United States Environmental Protection Agency
Office of Water
Washington, DC
EPA-841-R-14-007
National Coastal Condition Assessment
2015
Field Operations Manual
•>f
** the Ncrt\o*%
May 2015
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National Coastal Condition Assessment 2015 Field Operations Manual
Version 1.0 May 2015 Page i
NOTICE
The National Coastal Condition Assessment provides a comprehensive assessment for coastal
waters across the United States. The complete documentation of overall project
management, design, methods, and standards is contained in four documents:
• National Coastal Condition Assessment 2015: Quality Assurance Project Plan
(EPA-841-R-14-005)
• National Coastal Condition Assessment 2015: Site Evaluation Guidelines (EPA-
841-R-14-006)
• National Coastal Condition Assessment 2015: Field Operations Manual (EPA-
841-R-14-007)
• National Coastal Condition Assessment 2015: Laboratory Operations Manual
(EPA-841-R-14-008)
This Field Operations Manual contains a brief introduction and base and site location
procedures for sampling water chemistry (grabs and in situ measurements), benthic
macroinvertebrates, sediment composition and toxicity, fish tissue, a pathogen indicator, and
physical habitat. These methods are based on the guidelines developed and followed in the
Coastal 2000 and National Coastal Assessment Monitoring and Assessment Program (USEPA,
2001). All National Coastal Condition Assessment Project Cooperators must follow the
methods and guidelines in this Field Operations Manual. Mention of trade names or
commercial products in this document does not constitute endorsement or recommendation
for use. Details on specific methods for site evaluation and sample processing can be found in
the appropriate companion document.
The citation for this document is:
USEPA. 2014. National Coastal Condition Assessment: Field Operations Manual. EPA-841-
R-14-007. U.S. Environmental Protection Agency, Washington, DC.
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1 TABLE OF CONTENTS
1 TABLE OF CONTENTS iii
LIST OF TABLES viii
LIST OF FIGURES ix
ACRONYMS/ABBREVIATIONS x
CONTACT LIST xii
2 BACKGROUND 1
2.1 SURVEY DESIGN 1
2.2 TARGET POPULATION AND SAMPLE FRAME 2
2.3 SITE EVALUATION 3
2.3.1 Site Sample-ability 3
2.3.2 Replacing Sites 4
2.4 DESCRIPTION OF NCCA INDICATORS 5
2.4.1 In Situ Water Column Measurements 5
2.4.2 Water Chemistry (CHEM) and Associated Measurements 5
2.4.3 Algal Toxin (ALGX), Microcystin (MICX) 5
2.4.4 Underwater Video (UVID) 6
2.4.5 Sediment Assessment (SEDG, SEDC, SEDX, SEDO) 6
2.4.6 Benthic Macroinvertebrate Assemblage (BENT) 6
2.4.7 Enterococci Fecal Indicator (ENTE) 6
2.4.8 Fish Tissue (FTIS, FPLG, HTIS) 6
2.5 SUPPLEMENTAL MATERIAL TO THE FIELD OPERATIONS MANUAL 7
2.6 RECORDING DATA AND OTHER INFORMATION 7
2.6.1 Electronic Field Forms 8
2.6.2 Paper Field & Tracking Forms 8
2.7 DATA MANAGEMENT 9
2.8 SAFETY AND HEALTH 10
2.8.1 General Considerations 10
2.8.2 Safety Equipment 11
2.8.3 Safety Guidelines for Field Operations 11
2.8.4 General Safety Guidelines for Field Operations 12
3 INTRODUCTION TO SAMPLING 13
3.1 SITE VISIT DURATION 13
3.2 FIELD CREW MAKEUP 13
3.3 SAMPLING SEQUENCE 13
3.4 SAMPLING CONSIDERATIONS 13
3.4.1 Considerations for Fish Tissue Collection 13
3.4.2 Considerations for Enterococci Collection 14
3.4.3 Other Considerations 14
4 PRE-DEPARTURE ACTIVITIES 17
4.1 DAILY ITINERARIES 18
4.2 INSTRUMENT CHECKS AND CALIBRATION 18
4.2.1 Initial Assembly and Setup Procedures for LI-COR frame, sensor and LI-1400 Datalogger. 19
4.3 EQUIPMENT AND SUPPLY PREPARATION 21
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5 INITIAL SITE PROCEDURES 23
5.1 SITE VERIFICATION 23
5.1.1 Equipment & Supplies 23
5.1.2 Site Verification Procedures 23
5.1.3 Site Relocation 24
5.1.4 Site Characteristics 24
5.2 SITE PHOTOGRAPH 25
5.3 SAMPLE COLLECTION 25
5.4 SECONDARY SEDIMENT OR FISH COLLECTION ZONES 26
5.4.1 Sediment Samples 26
5.4.2 Fish Samples 28
6 WATER QUALITY MEASUREMENTS 30
6.1 SUMMARY OF METHOD FOR IN SITU MEASUREMENTS OF WATER COLUMN TRANSPARENCY, DISSOLVED OXYGEN, pH,
SALINITY, CONDUCTIVITY, TEMPERATURE, AND LIGHT ATTENUATION 30
6.1.1 Equipment and Supplies 30
6.2 SAMPLING PROCEDURE- WATER COLUMN TRANSPARENCY (SECCHI DEPTH) 30
6.3 SAMPLING PROCEDURE - MULTI-PARAMETER SONDE 31
6.3.1 Calibration 31
6.3.2 Dissolved Oxygen Meter 33
6.3.3 pH Meter 33
6.3.4 Salinity/Conductivity Meter 33
6.3.5 Temperature Meter 34
6.4 SAMPLING PROCEDURE - DISSOLVED OXYGEN, pH, TEMPERATURE AND SALINITY/ CONDUCTIVITY 34
6.5 PHOTOSYNTHETICALLY ACTIVE RADIATION (PAR) METER 35
6.5.1 Sampling Procedure—Light Attenuation (LI-1400 Datalogger) 35
7 WATER CHEMISTRY [CHEM], CHLOROPHYLL-^ [CHLA], AND NUTRIENTS [NUTS] SAMPLE
COLLECTION AND PRESERVATION 37
7.1 SUMMARY OF METHOD 37
7.2 EQUIPMENT AND SUPPLIES 37
7.3 SAMPLING PROCEDURE 37
8 ALGAL TOXINS [ALGX] INCLUDING MICROCYSTINS [MICX] 39
8.1 SUMMARY OF METHOD 39
8.2 EQUIPMENT AND SUPPLIES 39
8.3 SAMPLING PROCEDURE 40
8.3.1 Sample Collection 40
8.3.2 Sample Storage 40
9 FECAL INDICATOR (ENTEROCOCCI, [ENTE]) 41
9.1 SUMMARY OF METHOD 41
9.2 EQUIPMENT AND SUPPLIES 41
9.3 SAMPLING PROCEDURE 41
10 PHYTOPLANKTON [PHYT] (GREAT LAKES ONLY) 43
10.1 SUMMARY OF METHOD 43
10.2 EQUIPMENT AND SUPPLIES 43
10.3 SAMPLING PROCEDURE 43
11 UNDERWATER VIDEO [UVID] (GREAT LAKES ONLY) 45
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11.1 SUMMARY OF METHOD 45
11.1.1 Equipment and Supplies 45
11.2 SAMPLING PROCEDURE 45
11.2.1 Initial Setup of Underwater Camera System 45
11.2.2 Initial Setup of Underwater Camera System GPS 46
11.2.3 Underwater Video Recording Procedure 46
11.2.4 Reviewing Underwater Video Files Procedure 47
11.2.5 Directions for Uploading Video Files from DVR 48
12 SEDIMENT COLLECTION 49
12.1 SUMMARY OF METHOD 49
12.2 EQUIPMENT AND SUPPLIES 50
12.3 SAMPLING PROCEDURE 50
12.4 PROCESSING PROCEDURE - BENTHIC MACROINVERTEBRATE [BENT] COMPOSITION AND ABUNDANCE 52
12.5 PROCESSING PROCEDURE- SEDIMENT COMPOSITION, CHEMISTRY, ANDToxiciTY 54
13 FISH TISSUE COLLECTION 57
13.1 ECOLOGICAL CONTAMINATION FISH TISSUE COLLECTION [FTIS] 57
13.1.1 Summary of Method 57
13.1.2 Equipment and Supplies 59
13.1.3 Sampling Procedure 59
13.1.4 Sample Storage and Shipping Preparation 61
13.2 FISH TISSUE PLUG [FPLG] 66
13.2.1 Summary of Method 66
13.2.2 Equipment and Supplies 66
13.2.3 Sampling Procedure 67
13.2.4 Sample Storage 69
13.3 HUMAN HEALTH FISH TISSUE COLLECTION [HTIS] (SELECT GREAT LAKES SITES ONLY) 71
13.3.1 Summary of Method 71
13.3.2 Equipment and Supplies 72
13.3.3 Sampling Procedure 73
13.3.4 Sample Storage and Shipping Preparation 74
14 FINAL SITE ACTIVITIES 77
14.1 GENERAL SITE ASSESSMENT 78
14.1.1 Shoreline Activities and Disturbances 78
14.1.2 Site Characteristics 78
14.1.3 General Assessment 78
14.2 PROCESSING THE FECAL INDICATOR 79
14.2.1 Summary of Method 79
14.2.2 Equipment and Supplies 79
14.2.3 Processing Procedure - Fecal Indicator Filter Blank 79
14.2.4 Processing Procedure - Fecal Indicator Sample 80
14.3 PROCESSING THE CHLOROPHYLL-^ & DISSOLVED NUTRIENTS INDICATORS 82
14.3.1 Summary of Method 82
14.3.2 Equipment and Supplies 82
14.3.3 Processing Procedure 82
14.4 POST-MEASUREMENT CALIBRATION CHECK OF MULTI-PARAMETER SONDE 85
14.5 FIELD DATA & TRACKING FORM REVIEW 85
14.6 SAMPLE PACKAGING AND LABEL REVIEW 86
14.7 SAMPLE SHIPMENT & TRACKING FORM SUBMITTAL 86
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14.7.1 Time-Sensitive Samples 86
14.7.2 Other Samples 87
14.8 EQUIPMENT CLEANUP & CHECK 87
14.8.1 Boat & Trailer Cleanup 88
14.8.2 Post Sampling Equipment Care 88
14.8.3 Additional Decontamination Information 88
15 POST-SAMPLING ACTIVITIES 90
15.1 SAMPLE SHIPPING 90
15.2 TRACKING FORM SUBMITTAL 90
15.3 DATA SUBMITTAL 91
15.3.1 App Users 91
15.3.2 Paper Form Users 91
15.4 TRACKING REMINDERS 92
15.5 SITE EVALUATION SPREADSHEET SUBMITTAL 92
16 FIELD QUALITY CONTROL 93
16.1 STANDARDIZED TRAINING 93
16.2 STANDARDIZED FIELD DATA FORMS 93
16.3 REPEAT SAMPLING 93
16.4 FIELD EVALUATION AND ASSISTANCE VISITS 94
16.4.1 Specifications for QC Assurance 94
16.4.2 Reporting 95
17 LITERATURE CITED 97
APPENDIX A: EQUIPMENT AND SUPPLIES LISTS 99
BASE KIT 99
ADDITIONAL BASE KIT ITEMS - GREAT LAKES CREWS 100
SITE KIT 100
FORM & LABEL PACKET 101
HUMAN HEALTH FISH TISSUE SAMPLING SITE KIT 102
CREW SUPPLIED EQUIPMENT 102
APPENDIX B: FIELD FORMS, LABELS a TRACKING FORMS 105
SITE VERIFICATION (FRONT) 106
SITE VERIFICATION (BACK) 107
FIELD MEASUREMENT (FRONT) 108
FIELD MEASUREMENT (BACK) 109
SAMPLE COLLECTION (FRONT) 110
SAMPLE COLLECTION (BACK) 111
Eco FISH COLLECTION (FRONT) 112
Eco FISH COLLECTION (BACK) 113
SITE ASSESSMENT (FRONT) 114
SITE ASSESSMENT (BACK) 115
HUMAN HEALTH FISH COLLECTION (FRONT) 116
HUMAN HEALTH FISH COLLECTION (BACK) 117
SAMPLE LABELS (MARINE) 118
SAMPLE LABELS (G REAT LAKES ) 119
SITE AND SAMPLE STATUS/WATER CHEMISTRY LAB TRACKING 120
TRACKING: BATCH SAMPLES - OVERNIGHT (CHILLED) 121
TRACKING: BATCH SAMPLES - OVERNIGHT (DRY ICE) 122
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TRACKING: BATCH SAMPLES - GROUND (No ICE) 123
TRACKING: Eco FISH TISSUE - OVERNIGHT (DRY ICE) 124
TRACKING: PACKS 125
TRACKING: HUMAN HEALTH WHOLE FISH SAMPLE - OVERNIGHT (DRY ICE) 126
TRACKING: UVID 127
APPENDIX C: SHIPPING AND TRACKING GUIDELINES 128
TRACKING FORMS 128
T1 - Site & Sample Status/Water Chemistry Lab Tracking Form 129
72 -Tracking: Batch Samples - Overnight (Chilled) Form 129
T3 - Tracking: Batch Samples - Overnight (Frozen) Form 129
T4 - Tracking: Batch Samples - Ground (No Ice) Form 130
T5 - Tracking: Eco Fish Tissue - Overnight (Dry Ice) Form 130
T6 - Tracking: Packs Form 130
77 - Tracking: Human Health Whole Fish Sample - Overnight (Dry Ice) Form [Select Great Lakes
sites Only] 131
T8 - Tracking: UVID Form [Great Lakes Only] 131
SHIPPING GUIDELINES 131
APPENDIX D: FIELD EVALUATION AND ASSISTANCE VISIT CHECKLIST 139
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LIST OF TABLES
TABLE 1.1 CONTACTS xii
TABLE 1.2 REGIONAL COORDINATORS xii
TABLE 2.1 GUIDELINES FOR RECORDING FIELD MEASUREMENTS & TRACKING INFORMATION 9
TABLE 2.2 GENERAL HEALTH & SAFETY CONSIDERATIONS 10
TABLE 4.1 STOCK SOLUTIONS, USES &METHODS FOR PREPARATION 22
TABLE 5.1 EQUIPMENT & SUPPLIES: SITE VERIFICATION 23
TABLE 6.1 EQUIPMENT & SUPPLIES: TRANSPARENCY, DO, pH, SALINITY/CONDUCTIVITY, TEMPERATURE, & LIGHT
ATTENUATION 30
TABLE 6.2 EXAMPLE DEPTH MEASUREMENT INTERVALS 35
TABLE 7.1 EQUIPMENT & SUPPLIES: WATER CHEMISTRY & CHLOROPHYLL-A SAMPLE COLLECTION 37
TABLE 8.1 EQUIPMENT & SUPPLIES: ALGAL TOXINS, MICROCYSTINS 39
TABLE 9.1 EQUIPMENT & SUPPLIES: FECAL INDICATOR (ENTEROCOCCI) SAMPLING 41
TABLE 10.1 EQUIPMENT & SUPPLIES: PHYTOPLANKTON 43
TABLE 11.1 EQUIPMENT & SUPPLIES: UNDERWATER VIDEO 45
TABLE 12.1 EQUIPMENT & SUPPLIES: SEDIMENT COLLECTION 50
TABLE 13.1 EQUIPMENT & SUPPLIES: ECO FISH TISSUE COLLECTION 59
TABLE 13.2 NORTHEAST REGION PRIMARY AND SECONDARY MARINE TARGET SPECIES - WHOLE BODY FISH TISSUE COLLECTION
(ECOFISH) 62
TABLE 13.3 SOUTHEAST REGION PRIMARY AND SECONDARY MARINE TARGET SPECIES - WHOLE BODY FISH TISSUE COLLECTION
(ECOFISH) 62
TABLE 13.4 GULF REGION PRIMARY AND SECONDARY MARINE TARGET SPECIES - WHOLE BODY FISH TISSUE COLLECTION
(ECOFISH) 63
TABLE 13.5 WESTERN REGION PRIMARY AND SECONDARY MARINE TARGET SPECIES - WHOLE BODY FISH TISSUE COLLECTION
(ECOFISH) 64
TABLE 13.6 GREAT LAKES PRIMARY AND SECONDARY TARGET SPECIES - WHOLE BODY FISH TISSUE COLLECTION (ECOFISH) ... 65
TABLE 13.7 EQUIPMENT & SUPPLIES: FISH TISSUE PLUGS 66
TABLE 13.8 PRIMARY AND SECONDARY MARINE TARGET SPECIES FOR FISH PLUG COLLECTION 70
TABLE 13.9 PRIMARY AND SECONDARY GREAT LAKES TARGET SPECIES FOR FISH PLUG COLLECTION 71
TABLE 13.10 EQUIPMENT & SUPPLIES: HUMAN HEALTH FISH TISSUE COLLECTION 73
TABLE 13.11 PRIMARY AND SECONDARY TARGET SPECIES FOR HUMAN HEALTH FISH TISSUE COLLECTION 76
TABLE 14.1 EQUIPMENT &SUPPLIES: ENTEROCOCCI PROCESSING 79
TABLE 14.2 EQUIPMENT & SUPPLIES: CHLOROPHYLL-A & DISSOLVED NUTRIENTS PROCESSING 82
TABLE 16.1 GENERAL INFORMATION NOTED DURING FIELD EVALUATION 95
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LIST OF FIGURES
FIGURE 2.1 EXAMPLE OF AN ESTUARINE SYSTEM COMPRISED OF AN EMBAYMENT PLUS A COMPLEX OF BAYS AND TIDAL RIVERS AND
CREEKS 3
FIGURE 2.2 EXAMPLE OF AN INTER-COASTAL ESTUARINE SYSTEM 3
