030B according to the manufacturer's specifications.

100

Mercury Digestion Method

Chemicals 70.0-71.0%, "Baker Instra-Analyzed" Reagent for Trace Metal Analysis

1.

HN0 3

2.

H 2 S04 , 95.0-98.0%, "Baker Instra-Analyzed" Reagent for Trace Metal Analysis

3.

KMn04

4.

K2S 2

,

8

,

,

"Baker Instra-Analyzed" Reagent for Hg Determination "Baker Instra-Analyzed" Reagent for Hg Determination

Solutions Needed 1.

Potassium permanganate solution: distilled water and dilute to 500 ml.

2.

Potassium persulfate solution: distilled water and dilute to 500 ml

5%

5%

Dissolve 25 g KMn04 in deionized

Dissolve

25

g

K2S 2

8

in

deionized

Procedure for Shellfish Tissue Digestion

1.

Weigh approximately 2 grams of blended sample, to the nearest 0.1 mg, in a pre-weighed or tared 125 ml Erlenmeyer reaction flash.

2.

Add 7.0 ml cone. H 2 S04 and 3.0 ml HN0 3 to each flask and place in a 70 °C water bath.

3.

Remove samples to be spiked from water bath when a colored liquid with no visible tissue has formed. Spike appropriate Q.C. samples with 1.0 ml of 100 ng/ml Hg. This will yield 50 ng Hg enrichment in final sample (refer to Step 8). Return samples to water bath.

4.

Samples should remain in the water bath for four (4) hours.

5.

Remove samples from water bath. Allow to cool to room temperature. Add 5.0 mL deionized distilled water to the samples to cause precipitation of waxy digestion products and decrease the acidity of the sample solutions.

6.

Filter samples through VWR grade 615 9 cm or equivalent filter paper into a stoppered glass 25.0 mL graduated cylinder to remove the waxy precipitate. Rinse the sample flask twice with small amount of 20% v/ HNO3. Rinse filter paper with small amount of 20% HN0 3 taking care not to exceed 25.0 mL of liquid in cylinder.

7.

Q.S. to 25.0 mL with 20% HN0 3

.

Stopper cylinder and shake well.

101

8.

Using acid washed disposable 9 inch Pasteur pipets, divide sample into two Rinse equal portions and place in two clean 125 mL Erlenmeyer flasks. cylinder with two 2.5 mL portions of 20% HN0 3 solution, divide rinses equally between the two sample flasks.

9.

Ice samples.

10.

Add 10 mL KMn04 solution to each flask and let stand 15 minutes in ice bath.

11.

Add 8 mL K2S 2 Og solution to each flask while still in the ice bath.

12.

Add 0.5 - 1.5 g of KMn04 crystals as needed to keep the solutions purple. Remove from ice bath. Samples are left overnight to digest or are placed in a 70°C water bath for 2 to 4 hours. Please note, solutions must remain purple until analysis. Analysis must be with 24 hours.

Washing Procedure for All Labware Used for Metals Analysis

1.

A 12 hour presoak is used if glassware has an organic/waxy film. The presoak solution is made from Terg-A-Zyme (as instructions indicate on the carton)

2.

Wash with soap (Liquinox) and tap water, rinse well with tap water.

3.

Rinse thoroughly with 1:1 HN0 3 followed by 1:1 H 2 S04 (twice). bottle is used to deliver the rinse.

4.

Rinse thoroughly with deionized distilled water at least three times. The deionized distilled water should have a resistance of 2Mohm or higher.

5.

Air dry or place in oven to dry.

6.

Store clean labware in assigned areas, stoppers, whichever is appropriate.

102

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