FIGURE 3.1 FIELD SAMPLING SCENARIO - ACTIVE FISHING METHODS 15
FIGURE 3.2 FIELD SAMPLING SCENARIO - PASSIVE FISHING METHODS 16
FIGURE 4.1 OVERVIEW OF BASE SITE ACTIVITIES 17
FIGURE 4.2 ATTACHMENT OF THE UNDERWATER SENSOR TO THE MOUNTING RINGS (ADAPTED FROM LI-COR, 2006) 20
FIGURE 4.3 LOWERING FRAME ASSEMBLY WITH SENSOR, WEIGHT, AND LOWERING LINE (ADAPTED FROM LI-COR, 2006).... 20
FIGURE 5.1 PRIMARY, SECONDARY AND TERTIARY SAMPLE COLLECTION ZONES 28
FIGURE 5.2 PRIMARY AND SECONDARY FISH COLLECTION ZONES 29
FIGURE 11.1 SETUP DIAGRAM OF UNDERWATER VIDEO SYSTEM 46
FIGURE 12.1 ILLUSTRATION OF ACCEPTABLE & UNACCEPTABLE GRABS FOR BENTHIC COMMUNITY ANALYSIS. AN ACCEPTABLE
GRAB IS AT LEAST 7 CM IN DEPTH (USING A 0.04M2 VAN VEEN SAMPLER), BUT NOT OOZING OUT OF THE TOP OF THE
GRAB, AND HAS A RELATIVELY LEVEL SURFACE 52
FIGURE 14.1 FINAL SITE ACTIVITIES SUMMARY 77
FIGURE 14.2 FILTERING SET-UP FOR ENTEROCOCCI FILTERING 81
FIGURE 14.3 FILTERING SET-UP FOR CHLOROPHYLL-A AND NUTRIENTS FILTERING 85
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ACRONYMS/ABBREVIATIONS
CPR Cardiopulmonary resuscitation
Dl Deionized
DO Dissolved oxygen
DVR Digital video recorder
EAAAP Environmental Monitoring and Assessment Program
EPA Environmental Protection Agency
FLC Field Logistics Coordinator
GED Gulf Ecology Division, U.S. EPA Office of Research and Development
GIS Geographic information system
GL Great Lakes
GPS Global positioning system
GRTS Generalized Random Tessellation Stratified survey design
HOPE High density polyethylene
HQ Headquarters
IM Information Management
MED Mid-Continent Ecology Division, U.S. EPA Office of Research and Development
NAD North American Datum
NARS National Aquatic Resource Surveys
NAWQA National Water-Quality Assessment Program
NCA National Coastal Assessment
NCCA National Coastal Condition Assessment
NEP National Estuaries Program
NHD National Hydrography Dataset
NIST National Institute of Standards
NM Nautical miles
NOAA National Oceanographic and Atmospheric Administration
NRSA National Rivers and Streams Assessment
ORD Office of Research and Development, U.S. EPA
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OSHA Occupational Safety and Health Administration
PAH Polycyclic aromatic hydrocarbon
PAR Photosynthetically active radiation
PBS Phosphate Buffer Solution
PFD Personal flotation device
PSI Pounds per square inch
QAPP Quality Assurance Project Plan
QA/QC Quality assurance/quality control
QCS Quality Check Solution
QRG Quick Reference Guide
SAV Submerged aquatic vegetation
SOPs Standard Operating Procedures
SRM Standard Reference Material
TOC Total organic carbon
TP Total phosphorus
TSS Total suspended solids
USGS United States Geological Survey
WSA Wadeable Streams Assessment
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CONTACT LIST
Table 1.1 Contacts
Role
NCCA Team
Lead
NCCAQA
Coordinator
NARSQA
Coordinator
EPA Logistics
Coordinator
Contractor Field
Logistics
Coordinator
NARSIM
Coordinator
Great Lakes
Human Health
Fish Tissue
Manager
Name
Treda Grayson
Hugh Sullivan
Sarah Lehmann
Colleen Mason
Chris Turner
Marlys Cappaert
Leanne Stahl
Phone/Email
202-566-0916
grayson.treda@epa.gov
202-564-1763
sullivan.hugh@epa.gov
202-566-1379
lehniann.sarah@epa.gov
202-343-9641
niason.colleen@epa.gov
715-829-3737
cturner@glec.coni
541_754_4467
541-754-4799 (fax)
cappaert.rnarlys@epa.gov
202-566-0404
stahl.leanne@epa.gov
Address
US EPA Office of Water
1200 Pennsylvania Avenue NE
(4503T)
Washington DC 20460
US EPA Office of Water
1200 Pennsylvania Ave NE
(4504T)
Washington DC 20460
US EPA Office of Water
1200 Pennsylvania Ave NE
(4503T)
Washington DC 20460
US EPA Office of Water
1200 Pennsylvania Ave NE
(4503T)
Washington DC 20460
Great Lakes Environmental
Center, Inc.
739 Hastings Street
Traverse City, MI 49686
SRA International
200 SW 35th Street
Corvallis OR 97333
US EPA Office of Water
1200 Pennsylvania Avenue NE
(4305T)
Washington DC 20460
Table 1.2 Regional Coordinators
EPA Region 1
EPA Region 3
Name
Phone/Email
Hilary Snook 617-918-8670. snook.hilary@epa.gov
Diane Switzer 617-918-8377. Swit2er.diane@epa.gov
EPA Region 2 Darvene Adams
Bill Richardson
732-321-6700
adams.darvene@epa.gov
215-814-5675
richardson.william@epa.gov
EPA Region 4 Dave Melgaard 404-562-9265, melgaard.david@epa.gov
Address
USEPA Region 1 - New
England Regional Laboratory
11 Technology Drive
North Chelmsford, MA 01863-
2431
USEPA Facilities
Raritan Depot
2890 Woodbridge Avenue
Edison, NJ 08837-3679
USEPA Region 3
1650 Arch Street
Philadelphia, PA 19103-2029
USEPA Region 4
61 Forsyth Street, S.W.
Atlanta, GA 30303-8960
EPA Region 5 Mari Nord 312-886-3017. nord.mari@epa.gov
Ed Hammer 312-886-3019,
| hanimer.edward@epa.gov
USEPA Region 5
77 West Jackson Boulevard
Chicago, IL 60604-3507
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Role
EPA Region 6
EPA Region 9
EPA Region 10
Name
Rob Cook
Forrest John
Mike Schaub
Janet Hashimoto
Sara Roser
Terrence
Fleming
Gretchen
Hayslip
Lillian Herger
Phone/Email
214-665-7141, cook.robert(g).epa.gov
214-665-8368, john.forrest(g).epa.gov
214-665-7314, schaub.mikefSlepa.gov
415-972-3452, hashimoto.janet(g).epa.gov
415-972-3513, roser.sarafSlepa.gov
415-972-3462,
fleming. terrence @.ep a.go v
206-553-1685,
hayslip.gretchen(g).epa.gov
206-553-1074. hero-er.lillian(a)£pa.Q-ov
Address
USEPA Region 6
1445 Ross Avenue
Suite 1200
Dallas, TX 75202-2733
USEPA Region 9
75 Hawthorne Street
San Francisco, CA 94105
USEPA Region 10
1200 Sixth Avenue
Seattle, WA 98101
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2 BACKGROUND
The National Coastal Condition Assessment (NCCA) is one of a series of water assessments
being conducted by states, tribes, the U.S. Environmental Protection Agency (EPA), and
other partners. In addition to coastal waters, the National Aquatic Resource Surveys
(NARS) focus on rivers and streams, lakes, and wetlands in a revolving sequence. The
purpose of these assessments is to generate statistically-valid reports on the condition of
our Nation's water resources and identify key stressors to these systems.
The goal of the NCCA is to address two key questions about the quality of the Nation's
coastal waters:
• What percent of the Nation's coastal waters are in good, fair, and poor
condition for key indicators of water quality, ecological health, and recreation?
• What is the relative importance of key stressors such as nutrients and
contaminated sediments?
The NCCA is designed to be completed during the index period of June through the end of
September. Field crews collect a variety of measurements and samples from preselected
sampling sites that are located at predetermined coordinates.
This manual describes field protocols and daily operations for crews in the NCCA. As a
probability-based survey of our Nation's coastal and estuarine waters, the NCCA is
designed to:
• Assess the condition of the Nation's coastal and estuarine waters at national
and regional scales, including the Great Lakes;
• Identify the relative importance of selected stressors to coastal and estuarine
water quality;
• Evaluate changes in condition from previous National Coastal Assessments
(NCA) starting in 2000; and
• Help build State and Tribal capacity for monitoring and assessment and
promote collaboration across jurisdictional boundaries.
2.1 SURVEY DESIGN
EPA selected sampling locations using a probability based survey design, allowing data
from a subset of sampled sites to be applied to the larger target population, and
permitting assessments with known confidence bounds.
The 2015 NCCA survey design produces:
1. National and regional estimates of the status of all coastal waters, including
major estuary groups and the Great Lakes; and
2. National and regional estimates of the change in status in coastal water
condition between 2010 and 2015.
With input from the states and other partners, EPA used an unequal probability, stratified
design to select 1000 probabilistic sampling events, of which 50% are resample sites (sites
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that were sampled in 2010 and will be sampled again in 2015). Approximately 10% of the
2010 resample sites are also designated "revisit sites," which indicates that they will be
sampled twice in 2015 to assess crew sampling and temporal variability. In addition to the
1000 probabilistic sampling events, a number of intensification sites have been added to
NCCA 2015, many of which were also selected using a stratified probabilistic design.
Sample site stratification is based on major estuaries using the National Oceanic and
Atmospheric Administration (NOAA) Coastal Assessment framework and National Estuary
Program (NEP). The Great Lakes sites are stratified based on the individual Great Lake,
depth zone, and country. Only the shallow nearshore depth zone is included in the design
for NCCA Great Lakes sites. The shallow nearshore depth zone is defined as the region
extending from the shoreline to a depth of 30 meters, and no more than 5 kilometers from
the shoreline.
Oversample sites were drawn to provide alternate sampling sites if primary sites are
rejected and to provide supplemental sampling locations for states that wish to conduct a
state level or NEP-level condition assessment. Also, sites were identified for the Canadian
nearshore zone although sampling of these sites is not a part of the NCCA.
Additional details on the NCCA survey design can be found in the NCCA survey design
documents.
2.2 TARGET POPULATION AND SAMPLE FRAME
The target population for the estuarine resources consists of all coastal waters of the
conterminous United States from the head-of-salt to confluence with the ocean, including
inland waterways tidal rivers and creeks, lagoons, fjords, bays, and major embayments
(see Figure 2.1 and Figure 2.2 for examples). For the purposes of this study, the head-of-
salt is defined as waters with salinity less than 0.5 parts per thousand (ppt) salinity,
representing the landward/upstream boundary. The seaward boundary extends out to
where an imaginary straight-line intersecting two land features would fully enclose a body
of coastal water. All waters within the enclosed area are defined as estuarine, regardless
of depth or salinity.
The target population for the Great Lakes consists of all waters of the Great Lakes of the
United States and Canada. The current target population is restricted to the shallow
nearshore zones of Lake Superior, Lake Michigan, Lake Huron, Lake Erie, and Lake
Ontario. The NCCA Great Lakes sites are restricted to waters within the United States.
Please refer to the Site Evaluation Guidelines and the NCCA Web site
(http://www.epa.gov/owow/monitoring/nationalsurveys.html) for more detailed
information on the target population.
The sample frame was derived from prior National Coastal Assessments developed by EPA
Office of Research and Development (ORD) Gulf Ecology Division (GED). The prior GED
sample frame was enhanced as part of the National Coastal Monitoring Network design by
including information from NOAA's Coastal Assessment Framework, boundaries of NEP and
identification of major coastal systems. For NCCA 2010, information on salinity zones was
obtained from NOAA. For the Delaware Bay, Chesapeake Bay, Puget Sound and the State
of South Carolina, the prior NCA sample frames were replaced by geographic information
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system (GIS) layers provided by the organizations that manage the coastal waters in these
areas, ensuring that prior areas sampled in NCA were not excluded and any differences
from the previous sample frames to the current sample frame are clearly identified in this
NCCA 2015 sample frame. For the Californian Province excluding San Francisco Bay, the
GED sample frame was changed to match a 2004 sample frame used for NCA 2004 study.
In 2013, the sample frame was updated to include information related to 1999-2001 and
2005-2006 NCA sample frames. This update is necessary to provide the information
required to estimate change between the periods of 2010 and 2015. The sample frame for
the Great Lakes sites were obtained from EPA ORD Mid-Continent Ecology Division (MED).
Please refer to the NCCA 2015: Site Evaluation Guidelines for more detailed information
on the target population and exclusion criteria.
Ecorifi^River
RrtriBttjftN
Apjlachee Bay
Legend
| | Estuarine System
uw
] Marine Nearshore Coastal Waters
Legend
| | Estuarine System
Land
Marine Nearshore Coastal Waters
Figure 2.1 Example of an estuarine system
comprised of an embayment plus a complex of
bays and tidal rivers and creeks
Figure 2.2 Example of an inter-coastal estuarine
system
2.3 SITE EVALUATION
Base site sampling points were drawn using a Generalized Random Tessellation Stratified
(GRTS) survey design, a stratified design that gives all points within a target population
equal probability of selection. Each point selected as a sample site is designated the "X-
site" and represents the point at which sample collections are targeted.
2.3.1 SITE SAMPLE-ABILITY
X-sites will be found in waterbodies of varied sizes and shapes depending on coastal
morphology. Site depth and salinity are considered when the initial site draw is made;
therefore, those conditions should not generally be a factor when choosing to replace a
planned sampling site. However, there may be instances when a field crew determines
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that an X-site does not meet the operational definition of an estuary in marine
environments, or lacustrine and nearshore coastal waters in the Great Lakes. Sampleable
sites must:
• Have access to open water;
• Be navigable using a shallow-draw boat. Typically this means that the depth of
the X-site is generally > 1 meter. Actual sampleable depths, however, may be
adjusted based on the vessel and sampling equipment being used, and wave
action at the site observed by the field crew.
If the specific site does not fit the definition of a sampleable site, and every attempt to
relocate a site within the margin provided has been made (see Section 5.1.3), complete
the appropriate "Non-Sampleable-Permanent" category on the Site Verification (Front)
form. Document the reason for not sampling the site in the comments section of the form.
Add any additional explanation as required. (For complete details on the site evaluation
process, refer to the NCCA Site Evaluation Guidelines).
2.3.2 REPLACING SITES
It is likely that some sites will be determined to be unsampleable; therefore, a number of
backup sites, in the form of an oversample list, are provided to each state. A site can be
deemed unsampleable for any number of reasons, including being too shallow to properly
operate sampling equipment, in the middle of a navigational channel where it is unsafe,
or practically on top of a neighboring site.
When a site is determined to be unsampleable, field crews will document the sampling
status of the site and select the next oversample site within the same stratum (i.e. same
state and estuary type or Great lake) and the same base year (Base 10 sites must be
replaced with Base 10 oversamples sites and Base 15 sites must be replaced with Base 15
oversamples sites). This process maintains the probabilistic integrity of the survey. This
process is handled through the Site Evaluation Spreadsheets that EPA Headquarters (HQ)
has provided for each state. These spreadsheets are available on the NARS SharePoint
site. Please refer to the NCCA Site Evaluation Guidelines for more detailed information
on determining site sampling status and completion of the Site Evaluation Spreadsheets.
These spreadsheets will be turned in when sampling is completed, or throughout the field
season should it be necessary for communicating the replacement of specific sites to EPA
HQ and the Contractor Field Logistics Coordinator (FLC).
If a dropped site is designated as a revisit site (designated "RVT2" in the panel code)
and/or a human health fish tissue site (designated by "FT" in the panel code), then the
replacement site takes on the RVT2 and/or FT assignment. That is the site must be
visited twice in 2015 (if RVT2) and human health fish tissue collected (if FT).
If a site is generally sampleable, but one or more indicators cannot be collected (e.g. no
fish caught or site is too deep to collect sediment), the site should not be dropped.
Rather, the crew will flag that indicator and document the reason why the indicator could
not be collected. See Section 12 for information regarding the collection of sediment
samples, which is potentially the indicator crews may experience difficulty collecting.
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2.4 DESCRIPTION OF NCCA INDICATORS
Indicators for the 2015 survey will basically remain the same as those used in 2010 and
other past coastal surveys, with a few modifications. Again, sample collection methods
and laboratory methods will reflect freshwater and saltwater matrices to account for
marine and Great Lakes sampling.
2.4.1 IN SITU WATER COLUMN MEASUREMENTS
2.4.1.1 Hydrographic Profile
Measurements for dissolved oxygen (DO), pH, salinity (at marine sites) or conductivity (at
freshwater sites), and temperature will be taken with a calibrated water quality meter or
multi-parameter sonde at each site. Measurements will be taken at specific depth
intervals within 37 meters of the X-site. The specific location of the profile (and
subsequently the area where several samples are collected) is referred to as the Y-
location. This information will be used to detect extremes in condition that might indicate
impairment.
2.4.1.2 Light Attenuation
A Photosynthetically Active Radiation (PAR) meter will be used to obtain a vertical profile
of light in order to calculate the light attenuation coefficient at each station. PAR
measurements are taken at the same depths as other water column indicators.
2.4.1.3 Secchi Disk Transparency
A Secchi disk is a commonly used black and white patterned disk used to measure the
clarity of water within a visible distance.
2.4.2 WATER CHEMISTRY (CHEM) AND ASSOCIATED MEASUREMENTS
Water chemistry measurements will be used to determine nutrient enrichment, as well as
classification of trophic status. Parameters measured include total and dissolved nitrogen
and phosphorus.
2.4.2.1 Chlorophyll-a (CHLA)
Chlorophyll-a is the green pigment used in photosynthesis by plants and algae. Its
measurement is used to determine algal biomass in the water.
2.4.2.2 Dissolved Nutrients (NUTS)
A portion of the filtrate produced from the processing of the chlorophyll-a sample will be
collected in the field and processed in the laboratory for dissolved nutrients.
2.4.2.3 Phytoplankton Assemblage (PHYT)
Phytoplankton are plant microorganisms that float in the water, such as certain algae, and
are the primary source of energy in most lake systems (Schriver et al. 1995).
Phytoplankton are highly sensitive to environmental changes in ecosystems (e.g., turbidity
and nutrient enrichment). Phytoplankton will be collected in Great Lakes sites only.
2.4.3 ALGAL TOXIN (ALGX), MICROCYSTIN (MICX)
Algae are microscopic organisms found naturally at low concentrations in freshwater and
marine systems. They often form large blooms under optimal conditions, potentially
affecting water quality as well as human health and natural resources. Microcystis, for
example, is one organism that produces microcystin, a potent liver toxin. One water
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sample is taken to analyze for a suite of algal toxins and another will be taken specifically
for microcystin.
2.4.4 UNDERWATER VIDEO (UVID)
At Great Lakes sites only, crews will use an underwater video camera with recorder to
capture one minute of video focused on the substrate at the Y-location. Video will be
used in the lab to visually document the bottom composition, and record the presence or
absence of zebra mussels, Cladophora, or other organisms.
2.4.5 SEDIMENT ASSESSMENT (SEDG, SEDC, SEDX, SEDO)
Sediment grab samples will be obtained to measure sediment composition (e.g., grain size
[SEDG] and percent moisture, organic content, etc. [SEDC]), toxicity [SEDX], and
contaminant chemistry [SEDO] in order to determine sediment condition.
2.4.6 BENTHIC MACROINVERTEBRATE ASSEMBLAGE (BENT)
Benthic macroinvertebrates are bottom-dwelling animals without backbones
("invertebrates") that are large enough to be seen with the naked eye ("macro").
Examples of macroinvertebrates include: aquatic worms, mollusks, and crustaceans.
Populations in the benthic assemblage respond to a wide array of stressors in different
ways so that it is often possible to determine the type of stress that has affected a
macroinvertebrate assemblage (Klemm et al., 1990). Because many macroinvertebrates
have relatively long life cycles of a year or more and are relatively immobile, the
structure of the macroinvertebrate assemblage is a response to exposure of present
and/or past conditions. The benthic macroinvertebrate data will serve as the basis for
assessing aquatic community health.
2.4.7 ENTEROCOCCI FECAL INDICATOR (ENTE)
Enterococci are bacteria that are endemic to the guts of warm blooded creatures. These
bacteria, by themselves, are not considered harmful to humans but often occur in the
presence of potential human pathogens (the definition of an indicator organism).
Epidemiological studies of marine and fresh water bathing beaches have established a
direct relationship between the density of Enterococci in water and the occurrence of
swimming-associated gastroenteritis.
2.4.8 FISH TISSUE (FTIS, FPLG, HTIS)
The fish tissue indicator [FTIS], which measures bioaccumulation of persistent toxics, is
used to estimate the ecological risks associated with fish consumption by wildlife. In this
study fish will be collected and whole body tissue will be homogenized and analyzed to
estimate concentrations of target contaminants. Various studies have been conducted on
contaminants in different tissues of the fish (e.g., whole fish, fillets, or livers). For this
study the focus will be on analyzing whole fish [FTIS] for contaminants to generate data
for ecological purposes are referred to as the ecofish sample. At revisit sites, ecofish
samples will only be collected during visit 1.
Crews will also collect fish tissue plugs [FPLG] at all NCCA Sites. The plugs will be sent to
the lab for analysis of mercury contamination levels to assess the risk to humans of
consuming fish tissue. If the fish plug sample is taken from fish other than those being
collected for ecological analysis, the fish will be released back into the waters from which
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they were collected. At revisit sites, fish plug samples will only be collected during visit
1.
In the Great Lakes only, additional fish composite samples will be collected at 150 of the
225 sites (ideally the first 15 of both the Base 10 and Base 15 sites for a combined total of
30 sites per lake). Fillet tissue from these samples will be homogenized and analyzed to
generate fish contamination data related to human health [HTIS]. Fish submitted in the
human health fish tissue sample should remain intact and fish plugs are not to be taken
from these fish. At Great Lakes revisit sites that are also human health fish tissue sites,
crews that are unsuccessful at collecting the human health fish tissue sample during visit
1 are expected to attempt the collection of that sample during visit 2.
2.5 SUPPLEMENTAL MATERIAL TO THE FIELD OPERATIONS MANUAL
The Field Operations Manual describes field protocols and daily operations for crews to
use in the NCCA. Following these detailed protocols will ensure consistency across regions
and reproducibility for future assessments. Before sampling a site, crews should prepare a
Site Packet for each site containing pertinent information to successfully conduct
sampling. This site packet typically includes a road map or navigation chart and a set of
directions to the site, topographic/bathymetric maps, land owner access forms (where
applicable), sampling permits (if needed), site evaluation forms and other information
necessary to ensure an efficient and safe sampling day.
Field crews will also receive a Quick Reference Guide that contains tables and figures
summarizing field activities and protocols from the Field Operations Manual. This
waterproof handbook will be the primary field reference used by field crews after
completing the required field training session. Field crews are also required to keep the
Field Operations Manual as well as other equipment manuals (probes, etc.) available in
the field for reference and for possible protocol clarification.
Large-scale and/or long-term monitoring programs such as those envisioned for national
surveys and assessments require a rigorous Quality Assurance (QA) program that can be
implemented consistently by all participants throughout the duration of the monitoring
period. QA is a required element of all EPA-sponsored studies that involve the collection
of environmental data (USEPA 2000a, 2000b). Field crews will be provided a copy of the
integrated Quality Assurance Project Plan (QAPP). The QAPP contains more detailed
information regarding QA/ Quality Control (QC) activities and procedures associated with
general field operations, sample collection, measurement data collection for specific
indicators, data reporting activities, and the information management plan for this
project. For more information on the QA procedures, refer to the National Coastal
Condition Assessment 2015: Quality Assurance Project Plan (EPA-841-R-14-005).
2.6 RECORDING DATA AND OTHER INFORMATION
Field data and sample information must be recorded completely, legibly, accurately,
and consistently. The cost of a sampling visit coupled with the short index period
severely limits the ability to resample a site if the initial records are inaccurate or
illegible. Illegible or incorrect information can result in substantially increased time to
transfer information from field forms to the National Aquatic Resource Surveys
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Information Management (NARS IM) system. Guidelines for recording field measurements
are presented in Table 2.1.
Field crews may choose to record field data in one of two formats: electronic or paper
field forms. Paper tracking forms must be included in every cooler/box which contains
samples being shipped to the labs and must also be submitted to NARS IM electronically
(via App, fillable pdfs, scans, or fax). See additional information on each format below.
All samples need to be identified and tracked, and associated information for each sample
must be recorded. To assist with sample identification and tracking, tracking forms and
labels are preprinted and provided by EPA with sample ID numbers.
2.6.1 ELECTRONIC FIELD FORMS
Field crews may choose to utilize the NARS App to complete data collection. The NARS
App is available in both Android and iOS formats and is available on the NARS SharePoint
site. If a field crew is utilizing the iOS app, they must first provide (via email) the Unique
Device Identifier (UDID) for their device to NARS IM and the EPA Logistics Coordinator.
This will allow the App to be loaded on a particular device.
The NARS App is the preferred format for data submission as it cuts down on processing
time required in scanning paper field forms, prevents data entry errors, eliminates
redundant entry of common fields, eliminates issues caused by illegible entries, and
provides validation checks of fields. In addition, the app generates all sample IDs based on
the initial entry of the CHEM sample ID and includes fish pick lists for consistent naming.
If field crews are utilizing this form of data entry, they will upload site sketches of their
sites to the NARS SharePoint site.
2.6.2 PAPER FIELD & TRACKING FORMS
Paper field forms are utilized by some crews. Paper tracking forms must be included in
every cooler regardless of whether crews choose to use the NARS App or paper field
forms. It is important that field crews adhere to the guidelines listed in Table 2.1 when
completing paper forms.
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Table 2.1 Guidelines for recording field measurements & tracking information
Activity
Guidelines
Field Measurements
Data Recording
Data Qualifiers
(Flags)
• Record measurement values and observations on data forms preprinted on water-resistant
paper.
• Use No. 2 pencil only (fine-point indelible markers can be used if necessary) to record
information on forms.
• Record data and information using correct format as provided on data forms.
• Be sure to accurately record site and sample IDs.
• For all primary sampling visits indicate the event as Visit 1. For revisit sites use Visit 2 to
indicate the second sampling event during the same season.
• Print legibly (and as large as possible). Clearly distinguish letters from numbers (e.g., 0 versus
O, 2 versus Z, 7 versus T or F, etc.), but do not use slashes.
• When recording comments, print or write legibly. Make notations in comments field only;
avoid marginal notes. Be concise, but avoid using abbreviations or "shorthand" notations. If
you run out of space, attach a sheet of paper with the additional information, rather than trying
to squeeze everything into the space provided on the form.
• Use only defined flag codes and record on data form in appropriate field.
• F» = Miscellaneous flags (» = 1, 2, etc.) assigned by a field crew during a particular sampling
visit (also used for qualifying samples).
• Define each flag in comments section on data form. Flags must be re-defined on each form
and on forms from different stations.
Sample Labels
• Use adhesive labels with preprinted sample IDs and follow the standard recording format for
each type of sample.
• Use a fine tipped permanent marker to record information on label. Cover the completed label
with clear tape.
• Record sample ID from label and associated collection information on sample collection form
preprinted on water-resistant paper.
Sample Collection and Tracking
Sample
Qualifiers (Flags)
• Use only defined flag codes and record on sample collection form in appropriate field.
• F» = Miscellaneous flags (»=1, 2, etc.) assigned by a field crew during a particular sampling
visit (also used for field measurements).
• Define each flag in comments section on data form. Because the same flag may have different
meanings at different sites, re-define each flag when used on a form and when used on forms
from different stations.
Review of Labels
and Data
Collection Forms
• Before leaving site, compare information recorded on labels and sample collection form to
ensure agreement and accuracy.
• Before leaving site, review labels and data collection forms for accuracy, completeness, and
legibility.
• The Field Crew Leader must review and initial all data collection forms, verifying consistency,
correctness and legibility, before transfer to NARS IM.
2.7 DATA MANAGEMENT
All field crews will be given access to the NARS SharePoint site. This site will be a
resource for field crews to access important NCCA documentation as well as for
facilitating document transfer to and from field crews.
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2.8 SAFETY AND HEALTH
Sample collection and analysis can pose significant risks to personal safety and health.
This section describes recommended training, communications, safety considerations,
safety equipment and facilities, and safety guidelines for field operations.
2.8.1 GENERAL CONSIDERATIONS
Important considerations related to field safety are presented in Table 2.2. The Field
Crew Leader is responsible for ensuring that all field personnel have successfully
completed the necessary safety courses and follow all safety policies and procedures.
Please follow your own agency's health and safety protocols. Additional sources of
information regarding safety-related training include the American Red Cross (2006), the
National Institute for Occupational Safety and Health (1981), and U.S. Coast Guard (1989).
Field crew members should become familiar with the hazards involved with sampling
equipment and establish appropriate safety practices prior to their use. Make sure all
equipment is in safe working condition. Personnel must consider and prepare for hazards
associated with the operation of motor vehicles, boats, winches, tools, and other
incidental equipment. Boat operators must meet any state requirements for boat
operation and be familiar with U.S. Coast Guard rules and regulations for safe boating
contained in the pamphlet, "Federal Requirements for Recreational Boats," available
from a local U.S. Coast Guard Director or Auxiliary or State Boating Official (U.S. Coast
Guard, 1989). While on the water, all crew members must wear Personal Flotation
Devices (PFD). All boats with motors must be equipped with fire extinguishers, boat horns,
PFDs, and flares or other U.S. Coast Guard approved signaling devices.
Table 2.2 General health ft safety considerations
Recommended first aid and cardiopulmonary resuscitation (CPR)
Training vehicle safety (e.g., operation of 4-wheel drive vehicles, trailering boats, etc.)
field safety (weather, personal safety, navigation, site reconnaissance prior to sampling)
equipment design, operation, and maintenance
handling of chemicals and other hazardous materials
Communications check-in schedule
sampling itinerary (vehicle used & description, time of departure & return, travel route and
destination)
contacts for police, ambulance, hospitals, fire departments, search and rescue personnel
emergency services available near each sampling site and base location
cell (or satellite) phone and VHP radio.
Personal Safety field clothing and other protective gear including PFDs for all crew members
medical and personal information (allergies, personal health conditions)
personal contacts (family, telephone numbers, etc.)
physical exams and immunizations
Prior to beginning a sampling day, each field crew must develop an Emergency
Communications Plan. This plan will include contacts for police, fire departments,
emergency medical services, hospitals and search and rescue personnel. In addition, the
plan must include daily check-in procedures with personnel who will not be in the field. A
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copy of the plan should be filed with a supervisor, safety specialist or other staff member
who is not in the field. All field personnel must be fully aware of all lines of
communication and able to initiate emergency communications if needed. Field crew
members must carry clothing and equipment to protect from exposure to different
weather conditions. Inadequate clothing could lead to hypothermia, heat exhaustion or
heat stroke. Field personnel must be able to swim. A PFD and suitable footwear must be
worn at all times while on board a boat.
2.8.2 SAFETY EQUIPMENT
Crews may face many hazards when working in coastal areas. Broken glass or other sharp
objects may be embedded in the substrate. Infectious agents and toxic substances may be
present in the water or sediment. Dangerous weather may approach with little warning.
Vessels can lose power and navigation.
Field crews must stock appropriate safety apparel such as gloves, foul weather gear,
safety glasses, etc., and use them when necessary. All vessels must have first aid kits, fire
extinguishers and blankets available in the field, and crew members must be trained in
how to use them. All crews must carry cellular or satellite telephones and all crew
members must be proficient in how to use them. Crews must carry supplies such as clean
water, anti-bacterial soap, and ethyl alcohol for cleaning exposed body parts that may
have been contaminated by pollutants in the water.
2.8.3 SAFETY GUIDELINES FOR FIELD OPERATIONS
Personnel participating in field activities must be in sound physical condition and have a
physical examination annually or in accordance with organizational requirements.
Field crew members must become familiar with the health hazards associated with
collecting, preserving, and storing field samples. All surface waters and sediments are
considered potential health hazards due to the potential presence of toxic substances or
pathogens, and chemical fixing and/or preserving agents are often comprised of hazardous
materials. In addition, chemical wastes can be flammable, explosive, toxic, caustic, or
chemically reactive. Therefore, all chemical wastes must be discarded according to
standardized health and hazards procedures (e.g., National Institute for Occupational
Safety and Health [1981]; U.S. EPA [1986]).
During the course of field research activities, field crews may observe violations of
environmental regulations, discover improperly disposed hazardous materials, or observe
or be involved with an accidental spill or release of hazardous materials. In such cases
proper actions must be taken and field personnel must not expose themselves to
something harmful.
The following safety guidelines should be applied:
First and foremost, protect the health and safety of all personnel. Take necessary steps to
avoid injury or exposure to hazardous materials. If you have been trained to take action
such as cleaning up a minor fuel spill during fueling of a boat, do it. However, you should
always err on the side of personal safety.
Field personnel should never disturb or retrieve improperly disposed hazardous materials
from the field to bring them back to a facility for "disposal". To do so may worsen the
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impact, incur personal liability for the crew members and/or their respective
organizations, cause personal injury, or cause unbudgeted expenditure of time and money
for proper treatment and disposal of the material. Notify the appropriate authorities so
they may properly respond to the incident. For most environmental incidents, the
following emergency telephone numbers should be provided to all field crews: State or
Tribal department of environmental quality or protection, U.S. Coast Guard, and the U.S.
EPA regional office. In the event of a major environmental incident, the National
Response Center may need to be notified at 1-800-424-8802.
2.8.4 GENERAL SAFETY GUIDELINES FOR FIELD OPERATIONS
• At least two crew members must be present during all sample collection
activities, and no one should be left alone while out on the water.
• Use caution and wear a suitable PFD.
• Use caution using the Ponar-type samplers. The jaws are sharp and may close
unexpectedly.
• Exposure to water and sediments should be minimized as much as possible. Use
gloves if necessary, and clean exposed body parts as soon as possible after
contact.
• All electrical equipment must bear the approval seal of Underwriters
Laboratories and must be properly grounded to protect against electric shock.
• Use appropriate protective equipment (e.g. gloves, safety glasses) when
handling and using hazardous chemicals.
• Crews working in areas with venomous snakes must check with the local Drug
and Poison Control Center for recommendations on what should be done in case
of a bite from a venomous snake.
• Any person allergic to bee stings, other insect bites, or plants (i.e., poison ivy,
oak, sumac, etc.) must take proper precautions and have any needed
medications handy.
• Field personnel should be familiar with the symptoms of hypothermia and know
what to do in case symptoms occur. Hypothermia can kill a person at
temperatures much above freezing (up to 10°C or 50°F) if he or she is exposed
to wind or becomes wet. Immersion in the cool waters experienced during the
summer along most of the marine coasts and Great Lakes can also rapidly result
in hypothermia.
• Field personnel should be familiar with the symptoms of heat/sun stroke and
be prepared to move a suffering individual into cooler surroundings and hydrate
immediately.
• Handle and dispose of chemical wastes properly. Do not dispose of any
chemicals in the field.
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3 INTRODUCTION TO SAMPLING
This Field Operations Manual describes procedures for collecting samples for the NCCA
2015. Overall, the same indicators will be collected at both estuarine and coastal
freshwater Great Lakes sites, though some of the sampling will be conducted using
different equipment. Field crews at all Great Lakes sites will collect additional water
samples to be analyzed for phytoplankton and will record underwater video of the bottom
substrate. At selected Great Lakes sites, crews will collect an additional fish tissue
samples to be analyzed for human health risks.
This section presents a general overview of the field activities and guidelines for field
operations, recording data and labeling samples. This section also describes field crew
makeup and other sampling considerations.
3.1 SITE VISIT DURATION
NCCA field methods are designed to be completed in one field day. Depending on the time
needed for sampling and travel, crews may require an additional day to complete
sampling, pre-departure and post-sampling activities (e.g., cleaning equipment, repairing
gear, shipping samples, and traveling to the next site). Remote sites with lengthy or
difficult approaches may require more time, and field crews must plan accordingly.
Conversely, some sites may be in relatively close proximity to each other, allowing
multiple sites to be sampled in a single day.
3.2 FIELD CREW MAKEUP
A field crew typically consists of three to four people. However, a minimum of two people
may be able to properly execute sampling activities. To ensure safety, at least two people
are always required in a boat when conducting field work for the NCCA. In order to
organize field activities efficiently, each field crew should define roles and responsibilities
for each crew member prior to beginning field activities. One crew member is primarily
responsible for boat operation and navigation. Additional crew members assist with
sample collection/processing and/or provide logistical support.
3.3 SAMPLING SEQUENCE
The field crew arrives at the site in the early morning to complete the sampling in a single
day. The typical sampling scenarios are shown in Figure 3.1 and Figure 3.2.
3.4 SAMPLING CONSIDERATIONS
3.4.1 CONSIDERATIONS FOR FISH TISSUE COLLECTION
The sequence of daily field activities may differ depending on whether the field crew is
collecting fish that day or another day, or using active (trawling, seining, hook and line,
etc.) or passive (gill net, hoop net, long-lines, etc.) fish collection methods. Other minor
modifications to the sampling scenario may be made by crews; however, the sequence of
sampling events presented in the following figures (depending on the type and timing of
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fish collection) should be adhered to and is based on the need to protect some types of
samples from contamination and to minimize holding times once samples are collected.
3.4.2 CONSIDERATIONS FOR ENTEROCOCCI COLLECTION
Enterococci levels tend to be highest in the morning prior to high levels of solar
irradiation; therefore, these samples must be collected as early in the day and with as
little water and sediment disturbance as possible. Regardless of when the Enterococci
samples are collected, crews must complete filtration within six hours of collection.
Enterococci samples not filtered within six hours of collection must be discarded,
recollected, and filtered.
3.4.3 OTHER CONSIDERATIONS
Crew members responsible for collecting water chemistry, sediment grabs, and fish tissue
must remember to not apply sunscreen or other chemical contaminants until after each of
these samples is collected to avoid compromising the integrity of the sample (or
implement measures to reduce contamination by such chemicals if applied such as
washing, wearing long gloves, etc.).
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Confirm X-Site
[5.1.2;
Sampieable
Not Sampleabte
Site Characteristics
[5.1.4]
Relocate
[5.1.31
Hydro-graphic
Profile [6.2]
PHYT(GL only) [103J
ALGX, MICX [8.3?
VERIFICATION
NUTS(N,P)
Aliquot [14.3]
COLLECTION
SEDX, SEDO, SEDC,
SEDG[12,5I
Fishing HTJSfGlon
113.3.3,
PROCESSING
Filters [14.3]
Figure 3.1 Field Sampling Scenario - Active Fishing Methods
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Confirm X-Site
[5.1.2]
Sampteable ""-—r--'' NotSampteabte
Sits Ch a ract eristics
[5.1.4]
Fishing
(set gear)
jr ---..
Relocate
[5.1.3;
VERIFICATION
Hydrographic
Profile [6.21
.
YT(GL
[10.3]
ALGX, MICX [8.3]
^^^^m
UVIL
I "
UVID(GLonly}
[11.2.3]
BENT [1.2.4]
COLLECTION
SEDX, SEDO, SEDC,
SEDG [1.2.5]
Retrieve
Gear
HflS(GL only) [13.33]
PROCESSING
1
FPLG [13.2.3]
r ~
^T NUTS[N,P) /
^ Aliquot [14.3]
' CHLA
Filters [14.3]
/ ENTE A
\ Filters ! ^B
Figure 3.2 Field Sampling Scenario - Passive Fishing Methods
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4 PRE-DEPARTURE ACTIVITIES
Field crews conduct a number of activities at their base site (i.e. office or laboratory,
camping site, or hotel) before departure to the site and after returning from the field.
Before leaving the base site, the crews must know: (1) where they are going; (2) that the
site is accessible and that, if necessary, they have permission to sample it; and (3) that
equipment and supplies needed to complete the sampling effort are available and in good
working order. After sampling, crews must ensure that: (1) samples are labeled, packed,
and shipped appropriately; (2) the sampling event is communicated to EPA; and (3)
equipment and supplies are cleaned and replenished as necessary.
're-departure
Activities
Sample Site
Post-Sampling
Activities
Figure 4.1 Overview of base site activities
•Pre-departure Activities
•Crew Leader - Prepare daily itinerary
•Whole Crew - Site verification
•Crew Members - Instrument checks
& calibration, equipment & supplies
preparation
•Post-Sampling Activities
•Crew Leader
•Review forms and labels
•File status report via email to tracking team
•Crew Members
•Filter, preserve & inspect samples
•Clean boats with 1-10% bleach solution
•Perform safety checks on boat (when trailering
between one water body to distinctly different
water body)
•Clean (and repair, if needed) sampling gear
•Charge or replace batteries
•Refuel vehicle and boat
•Obtain ice and other consumable supplies as
needed
•Package and ship samples & data forms
Pre-departure activities are included here, while post-sampling activities are discussed in
Section 14: Final Site Activities and Section 15: Post-Sampling Activities. Pre-departure
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activities include the development of a daily itineraries, instrument checks and
calibration, and equipment and supply preparation.
4.1 DAILY ITINERARIES
Field Crew Leaders are responsible for developing daily itineraries and site information,
which are compiled as a Site Packet. This site packet typically includes maps,
navigational charts, contact information, copies of permission letters, permits, access
instructions, location of FedEx offices, and location and contact information of hospitals
or other emergency services. Additional activities include confirming the best access
routes, calling the landowners or local contacts, confirming lodging plans, and
coordinating rendezvous locations with individuals who must meet with field crews prior
to accessing a site.
Also, the Field Crew Leader must identify appropriate boat ramps or marinas and gas
docks. If the crew is planning a multiple day/multiple site trip, information for each day
and site must be developed and compiled into separate site packets.
4.2 INSTRUMENT CHECKS AND CALIBRATION
Each field crew must test and calibrate instruments prior to sampling. Equipment can be
calibrated either prior to departure for the site or at the site. However, due to variations
in elevation, DO probes must be calibrated at the site. The field crew will verify site
location using a global positioning system (GPS) receiver. They will collect measurements
using a Photosynthetically Active Radiation (PAR) meter and a multi-parameter unit for
measuring DO, pH, temperature, salinity (recorded at marine sites) and conductivity
(measured at freshwater sites). Field crews must have access to backup instruments if any
instruments fail the manufacturer performance tests or calibrations. Prior to departure,
field crews must perform the following checks and calibrations:
• If using a hand-held GPS unit, turn on the GPS receiver and check the batteries.
Replace batteries immediately if a battery warning is displayed. Boat-mounted
GPS units run off of the boat electrical system.
• Test and calibrate the multi-parameter meter (or sonde). Each field crew must
refer to and follow the manufacturer's calibration and maintenance procedures
to calibrate multi-para meter meters according to manufacturer specifications.
Once each week, crews must verify that the meter is functioning properly by
performing manufacturer recommended internal diagnostic readouts (e.g. pH
millivolts, cell constants, and/or other diagnostic readings). Records of these
checks should be saved in a logbook or other documentation. For those meters
that do not have internal check capabilities, crews will need to verify that the
meter is measuring pH and conductivity properly by measuring a commercially
available Quality Check Solution (QCS) with properties similar to YSI 5580
confidence solution.
• Ensure that the PAR meter's handheld display unit has fresh batteries, that the
unit is functioning properly, and that the correct calibration factors are
entered for each probe.
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Note: Calibration factors are supplied by the manufacturer and are specific to
each individual probe. PAR sensors require no field calibration; however, they
should be returned to the manufacturer at least every 2 years for calibration.
Field crews must use the "Procedures for the initial setup of the LI-COR Ll-
1400 Datalogger" (Section 4.2.1) to verify the setup of the unit or to enter
coefficient values should a new sensor need to be installed.
• Crews operating in the Great Lakes must ensure that the underwater video
system's battery is charged and all components are correctly connected. Crews
must ensure that the GPS attached to the video system is set up to output
information to the GPS overlay (Section 11.2.2). The GPS output will be set
prior to shipping to field crews, but the crews must verify proper settings
before use.
4.2.1 INITIAL ASSEMBLY AND SETUP PROCEDURES FOR LI-COR FRAME, SENSOR AND LI-1400
DATALOGGER
Field crews must use a pre-configured LI-COR system. Use the following instructions to
assemble the system if needed and the following section to reconfigure the LI-COR if
needed.
4.2.1.1 Assembly of the LI-COR lowering frame and sensor (from LI-COR 2006)
For NCCA, crews will need to attach one LI-192 Underwater Quantum Sensor to the LI-COR
lowering frame. IMPORTANT: Do not use LI-COR underwater cable to support the sensor
and lowering frame, as damage to the cable can result. The lowering line provided in your
base kit should be used to support the lowering frame and sensor by attaching the in-line
clip to the suspension ring at the top of the lowering frame. In addition, the cable should
not be bent sharply near the sensor.
The lowering frame provides for the placement of two cosine sensors, however, NCCA
crews will only attach a single underwater sensor. Each LI-COR underwater sensor has
three 6-32 tapped mounting holes on the underside of the sensor for connection to the
mounting ring (Figure 4.2). Corrosion resistant mounting screws are used with each
sensor.
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Insulating washer
Mounting ring
Screw (one of three)
Figure 4.2 Attachment of the underwater sensor to
the mounting rings (adapted from LI-COR, 2006)
Marked sounding line
(use in-lineclip)
.Undcrwatcrcable
Underwater sensor
n
Secchi disk clip
(not used for PAR
meter)
Weight (moderate)
Figure 4.3 Lowering frame assembly with sensor,
weight, and lowering line (adapted from LI-COR,
2006)
The underwater sensor will be attached using the mounting ring on the fin of the
lowering frame (Figure 4.3). To accommodate for any tilting of the frame and to ensure
a straight downward direction, a compact weight should be attached to the weight ring
at the bottom of the frame. Depending upon the speed of the current, moderate weights
will often suffice (4 kg). Weights over 25 kg should be avoided.
Once the sensor is installed to the mounting ring using the three screws and insulating
washer, plug the underwater cable into the sensor by aligning the sensor pins and
tightening the threaded connection. There is a yellow etched mark on the sensor bottom
that should be aligned with the raised nub on the cable. If the underwater sensor begins
reading negative values at startup, this likely indicates that the plug on the bottom of
the underwater sensor is plugged in backwards.
The underwater cable should be attached to the frame such that approximately 25 cm of
cable forms a smooth arc to the underwater sensor connector and is restrained from
being flexed or supporting any weight. Additionally, the cable must be securely attached
to the shaft of the lowering frame at multiple points so that the cable does not slip and
put strain on the sensor connector. However, the cable cannot be clamped so tightly as
to damage it. Possible methods to use are numerous nylon cable ties along the length of
the shaft, or a tight wrap of light cord around the shaft and cable, starting at the
suspension ring and extending downward at least 25 cm.
4.2.1.2 Setup Procedures for LI-COR LI-1400 Datalogger
The following example demonstrates the process for configuring the LI-1400 (with the
instrument keypad) to view or log instantaneous data from a single LI-190SA Quantum
Sensor.
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Example 1a. Configure channel 11 for a LI-COR LI-190SA Quantum Sensor with calibration
multiplier of -125.0umols-1m-2/uAmp (Each sensor has a unique multiplier value
supplied from the factory)
1.1. Connect the Quantum LI-190 ambient light sensor to the BNC connector on
top of the LI-1400 labeled 11.
1.2. Turn on the LI-1400 meter.
1.3. Press the [Setup] key.
1.4. Use the left ([<-]) or right ([^]) arrow keys to navigate to "SETUP
CHANNELS".
1.5. Press the [Enter] key to begin the sensor setup.
1.6. Use the left ([<-]) or right ([->]) arrow keys to navigate to "H=Light", press
Enter".
1.7. Using the [Shift] key and the number/ letter keys, type a description for
this channel. This description could describe the type of sensor (i.e.
"QUANTUM"), or describe what the reading will be used for in the NCCA
sampling (i.e. "AMB").
1.8. Press the down ([|]) arrow key to enter the multiplier. The multiplier value
is found on the Certificate of Calibration provided with the sensors. Each
sensor must have a unique certificate and calibration multiplier value.
1.9. Press the down ([|]) arrow key; enter "UW" for the unit label.
1.10. Press the down ([|]) arrow key; select "1 sec" to display instantaneous
values. The running average parameter will not be used, but could be set
here to any desired value.
1.11. Press the down ([|]) arrow key; select "Log Routin=none"
1.12. The remaining options do not need to be set as they apply only when
using a Log Routine.
1.13. Repeat this entire procedure for channel 12 to setup the underwater
sensor ("!2=Light").
4.3 EQUIPMENT AND SUPPLY PREPARATION
Field crews must check the inventory of forms, supplies, and equipment prior to
departure using Appendix A: Equipment and Supplies Lists; use of the lists is mandatory.
Inventory extra site kits prior to each site visit to ensure sufficient back-up supplies are
available. Store extra site kits in the vehicle so that replacement supplies will be readily
available in case of loss or damage while at the sampling site.
• Obtain sufficient wet and dry ice for sample preservation and storage.
• Pack meters, probes, and sampling gear, taking care to do so in a way that
minimizes physical shock and vibration during transport.
• Pack stock solutions as described in Table 4.1 below. Follow the regulations of
the Occupational Safety and Health Administration (OSHA).
Field crews must request paper field forms (if using), tracking forms and labels, and site
kits through the supply request form two weeks prior to sampling. Crews using the NARS
App must request tracking forms and labels for each site and may wish to request paper
form packets as a backup to electronic data collection. Field Crew Leaders collecting
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human health fish tissue samples in the Great Lakes must specifically request a human
health fish tissue supply kit through the supply request form. Field Crew Leaders MUST
provide a general schedule to the EPA and the Contractor Field Logistics Coordinator
two weeks prior to initiating sampling for the season.
Note: Site kits for all sites to be sampled in 2015 cannot be provided at the beginning of
the field season. Consequently, site kits must be sent out as requested throughout the
index period.
The site kit includes sample jars, bottles and other supplies (see complete list in
Appendix A: Equipment and Supplies Lists). After receipt, please inventory the site kit
against these lists. If items are missing, damaged or incorrect, please request
replacement supplies using the supply request form or by contacting the Contractor Field
Logistics Coordinator. The Contractor Field Logistics Coordinator will send replacement
supplies as quickly as possible.
Table 4.1 Stock solutions, uses ft methods for preparation
Bleach (1-10%)
Quality Check
Solution for multi-
parameter sonde
Buffered Formalin
Lugol's Solution
Clean nets, gear, and inside of boat
Weekly check of meter calibration
In place of weekly internal meter checks
Preserve benthic samples
Preserve phytoplankton samples
(Great Lakes sites only)
Add 10 - 100 mL bleach to 1 L distilled water.
No preparation needed (if purchased as ready-to-use
solution)
Add 8 tablespoons Borax to 2 gallons 100% Formalin (37%
formaldehyde) solution.
FOR USE AT ALL SITES: Add :/4 teaspoon Rose Bengal
crystals to above solution.
None (included in GL base kits); LugoPs Iodine solution is
light sensitive. Take care to avoid exposure to direct light.
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5 INITIAL SITE PROCEDURES
Upon arriving at the site, the field crew must confirm that it is the correct site and
determine if the site meets the criteria for sampling and data collection activities. The
crew verifies site access, safety, and general conditions to determine if the site can be
sampled within the swing of the anchored boat.
Note: Inability to collect samples for sediment, benthic or fish indicators does not
disqualify a site from meeting sample criteria. See Section 2.3.1 to determine site
sampleability.
5.1 SITE VERIFICATION
5.1.1 EQUIPMENT & SUPPLIES
Table 5.1 Equipment ft supplies: site verification
For locating and sampling permit and landowner access (if required)
verifying site site packet, including access information, site spreadsheet with map coordinates,
navigational charts with "X-site" marked
NCCA Fact Sheets for public outreach
GPS unit (preferably one capable of recording waypoints) with manual, reference card, extra
battery pack
For recording Site Verification form
measurements pencils (for data forms)
fine-tipped indelible markers (for labels)
clipboard
5.1.2 SITE VERIFICATION PROCEDURES
1. Create a waypoint in the GPS unit that corresponds to the target X-site
coordinates provided by EPA in the site list. This can be done ahead of time in
the office.
2. Navigate the sampling vessel as close as possible to the target X-site using GPS
(you must be no more than 0.02 nautical miles (NM) or 37 meters from the
target X-site). Compare the target X-site coordinates with the GPS coordinates
displayed at the sampling site.
• Sampling may start when the sampling vessel is within 37 meters of the X-
site. This distance provides the desired level of precision which is
approximately equal to that of the GPS receiver without differential fix
correction.
• With the exception of fish tissue and sediment samples (see Section 5.4)
crews are expected to collect all samples within a circle of 0.02 NM radius
around the X-site. This allowable deviation distance accounts for typical
"anchor swing" of the sampling vessel.
3. Anchor the sampling vessel in such a way as to minimize the possibility of the
anchor(s) dragging or becoming dislodged.
4. Once the anchor has been set and the vessel is essentially stationary, verify
that the X-site is still within 0.02 NM or 37 meters. This location (where
sampling will begin) is referred to as the Y-location. If the X-site is not within
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0.02 NM or 37 meters, reposition the vessel by following the steps outlined
above.
5. Determine if the site is sampleable. See Section 2.3.1 for specific guidelines.
• If not sampleable, proceed to Section 5.1.3.
• If sampleable, proceed to the steps below and then to Section 5.1.4.
Record the time of arrival to the Y-location on the Site Verification Form.
6. Record the coordinates of the Y-location on the Site Verification (Front) form
in decimal degrees in the NAD83 datum.
7. Record the number of satellites fixed as <3 or >4.
8. After anchoring, and throughout all subsequent sampling efforts, monitor the
GPS to ensure that the sampling vessel stays within the proper X-site radius.
9. Indicate any and all methods that were used to verify that you are at the
correct location.
10. Measure and record the water depth at the Y-location on the Site Verification
(Front) form. Make sure an accurate depth reading is taken at the site to
ensure the depth is adequate to conduct sampling.
5.1.3 SITE RELOCATION
Every attempt should be made to sample within a 0.02 NM (-37 m) radius of the X-site. If
the proposed initial sampling location is not sampleable, then relocate using the following
guidelines:
1. The Field Crew Leader should choose a specific compass heading (e.g., north,
south, east, west) and slowly motor the vessel in that direction.
2. After moving approximately 15-20 m, assess the relocated area using the Site
Verification guidelines given above.
3. Should the relocated area fail to meet the "sampleable" definition, then this
process may be continued using the same heading out to 37 meters from the X-
site.
4. If no suitable sampling location is found along the first chosen heading, return
to the X-site and follow a new heading until an acceptable sampling location is
found.
5. If after a sufficient amount of effort is expended and no suitable sampling
location is found, then the determination may be made that the site is
unsampleable.
6. If the site is non-sampleable or inaccessible and cannot be relocated within the
designated area, indicate the reason on the Site Verification (Front) form. No
further sampling activities are conducted.
7. Replace the original site with the next oversample site on the estuary/state
list.
• Notify the EPA Regional Coordinator and FLC that the site was replaced and
submit the Site & Sample Status/Water Chemistry Lab Tracking form to
NARS IM.
8. Return to Section 5.1.2.
5.1.4 SITE CHARACTERISTICS
1. If the site is sampleable, record the sampling status and method being used
(marine or Great Lakes).
2. Record the general habitat type and the dominant bottom type present at the
sampling site.
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• In many sites, it may not be possible to record the bottom type until after
the sediment collections are performed.
3. Record the presence and type of debris (if any), submerged aquatic vegetation
(SAV) present, and/or macroalgae present in the area.
4. Make any general comments about the site that may be important during the
data review portion of the assessment or any unusual characteristics about the
site, including weather conditions.
5. Record directions to the launch site from an easily recognizable location (city
or major road intersection).
6. On the back side of the Site Verification (Back) form, draw a simple sketch of
the area.
• Include the relative locations of the shoreline, launch point, X-site, Y-
location, and, if different from the Y-location, sediment and fish collection
locations. If sediment and fish were collected at different locations from
each other, please indicate them separately (see Section 5.4). Include any
other specific attributes of the site that may be important during data
analysis.
• A printed or copied section of a map with the pertinent information may be
submitted in place of the scene sketch. Upload this map to the NARS
SharePoint site when you submit your data forms. NARS App users will
upload their site sketch to the NARS SharePoint as well.
7. Record the name of all crew members and indicate their primary duties.
5.2 SITE PHOTOGRAPH
Although not required, EPA encourages crews to take site photographs, especially if the
site is associated with unusual natural or man-made features.
• Date-stamp any site photographs and include the site ID.
• Alternatively, start the photograph sequence with one image of an 8.5 x 11
inch piece of paper with the site ID, waterbody name, and date printed in
large, thick letters.
• Keep a brief photograph log (site ID, number of photographs, time and date
if not stamped by camera) and describe the subject of each photo ;/ it is
not self-explanatory.
• Field crews can upload these photos to the NARS SharePoint site.
5.3 SAMPLE COLLECTION
Even when the field crew makes every attempt to collect all samples, there will be some
circumstances that will prevent all samples from being collected. When site conditions
limit full completion of the standard sampling protocol, crews prioritize sample collection
and follow a "checklist" for determining the order of sample completion:
1. Measure in situ water parameters and collect all water samples at all sites.
2. Collect benthic grab samples at all sites. Any size sediment grab is acceptable
as long as it meets the definition of a "successful benthic grab" (see Section
12.3).
Note: Acceptable means:
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a) A sediment grab that meets the criteria for benthic samples; or
b) Enough sediment can be collected that will allow the crew to obtain the
surficial sub-sample required for the sediment composite to send to the
laboratory for abiotic indicator analysis (e.g., organics/metals, TOC, grain
size, toxicity).
3. Collect sediment composite material of sand-sized sediment grain or smaller
(preferred size). If an acceptable sediment grab cannot be obtained at the Y-
location or within a 37 m radius around the X-site, move to a secondary
sediment collection area following the procedures in Section 5.4.1 below. Flag
and note the reason for limited/missing sediment samples. In the case of
limited sediment, prioritize sample distribution in the following order of
preference:
a) Organics/Metals [SEDO]
b) Toxicity [SEDX]
c) Total Organic Carbon [SEDC]
d) Silt/Clay (Grain Size) [SEDG]
Indicate if any of the sediment samples were not successfully collected by
marking the "no sample collected" bubble(s) for each pertinent sample.
4. Collect fish for ecological contaminant [FTIS] analysis. For the ecological
assessment, fish collections are targeted to areas within a 500 m radius of the
X-site. After unsuccessful attempts within this area, crews may move outside
of this radius and attempt to collect fish up to 1000 meters from the X-site (see
Section 5.4.2). Unsuccessful deployment of fish collection gear or the absence
of fish in the catch should not necessarily be used as a determining factor for
rendering a site unsampleable.
5. Collect fish tissue plugs [FPLG].
6. Collect human health fish tissue sample [HTIS] (if applicable). If suitable fish
cannot be collected within 1000 meters of the X-site, crews may move out to a
maximum of 1500 meters from the X-site in an effort to collect the human
health fish tissue sample.
5.4 SECONDARY SEDIMENT OR FISH COLLECTION ZONES
All water, benthos, sediment, and fish samples are expected to be collected at the same
location (the Y-location), which is as close to the X-site as possible (within the 37 meter
radius around the X-site). However, circumstances may require the field crew to relocate
to a secondary location to collect an acceptable sediment grab or fish sample. If benthos,
sediment, and/or fish are collected from a secondary location, in situ measurements and
water collections do not need to be resampled. Guidelines for relocating to a secondary
sample collection zone are covered in the sections below.
5.4.1 SEDIMENT SAMPLES
1. If an acceptable sediment grab cannot be obtained at the Y-location where
water samples were collected, move the vessel within the 37 m radius margin
(of the X-site) and try to obtain the sediment sample. Use the site relocation
method described previously (Section 5.1.3). On the Sample Collection (Back)
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form, indicate the sediment collection zone by filling in the "within 37 m from
X-site" bubble.
2. In cases where sediment sampling cannot be successfully conducted within 37m
of the X-site, grabs may be taken in a secondary sediment collection zone
(e.g., > 37 m radius but within a 100 m radius (-0.05 NM) of the X-site) without
re-collecting the water samples (Figure 5.1).
Draw a second circle with a 100 m radius from the X-site on the site sketch
map. Place a mark on the map showing the relative location of the sediment
collection zone and the approximate distance and direction from the X-site.
Indicate in the comments section approximately how far and in what direction
from the X-site the sediment was collected. On the Sample Collection (Back)
form, indicate the sediment collection location by filling in the "between 37-
100m from X-site" bubble. The data will be flagged for subsequent review.
3. Crews may use the same relocation procedures to move out to a maximum
distance of 500 m from the X-site to locate suitable sediment sampling
locations (after attempting to collect sediment from within the primary and
secondary zones). Draw a 500 m radius circle on the site sketch map indicating
the sediment collection area and the approximate distance and direction from
the X-site. Indicate in the comments section approximately how far and in what
direction from the X-site the sediment was collected. On the Sample
Collection (Back) form, indicate the sediment collection zone by filling in the
"between 100-500m from X-site" bubble. The data will be flagged for
subsequent review.
4. If a suitable location to collect sediment samples has not been found after a
minimum of three collection attempts inside each of the acceptable relocation
radii, sediment sampling is considered "complete" for the site. All appropriate
field form flags and explanations must be completed, as well as pertinent "no
sample collected" bubbles.
Note: The Field Crew Leader may choose to make additional sediment grab
attempts.
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X-Site: Target sampling coordinates from the Site List
V-Location: Actual boat location after anchoring. Sample collection begins here.
(As close to X-site as possible; can be anywhere within 37 meters of the X-Site)
37m 100m 500m
Primary Sample Collection Zone:
0-37 meters from X-site
• In Situ
• Water
• Benthos (if possible)
• Sediment (if possible)
Secondary Sample Collection Zone:
37 - 100 meters from X-site
If sediment not available in primary zone
• Benthos
• Sediment
! Tertiary Sample Collection Zone:
! 100 - 500 meters from X-site
] If sediment not available in primary or secondary zones
] Crews may move out to a maximum
', distance of 500 meters from the X site < ;
I I
i in repeated attempts to locate suitable i [
! benthos/sediment sampling locations
Figure 5.1 Primary, secondary and tertiary sample collection zones
5.4.2 FISH SAMPLES
Secondary fish tissue collection sites may be selected up to an additional 500 m beyond the
original 500 m radius at all estuarine and Great Lakes sites (Figure 5.2).
Please observe the following guidelines:
1. In order to move to a secondary fish tissue collection site, crews must be
unsuccessful at obtaining target fish during a reasonable portion of the three
hours allotted to fishing (at least 30 minutes and no more than two hours)
within the original 500 m radius.
2. The crew must have attempted several sampling locations within the 500 m
radius without success.
3. Crews must observe signs of fish presence such as schools of bait fish just
below the surface, predator activity or prey escape behavior on the surface of
the water, overhead shading or favorable underwater habitat structure or
bathymetric features within an additional 500 m from the X-site.
4. When relocating outside of the original 500 m radius from the X-site, but inside
of the 1000 m radius of the X-site, crews must document:
a) The amount of time spent fishing within the original 500-meter radius.
b) The direction of travel from the X-site.
c) The coordinates of the site where fish were ultimately caught.
5. For the collection of the human health fish tissue sample ONLY, crews may
move out to a maximum of 1500 meters from the X-site.
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500m 1000m 1500m
Primary Fish Collection Zone (FTIS/FPLG/HTIS):
0 - 500 meters from X-site
• Spend at least 30 minutes attempting to collect
fish here, but no more than 2 hours is required
• Attempt fishing in several locations within
primary zone.
• If no suitable fish are collected,
consider moving to secondary zone
Secondary Fish Collection Zone (FTIS/FPLG/HTIS):
500 - 1000 meters from X-site
// no suitable fish are collected in primary zone, and
• Crew observes signs of fish or
favorable habitat/structure
Human Health Fish Tissue (HTIS1 ONLY: \" "
1000 - 1500 meters from X-site ; !
// no suitable human health fish are ' •
collected in primary or secondary zones, crews
may move out to a distance of 1500 meters from
the X-site in an attempt to collect this sample.
Figure 5.2 Primary and secondary fish collection zones
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6 WATER QUALITY MEASUREMENTS
This section describes the procedures and methods for the field collection and analysis of
the water quality indicators (in situ measurements, water column transparency, and light
attenuation) from freshwater and marine coastal areas.
6.1 SUMMARY OF METHOD FOR IN SITU MEASUREMENTS OF WATER COLUMN
TRANSPARENCY, DISSOLVED OXYGEN, pH, SALINITY, CONDUCTIVITY,
TEMPERATURE, AND LIGHT ATTENUATION
Field crews obtain a hydrographic profile at each site (at the Y-location) by measuring DO,
pH, salinity (marine sites) or conductivity (freshwater sites), and temperature using a
multi-parameter water quality meter (or sonde). They also assess water column
transparency using a Secchi disk and light attenuation using a PAR meter. The protocol
requires measurements at the prescribed depths as the probe/sensor is both lowered and
retrieved, starting just below the surface, progressing down to 0.5 m from the bottom,
and returning to just below the surface.
6.1.1 EQUIPMENT AND SUPPLIES
Table 6.1 lists the equipment and supplies used to measure water column transparency,
DO, pH, salinity/conductivity, temperature, and light attenuation. Crews record in situ
measurements on the Field Measurement (Front) form.
Table 6.1 Equipment ft supplies: transparency, DO, pH, salinity/conductivity, temperature, ft light
attenuation
For taking measurements and I multi-parameter water quality meter with DO, pH, salinity/conductivity, I
calibrating the water quality meter | and temperature probes. |
I extra batteries I
I de-ionized water (lab certified preferred, but not required) I
I calibration cups and standards I
I QCS (used if internal meter checks an not possible) I
| barometer to use for calibration |
| thermometer |
j Secchi disk (20 cm diameter, weighted) & 100' line with clip (marked in |
I 0.5 m intervals) I
| PAR meter (with LI-190 Quantum Sensor and LI-192 Underwater I
| Quantum Sensor & cables, independent datalogger) j
For recording measurements I Field Measurement form ;
I pencils (for data forms) :
6.2 SAMPLING PROCEDURE - WATER COLUMN TRANSPARENCY (SECCHI
DEPTH)
A Secchi disk is a 20 cm black and white disk suspended from a non-stretch line that is
marked in 0.5 m intervals. Field crews use a Secchi disk to measure water column to
nearest 0.1m transparency at every site (at the Y-location). The resulting measurement is
called the Secchi disk transparency depth, or "Secchi depth" for short. Below are step-by-
step procedures for measuring water column transparency.
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Note: For valid Secchi depth readings, no sunglasses, hats, or any other devices that
shade the eyes may be used by the person who is observing the disappearance and
reappearance depths. The Secchi depth is assessed from the shady side of the boat and
can only be measured during daylight hours. One crew member must make all three sets
of Secchi measurements at a site, and it is desirable to have the same crew member
complete Secchi depth readings throughout the entire field season whenever possible.
1. In the "Secchi Depth" section of the Field Measurement (Front) form, record
the time Secchi depth readings were started.
2. Slowly lower the Secchi disk until it is no longer visible. In the "DISAPPEARS"
column, record the depth where the marking on the line meets the water level.
Interpolate between the 0.5m markings on the rope to the nearest 0.1m.
• If the disk hits the bottom before disappearing, water column transparency
depth is greater than the water depth. Fill in the "Yes" circle on the data
sheet under "Clear to Bottom?" and record the station depth as both the
disappearance and reappearance depth in the "Reading 1" row on the data
form.
3. Slowly raise the Secchi disk until it just becomes visible and record the depth
in the "REAPPEARS" column. Interpolate between the 0.5m markings on the
rope to the nearest 0.1m.
4. Repeat steps 1 -3 two more times, recording both disappearance and
reappearance depths each time.
5. Use the comment space provided on the Field Measurement (Front) form to
flag any measurements that the crew feels needs further comment or when a
measurement cannot be made.
6. Repeat the entire process if any one disappearance or reappearance
measurement differs from the others by more than 0.5 m.
6.3 SAMPLING PROCEDURE - MULTI-PARAMETER SONDE
6.3.1 CALIBRATION
Crews calibrate the DO, pH, and salinity/conductivity meter functions of the multi-
parameter water quality meter (or sonde) before collecting data at each site. If a crew is
sampling multiple sites in a single day, a single calibration is sufficient for the day.
• Crews record the manufacturer and model number of the instruments in the
"Calibration Information" section of the Field Measurement (Front) form.
• Crews must calibrate their pH probe according to the manufacturer's instructions
and their own laboratory policies, by using at least a 2-point calibration method.
Crews will supply commercially purchased calibration standards (typically pH of 7
and 10 for 2-point calibration and pH of 4, 7, and 10 for 3-point calibration). Any
pH standards used must reference NIST Standard Reference Material (SRM)
certifications to be used in the calibration of the pH probe. This applies for
calibrations done both pre-sampling and post-sampling.
• The calibration buffers must be accurate to 0.02 pH units or better.
• The calibration buffers should be replaced with fresh solutions every 3-4
days or sooner if the crew suspects it has become contaminated
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• Crews will also calibrate their conductivity/salinity probe according to the
manufacturer's specifications and their own laboratory policies, using a
commercially supplied, traceable conductivity standard.
• Crews will re-check pH and conductivity/salinity calibration again after daily
measurements are complete to document potential meter drift throughout the
day.
• For instruments that are factory calibrated and checked (e.g. Sea-Bird Electronics
meters, etc.), crews must ensure that factory-certified diagnostics have been
completed according to manufacturer specifications (preferably conducted
immediately prior to the sampling season) and provide documentation copies
during assistance visits. Meters such as these do not require the daily calibration
steps or the weekly diagnostic/QCS checks.
• Once each week, crews must verify that the meter is functioning properly by
performing manufacturer recommended internal diagnostic checks. This is
manufacturer and model specific, but typically involves accessing internal
diagnostic readouts (e.g. pH millivolts, cell constants, and/or other diagnostic
readings). Results of these checks must be recorded in a logbook or other
documentation and saved for potential review.
• For those meters that do not have internal check capabilities, crews will check pH
and conductivity against a commercially available QCS with properties similar to
YSI 5580 confidence solution. The QCS is provided by the crew. Crews record the
successful completion of the internal checks or the expected values and measured
values of the QCS in the "Quality Control Check" section of the Field
Measurement (Front) form.
• Crews using a commercially purchased pH QCS for the weekly quality checks should
follow the guidelines below:
• The pH QCS containers should be labeled with expected values and
preparation dates.
• The pH of the QCS should approximate the pH expected at sampling sites.
• Crews should have centrally located bulk solutions to replenish allotments
needed for quality checks every 3-4 days or sooner if the crew suspects it
has become contaminated.
o Bulk solutions should be replaced according to the manufacturer's
specifications or at any time if crew suspects it has become
contaminated.
• Crews use a commercially purchased primary conductivity/seawater standards to
be used as the QCS for weekly quality checks of conductivity/salinity.
• A secondary conductivity/seawater standard that is referenced against a
certified standard may also be used.
o If a secondary standard is used, then preparation and certification
test procedures and results must be logged in a QA notebook and
maintained by the state or contractor in-house QA personnel.
o The standard should be representative of the conditions expected in
the field (-0.5-35 ppt for marine waters).
o The conductivity/seawater calibration standard and QCS containers
must be labeled with expected values and preparation dates.
• The standards should be replaced with fresh solutions every 3-4 days or
sooner if the crew suspects they have become contaminated.
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o Bulk supplies of calibration standards and primary or secondary QCS
may be maintained in a central location and used to replenish QA
allotments.
o Bulk solutions should be replaced according to manufacturer's
specifications or if the crew suspects that they may have become
contaminated.
• At least once per sampling season (usually in a laboratory before crews begin
sampling), calibrate the temperature sensor against a National Institute of
Standards and Technology (NIST)-traceable thermometer.
• If you observe any irregularities or calibration measurements that fall outside of
the specified tolerance ranges use an alternate instrument if available and flag any
affected data.
Specific information about calibrating each probe function is presented below.
6.3.2 DISSOLVED OXYGEN METER
Calibrate the DO probe in the field against an atmospheric standard (i.e. ambient air
saturated with water or water saturated with air) according to manufacturer's
specifications and NCCA QA protocols prior to launching the boat. In addition, follow any
of the manufacturer's recommendations for periodic comparisons with internal quality
checks (cell constants, millivolt output, or other readings), or a DO chemical analysis
procedure (e.g., Winkler titration) to check accuracy and linearity. Record results and
report irregularities as described above.
6.3.3 PH METER
Calibrate the pH meter in accordance with the manufacturer's instructions and with the
field crew organization's existing Standard Operating Procedure (SOP).
After all in situ measurements have been completed for the sampling day, crews perform
a post-measurement calibration check of the pH meter. Crews will record the Calibration
Standard Value pH and the post-sampling measurement in the appropriate locations in the
"Post-Measurement Calibration Check" section of the Field Measurement (Front) form. If
significant drift (outside of manufacturer's specification) is detected, it may indicate that
the meter is in need of service. Perform the required service or exchange devices as
appropriate and if necessary, and flag any suspect measurements. Discontinue use of any
meter that is not functioning properly.
Once a week, each crew must check their multi-parameter sonde using manufacturer
recommended internal diagnostic checks (cell constants, millivolt output, or other
readings) or against the QCS that they provide. In addition to recording the expected
values and results, record the QCS date prepared in the appropriate sections of the
"Quality Control Check" section of the Field Measurement (Front) form. Report any
calibration or QC irregularities as described above.
6.3.4 SALINITY/CONDUCTIVITY METER
Prior to sampling each site, calibrate the salinity/conductivity meter in accordance with
the manufacturer's instructions. After the sampling day is complete, measure the
salinity/conductivity of the calibration standard that was used earlier in the day to
calibrate the instrument. Record the expected and post-measurement values as described
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above. Once a week, crews check the conductivity/salinity function using manufacturer
recommended internal diagnostic checks (cell constants, millivolt output, or other
readings) or against the QCS that they provide. Record results and report irregularities as
described above.
6.3.5 TEMPERATURE METER
When performing the once-a-season temperature sensor check, incorporate the entire
temperature range encountered in the NCCA into the testing procedure and keep a record
of test results on file. For use in this accuracy check, the following are the temperature
ranges from the NCCA 2010 dataset:
• Northeast: 6.8 °C < T < 32.3 °C
• Southeast: 21.2 °C
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Table 6.2 Example depth measurement intervals
EXAMPLE 1: EXAMPLE 2:
Water Depth = 7.2 meters Water Depth = 23.9 meters
0.1 m 0.1 m
0.5 m 0.5 m
1.0m 1.0m
2.0 m 2.0 m
3.0 m 3.0 m
4.0 m 4.0 m
5.0 m 5.0 m
6.0 m 6.0 m
6.7 m 7.0 m
8.0m
9.0m
10.0m
15.0m
20.0m
23.4m
6.5 PHOTOSYNTHETICALLY ACTIVE RADIATION (PAR) METER
Field crews measure photosynthetically active radiation using a PAR meter attached to a
LI-COR® data logger. The PAR meter measures a vertical profile of light attenuation at
each station. Measured light values are entered into a regression equation and used to
determine the coefficient of attenuation in the water column. PAR sensors require no
field calibration; however, they should be returned to the manufacturer at least every 2
years for calibration. Crews measure PAR at the same depths as other water column
indicators. See procedures below for measuring light attenuation.
6.5.1 SAMPLING PROCEDURE—LIGHT ATTENUATION (LI-1400 DATALOGGER)
1. Connect a deck sensor (LI-190 Quantum Sensor) and an underwater sensor (Ll-
192 Underwater Quantum Sensor) to the PAR meter as described in Section
4.2.1. Enter the calibration factors (supplied by the manufacturer) for each
probe.
2. Place the deck sensor in an unshaded location on the boat to record the
available ambient light.
3. Turn on the LI-1400 meter.
4. Press the View key.
5. Using the left or right keys, navigate to "NEW DATA" and press Enter.
6. Using the left or right keys, navigate until channel 111 is displayed; this shows
the instantaneous reading for that channel. Scrolling down will allow viewing of
2 channels at once.
7. Lower the underwater sensor, making sure that the sensor is facing up, on the
SUNNY (or at least unshaded) side of the boat to a depth of 10 cm (represents
"surface"). Allow the readings to stabilize and press "Enter" to manually log
the ambient (AMB) and underwater (UW) light readings in the datalogger.
NOTE: crews may choose to use alternate methods of recording the two sensor
readings as long as both readings are recorded at the same instant. This may
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include using two people to view the two readings, taking a photograph of the
screen, etc.
8. Continue to lower the underwater sensor to each of the required depths (same
as other water quality measurements):
a) 0.5 m
b) Every 1 m from 1.0 m to 10.0 m
c) Every 5 m thereafter for sites greater than 10m
d) 0.5 m from the bottom
9. Allow the readings to stabilize at each depth before pressing "Enter" or
recording the values on the data form.
10. Repeat the procedure at the same depths, but in reverse order on the upcast.
11. Review the saved data by pressing Esc and using the right or left key to select
"LOG DATA" and pressing Enter.
12. Select "View=ALL." Press Enter.
13. Use the down key to scroll through stored data by date and time to find the
data that were just logged. Press Enter to access logged data. Use the down
key to view both of the sensor readings.
14. Record the values from both sensors (/jE/m2/s), at the appropriate water
depths of the underwater sensor, on the datasheet. Record the deck sensor
reading in the ambient (AMB) column, and the underwater sensor reading in the
underwater (UW) column.
15. If the sensor hits bottom, allow 2-3 minutes for the disturbance to settle before
taking the reading.
16. If the light measurements become negative before reaching the bottom
measurement, terminate the profile at that depth.
17. If the underwater sensor begins reading negative values at startup, this likely
indicates that the plug on the bottom of the underwater sensor is plugged in
backwards. There is a yellow etched mark on the sensor bottom that should be
aligned with the raised nub on the cable.
Note: Pressing the On/Off key will only turn off the screen. To shut down the LI-1400
press the Fct key and use the right or left keys to navigate to "SHUTDOWN". Press
Enter to shut down.
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7 WATER CHEMISTRY [CHEM], CHLOROPHYLL-^ [CHLA],
AND NUTRIENTS [NUTS] SAMPLE COLLECTION AND
PRESERVATION
This section describes the procedures and methods for the field collection and
preservation of the water chemistry, chlorophyll-a, and dissolved nutrients samples from
freshwater and marine coastal areas.
7.1 SUMMARY OF METHOD
The water chemistry samples will be analyzed for chlorophyll-a [CHLA], total nutrients
including nitrogen and phosphorus [CHEM], and dissolved ammonia, nitrites, nitrates, and
phosphorus [NUTS]. Collect the water samples at the Y-location, 0.5 meters below the
surface (or mid-depth if station depth is less than 1.0 meter), with either a water
pumping system or water sampling device such as a Niskin, Van Dorn, or Kemmerer bottle
and transfer to a rinsed 250 ml amber HOPE bottle. Water for the chlorophyll-a sample
will be collected and transferred to a separate 2 L amber HOPE bottle. Store all samples
in darkness on ice in a closed cooler. After you filter the chlorophyll-a sample, the filter
must be kept frozen until ready to ship. A portion of the filtrate from the chlorophyll-a
processing will be collected for the dissolved nutrient sample.
Note: Fecal indicator sample IS NOT collected with these samples.
7.2 EQUIPMENT AND SUPPLIES
Table 7.1 Equipment & supplies: water chemistry & chlorophyll-a sample collection
For collecting samples | water sampling device or water pumping system |
I nitrile gloves I
| HDPE bottle (250 mL, amber) [CHEM] |
; HDPE bottle (2 L, amber) [CHLA] i
: cooler with wet ice :
For recording ; Sample Collection form j
measurements | water chemistry sample label |
| pencils (for data forms) |
I fine-tipped indelible markers (for labels) I
| clear tape strips |
7.3 SAMPLING PROCEDURE
The following describes the sampling procedures for collecting water chemistry samples.
Note: Do not apply sunscreen or other chemical contaminants until after the sample is
collected (or implement measures to reduce contamination by such chemicals if applied
such as washing, wearing long gloves, etc.).
1. Collect the water chemistry samples at the Y-location, which is no more than
37 meters from the X-site (located via GPS).
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2. Complete the CHEM sample label with Site ID, date collected, and visit
number.
3. Attach the completed label to the 250 ml amber HOPE sample bottle and cover
with clear plastic tape.
4. Put on nitrile gloves.
5. Using either a water sampling device or water pumping system, collect a water
sample at 0.5 m below the surface (or mid-depth if station depth is less than
1.0 meter).
a) Rinse the sampling device and the sample containers three times with
water from the site. To rinse a pumped sampling system follow your
agency's SOP. If no SOP exists, flush long enough so that the amount of
site water flushed is equal to at least three times the total volume of
the sampling system (including tubing). Be sure to cap the bottles and
rotate them so that the water contacts all the surfaces. Discard the
water away from the sampling location if additional water is to be
collected.
6. Fill the 250 ml amber HOPE bottle (for water chemistry) and the 2 L amber
HOPE bottle (for chlorophyll-a and nutrients) with sample water.
7. Replace the lids and seal the lid of the 250 ml bottle tightly with electrical
tape.
8. Place both samples in a cooler on ice at 4°C.
9. Record the collection data on the Sample Collection (Front) form.
a) Note anything that could influence sample chemistry (heavy rain,
potential contaminants, etc.) in the Comments section.
b) If the samples were not taken at the Y-location, enter the GPS
coordinates of the sampling location and the reason for relocation in
the comments field on the Sample Collection (Front) form.
10. Proceed to Section 14.3 for instructions on processing chlorophyll-a and
nutrients water sample to obtain a chlorophyll-a filter and the nutrients
filtrate.
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8 ALGAL TOXINS [ALGX] INCLUDING MICROCYSTINS
[MICX]
Algae, including Microcystis, are microscopic organisms found naturally at low
concentrations in water. Under optimal conditions (such as high light and calm weather,
usually in summer), these organisms occasionally form a bloom, or dense aggregation of
cells, that floats on the surface of the water forming a thick layer or "mat." At higher
concentrations, algal blooms are so dense that they resemble bright green paint that has
been spilled in the water. These blooms potentially affect water quality as well as human
health (some algae produce toxins) and natural resources. Decomposition of large blooms
can lower the concentration of dissolved oxygen in the water, resulting in hypoxia (low
oxygen) or anoxia (no oxygen). Sometimes, this condition results in fish kills. The blooms
can also be unsightly, often floating at the surface in a layer of decaying, odiferous,
gelatinous scum.
Although the likelihood of people being affected by algal blooms is low, various health
effects can occur following contact with or ingestion of algal toxins. People recreationally
exposed (e.g., swimmers or personal watercraft operators) to algal blooms have also
reported adverse effects. Health problems may occur in animals if they are chronically
exposed to water with algal toxins present. Fish and bird mortalities have been reported
in waterbodies with persistent algal blooms.
8.1 SUMMARY OF METHOD
Two water samples for algal toxin analysis are taken from the Y-location: one for a broad
suite of algal toxins [ALGX] and another specifically for microcystins [MICX]. All field
crews must collect water grab samples using the water chemistry sample collection device
to fill two, 500 ml bottles. Collect these samples after the in situ measurements and
water chemistry sample are collected. Store all samples on ice in a closed cooler.
8.2 EQUIPMENT AND SUPPLIES
Table 8.1 Equipment & supplies: algal toxins, microcystins
For collecting samples nitrile gloves
water chemistry sample collection device
2 HDPE bottles (500 mL, white, wide-mouth) [ALGX] [MICX]
cooler with ice
For recording Sample Collection form
measurements algal toxin sample label
microcystin sample label
pencils (for data forms)
fine-tipped indelible markers (for labels)
clear tape strips
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8.3 SAMPLING PROCEDURE
See below for step-by-step procedures for collecting both algal toxins and microcystins
samples. Collect both samples from the Y-location.
Note: Make sure not to handle sunscreen or other chemical contaminants until after the
sample is collected (or implement measures to reduce contamination by such chemicals if
applied such as washing, wearing long gloves, etc.).
8.3.1 SAMPLE COLLECTION
1. Complete the ALGX and MICX sample labels with Site ID, date collected, and
visit number.
2. At marine sites, also write the salinity (in ppt) on both of the labels.
3. Attach the completed labels to each of the 500 ml HOPE sample bottles and
cover with clear plastic tape.
4. Put on nitrile gloves.
5. Rinse the first 500 ml bottle three times with site water. Be sure to cap the
bottle and rotate it so that the water contacts all the surfaces. Discard the
water away from the sampling location if additional water is to be collected.
6. Fill the 500 ml bottle. Leave at least one inch of head space in the bottle to
allow for expansion when frozen.
7. Replace the lid and seal tightly with electrical tape.
8. Repeat Steps 5-7 for the second 500 ml bottle.
8.3.2 SAMPLE STORAGE
1. Place the 500 ml bottles in a cooler (on ice) and shut the lid.
2. Record the Sample ID on the Sample Collection (Front) form along with the
pertinent site information (site ID, date, etc.).
3. As soon as you return to your base site (hotel, lab, office, etc.), freeze sample
bottles and keep frozen until shipping.
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9 FECAL INDICATOR (ENTEROCOCCI, [ENTE])
Crews collect water samples to be tested for the presence of Enterococci. They filter
water at the field site or a nearby location. The filters are sent to the lab for quantitative
polymerase chain reaction (qPCR) analysis. Two filters must be collected and frozen
within six hours of collecting the water sample or the sample must be discarded and
recollected. Because of the time-sensitive nature of this technique, the position of the
Enterococci water sample collection in the sampling sequence varies based upon whether
and how fish will be collected at the site and how quickly the crew will be able to begin
filtration.
In short, if the crew is using a passive fishing method or is able to filter the samples on
the vessel, the Enterococci collection takes place immediately following the hydrographic
profile. If the crew is using active fishing methods or will not be able to filter the sample
until off the water, the collection of the Enterococci sample takes place at the end of the
sampling day. This variation is based on balancing the need to protect the Enterococci
sample from potential contamination with minimizing holding times once the sample is
collected.
9.1 SUMMARY OF METHOD
Crews collect and preserve the fecal indicator sample at the Y-location using the method
described in the Sampling Procedure (Section 9.3 below). In addition, crews observe the
area around the X-site and record (on the Site Assessment (Front) form) signs of
disturbance that may contribute to the presence of fecal contamination to the waterbody.
9.2 EQUIPMENT AND SUPPLIES
Table 9.1 Equipment ft supplies: fecal indicator (Enterococci) sampling
For collecting samples nitrile gloves
HDPE bottle (250 mL, clear, pre-sterilized)
sodium thiosulfate tablet
wet ice
cooler
For recording measurements Sample Collection form
fecal indicator sample labels (2 vial labels and 1 bag label)
pencils (for data forms)
fine-tipped indelible markers (for labels)
clear tape strips
9.3 SAMPLING PROCEDURE
The following outlines the procedure for collecting the fecal indicator sample.
1. Put on nitrile gloves.
2. Using either a gloved hand (on smaller boats) or pole dipper (on larger vessels),
lower the un-capped, inverted 250 ml sample bottle to a depth of 0.5 meters
below the water surface (or mid-depth if station depth is less than 1.0 meter).
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• Avoid surface scum, vegetation, and substrates. Point the mouth of the
container away from the boat. Right the bottle and raise it through the water
column, allowing bottle to fill completely.
3. After removing the container from the water, discard a small portion of the
sample to allow for proper mixing before filtering.
4. Add the sodium thiosulfate tablet, cap, and shake the bottle 25 times.
5. Store the sample in a cooler on wet ice to chill (not freeze) for at least 15
minutes. Do not hold samples longer than six hours before filtration and
freezing.
6. The filtration procedure is contained in Section 14.2.
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10 PHYTOPLANKTON [PHYT] (GREATLAKES ONLY)
10.1 SUMMARY OF METHOD
In all Great Lakes sites, crews will collect a sample for phytoplankton analysis. Collect this
sample from the Y-location at the same time and depth as the other water samples. Fill a
1 L white narrow-mouth HOPE bottle with water from the water sampling device or water
pumping system. The phytoplankton sample must be preserved with Lugol's solution
within two hours of collection. Store the samples in darkness inside a cooler with ice or in
a refrigerator.
10.2 EQUIPMENT AND SUPPLIES
Table 10.1 Equipment & supplies: phytoplankton
For collecting and water sampling device or water pumping system
preserving samples nitrile gloves
HDPE bottle (1 L, white, narrow mouth)
wet ice
cooler
LugoPs solution
For recording Sample Collection form
measurements phytoplankton sample label
pencils (for data forms)
fine-tipped indelible markers (for labels)
clear tape strips
10.3 SAMPLING PROCEDURE
The text below describes the sampling and preservation procedures for phytoplankton
samples. Collect the phytoplankton water sample at the Y-location along with the other
water samples.
Note: Make sure not to apply sunscreen or other chemical contaminants until after the
sample is collected (or implement measures to reduce contamination by such chemicals if
applied such as washing, wearing long gloves, etc.).
1. Complete the PHYT sample label with Site ID, date collected, and visit number.
2. Attach the completed label to the 1 L white narrow-mouth HDPE sample bottle
and cover with clear plastic tape.
3. Put on nitrile gloves.
4. Using either a pre-rinsed pump system or a water sampling device, collect a
water sample at 0.5 m below the surface (or mid-depth if station depth is less
than 1.0 meter).
5. Rinse the sample bottle three times with site water. Be sure to cap the bottle
and rotate it so that the water contacts all the surfaces. Discard the water
away from the sampling location if additional water is to be collected.
6. Fill the sample bottle with sample water, leaving enough head space for 10 ml
of Lugol's solution, and place in a cooler on ice at 4°C. Store the sample
chilled and in darkness at all times.
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7. The sample must be preserved by adding 10 ml of Lugol's solution to the bottle
within 2 hours of collection.
8. After preservation, replace the lid and seal tightly with electrical tape.
9. Record the collection data on the Sample Collection (Front) form. Include the
depth of collection, time of collection, and time of preservation.
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11 UNDERWATER VIDEO [UVID] (GREATLAKES ONLY)
At all Great Lakes sites, a 1 minute video image of the substrate at the Y-location will be
collected using an underwater video camera system. The video will be enhanced and
examined in the lab to visually document the bottom composition, and record the
presence or absence of zebra mussels, Cladophora, or other organisms.
11.1 SUMMARY OF METHOD
High quality underwater video will be best achieved if the field crew deploys the camera
and records the video at approximately the same time as the in situ measurements and
water collection activities. Avoid heavy disturbance of the bottom with anchors or
sediment samplers before capturing the video images.
At the Y-location, lower the camera into the water on the windward side of the boat and
wait for a clear view of the bottom. Record until you have captured 1 min of good bottom
footage.
11.1.1 EQUIPMENT AND SUPPLIES
Table 11.1 Equipment & supplies: underwater video
For recording Seaviewer underwater camera
underwater video Seaviewer digital video recorder (DVR)
Seaviewer SeaTrak GPS overlay
Garmin Etrex GPS
camera cable (100')
cable from GPS overlay to DVR
cable from GPS overlay to GPS
12v 18ah battery
charger for 12v battery
power cord (DVR ,Camera ,GPS overlay)
power adapters (110VAC - 12VDQ (3) for camera, DVR, and GPS overlay
EPA provided USB flash drive
stop watch (or similar time keeping device)
Seaviewer case (all components will fit into case for transport)
10 amp fuses (Automotive blade (large) type)
AA batteries (for GPS)
For recording Sample Collection Form
measurements pencils (for data forms)
11.2 SAMPLING PROCEDURE
11.2.1 INITIAL SETUP OF UNDERWATER CAMERA SYSTEM
Underwater camera systems will be assembled and set up prior to shipment to field crews.
However, information contained within this section will allow a field crew to verify
equipment setup or troubleshoot potential connection problems. The underwater camera
system and cables should be set up as shown in Figure 11.1. The system should not be
disassembled between sites other than to remove the battery clips. Initial one-time setup of
the camera system GPS unit is described below.
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cable
o button record button
| an/off button
DVR
Figure 11.1 Setup diagram of underwater video system
11.2.2 INITIAL SETUP OF UNDERWATER CAMERA SYSTEM GPS
1. Set the GPS to output NMEA data in the GPS Menu section of the settings to
send position information to the GPS overlay system. (This step will be
completed prior to shipment of the system, but the steps below can be
completed to verify correct setup).
2. Press "page" button 4 times to reach menu page.
3. Select "set up" by pressing arrow down button until "set up" is highlighted,
then press enter.
4. Select "interface" by pressing arrow down button until "interface" is
highlighted, then press enter.
5. Press enter again, select "NMEA out" by pressing arrow down button until
"NMEA out" is highlighted, then press enter.
6. Press page button 3 times to return to satellite tracking page.
11.2.3 UNDERWATER VIDEO RECORDING PROCEDURE
The following describes the procedure for recording the underwater video at the Y-
location as well as the procedure for archiving the video file after the recording has been
completed.
1. Power up the GPS and wait until it displays: "ready to navigate".
2. If the GPS is not set to output NMEA data, see Section 11.2.2.
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3. Connect either the battery wire to the internal battery or attach the alligator
clips to an external 12v battery. Attach red clip to the positive (+) terminal and
black clip to the negative (-) terminal.
4. Send power to the camera, GPS overlay, and DVR by switching the battery
switch to "Battery" (I) if using the internal battery or "External Power" (II) if
using an external 12v battery.
5. Turn on the GPS overlay by pressing its power button. The green light will
illuminate.
6. Turn on the DVR by pressing the power button (upper right side of DVR) for 3
seconds. A flash screen will appear for a short time then disappear.
Note: DO NOT PRESS POWER ON AGAIN DURING THIS TIME! A windows type menu
screen will appear.
7. Initialize the DVR by pressing the video ( -Q ) button located at top right of
the DVR unit (not on the screen). A video image from the camera should appear
on the DVR screen with a latitude / longitude display. (If not, see the trouble
shooting section supplied with manufacturer's literature).
8. While the camera is still out of the water, start recording by pressing the
record button (located at the top right of the unit, to the right of the video
button). The word "Recording" appears on the screen in red for 10 sec, then
disappears. (To pause recording, press the enter button then press it again to
resume recording).
9. Once recording, lower the camera into the water on the windward side of the
boat. One person is needed to operate the DVR and one to lower the camera.
The person operating the DVR should instruct the camera person on descent
speed and depth of camera.
10. Once a clear and close-up image of the bottom is displayed (do the best you
can in the conditions), record an additional 1 minute of good bottom footage.
During this 1 minute recording, slowly rotate the camera 360 degrees while
maintaining a clear image of the bottom.
11. In low light conditions, turn on the camera light by pressing the red button on
the DVR end of camera cable. Experiment with the light while monitoring the
screen for best picture results.
12. Stop recording by pressing the video key (top of unit), or the X button.
11.2.4 REVIEWING UNDERWATER VIDEO FILES PROCEDURE
Upon completing the 1 minute of underwater video, it is important to verify that the
video has been saved, record the file name on the Sample Collection (Front) form, and
preview the video to ensure adequate quality.
1. Select browser by pressing the enter button (upper right key on front of DVR).
2. Arrow down to "DVR".
3. Select "DVR" by pressing the enter button (upper right key front of DVR).
4. Arrow down to the last file listed. This should be the video you just recorded.
5. Record the file name on the Sample Collection (Front) form. The format of the
file is: DVRyymmdd_hhmm_xxx.avi (yymmdd is the date in year, month and
day; hhmm is the time in hours and minutes; xxx is a file number assigned by
the DVR, typically 001; and avi is the file format). Check that the date and
time on the file name match the date and time of the recording you just made.
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6. Press enter to play video to evaluate the quality of the video.
7. If the video clearly shows the composition of the bottom then the video is
deemed acceptable; continue to step 9.
8. If the video is un-viewable or is of poor quality, repeat the recording steps
above.
9. Shut down the system by the following the steps below.
a) Power down the DVR by pressing and holding the power button (upper right
side).
b) Power down the GPS overlay by pressing the power button.
c) Power down the camera by disconnecting the alligator clips from the
battery posts.
d) Power down GPS by pressing and holding its power button.
10. Recharge the 12v battery at the end of each day (it is a good idea to assign this
task to an individual crew member).
11.2.5 DIRECTIONS FOR UPLOADING VIDEO FILES FROM DVR
Follow these procedures to upload video files to another storage device (EPA-provided USB
flash drive).
1. Insert the USB flash drive provided by EPA into an available USB port in your
computer.
2. Open the silver flap on the top left of the DVR and locate the "USB2" slot on
the right hand side.
3. Locate a cord in your kit that has a "USB2" end (doesn't appear to directly
match the opening shape on the DVR, but it is correct!) and a standard
computer USB end. It may be in a bag, or loose in the kit under the DVR.
4. Power on the DVR.
5. Once the home screen appears, connect the DVR and computer using the cord.
6. The DVR screen will turn black and flash USB 2.0, once it is connected.
7. On your computer, go My Computer and locate the DVR, "DPA1" -Select
8. Select the DVR folder.
9. All video files should be visible. Select all files - right click - copy - go to the
flash drive under My Computer - right click - paste.
10. Once files are transferred, close all windows.
11. On your computer, go to My Computer and select the flash drive. Confirm all
DVR files are present.
12. Disconnect computer and DVR and remove flash drive.
13. Power off DVR.
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12 SEDIMENT COLLECTION
Crews collect sediments for a variety of analyses. Field crews will sieve one or two
sediment grabs and submit the resulting benthic infauna collection to the lab to be
analyzed for species composition and abundance. Additional sediment grabs will be
analyzed for chemical contaminants (organics/metals and TOC), grain size determination,
and acute whole sediment toxicity. In order to provide the minimum volume of sediment
for all analyses, crews may need to collect different numbers of grabs at different sites,
based on sediment characteristics. While the biology (benthic assemblage) grab is being
processed (sieved) by one crew member, other personnel collect the necessary grabs for
chemistry, grain size, and toxicity tests. They composite the grabs, mix them and then
split them into four separate sample containers. Crews must collect a minimum of 3L of
sediment at marine sites and 2L of sediment at Great Lakes sites to submit for chemistry
(contaminants), toxicity, and grain size analyses.
12.1 SUMMARY OF METHOD
A 1725 (0.04) m2, stainless steel, Young-modified Van Veen Grab (or similar) sampler is
appropriate for collecting sediment samples for both biological and chemical analyses.
The top of the sampler is either hinged or otherwise removable so the top layer of
sediment can be easily removed for chemical and toxicity sample collection. This gear is
relatively easy to operate and requires little specialized training. For crews sampling in
the Great Lakes, a standard Ponar grab (box size 22.9 cm x 22.9 cm with depth of 9 cm)
with removable top screens should be used for collecting sediments for benthic
invertebrate analysis (USEPA 2001); other sediment grab devices may be used for
sediment toxicity and contaminant samples at the crew's discretion. Record the
dimensions and sample area of the grab used on the Sample Collection (Back) form. The
area of sediment the grab collects is important for data analysis. If the grab sampler size
is less than 0.03 m2, take two grabs for the benthic macroinvertebrate collections and
composite the sediment into the sieve.
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12.2 EQUIPMENT AND SUPPLIES
Table 12,1 Equipment Et supplies: sediment collection
For collecting samples
For recording
measurements
Young-modified Van Veen (or Ponar) grab with grab stand
weights and pads for grab
nitrile gloves
plastic tub or bucket
0.5 mm stainless steel sieve
sieve box or bucket
electrical tape
forceps (fine-tipped)
funnel (wide-mouth)
Alconox
Formalin (100% buffered) with stain
Graduated cylinder for measuring formalin
Rose Bengal Stain (for staining formalin solution)
Borax
ruler (cm)
squirt bottle (for ambient water)
stainless steel mixing pot or bowl with lid
stainless steel or Teflon spoons (15")/scoops/spatula
HDPE bottie(s) (1 L, wide-mouth) [BENT]
glass jar (60 mL, amber) [SEDC]
glass jar (120 mL, amber) [SEDO]
plastic bags (2,1 quart) [SEDG]
screw-top bucket (0.6 gallon) [SEDX]
scrub brush
cooler with wet ice
Sample Collection form
pencils (for data forms)
fine-tipped indelible markers (for labels)
clear tape strips
12.3 SAMPLING PROCEDURE
The following describes the sampling procedure to obtain sediment samples.
Note: The sampler, spoons and mixing bowl or bucket must be thoroughly rinsed with
ambient water after sampling at each site to ensure no sediments remain, and then at
the next station washed with Alconox and rinsed with ambient water prior to use. This
practice reduces the risk of the equipment carrying contaminants from site to site.
Do not apply sunscreen or other chemical contaminants until after the sample is
collected (or implement measures to reduce contamination by such chemicals if applied
such as washing, wearing long gloves, etc.). Be sure to use new clean nitrile gloves or
wash gloves between stations if they are reused from one station to the next.
1. Attach the sampler to the end of the winch cable with a shackle and tighten
the pin.
2. Set the grab according to the manufacturer's instructions and disengage any
safety device designed to lock the sampler open.
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3. Lower the grab sampler through the water column such that travel through the
last 5 meters is no faster than about 1 m/sec. This minimizes the effects of
bow wave disturbance to surficial sediments.
4. Allow a moment for the sampler to settle into the substrate and then allow
slack on the cable. Letting the cable go slack serves to release the jaws of the
sampler so they will close as the sampler is retrieved.
5. Retrieve the sampler and lower it into its cradle or a plastic tub on-board.
Open the top and determine whether the sampling is successful or not.
• A successful grab is one having relatively level, intact sediment over the
entire area of the grab, and a sediment depth at the center of at least 7
centimeters for the benthic macroinvertebrate grab (see Figure 12.1).
• Grabs containing no sediment, partially filled grabs, or grabs with
shelly/rocky substrates or grossly slumped surfaces are unacceptable.
• Grabs completely filled to the top, where the sediment is in direct contact
with the hinged top, are also unacceptable.
• It may take several attempts using different amounts of weight to obtain
the first acceptable sample. More weight will result in a deeper bite of the
grab. In very soft mud, pads may be needed to prevent the sampler from
sinking into the mud. If pads are used, the rate of descent near the bottom
should be slowed even further to reduce the bow wave.
6. If, after several attempts, only grabs less than 7 centimeters deep can be
obtained, use the next successful grab regardless of the depth of sediment at
the center of the grab.
• Use the comments on the Sample Collection (Back) form to describe your
efforts and be sure to accurately record the depth of the sediment
captured by the grab.
• Carefully drain overlying water from the grab. If the grab is used for
benthic community analysis, the water must be drained into the container
that will receive the sediment to ensure no organisms are lost.
• Enter notes on the condition of the sample (smell, substrate, presence of
organisms on the surface, etc.) in the Sediment Characteristics section of
the Sample Collection (Back) form.
7. If the grab sampler size is less than 0.03 m2, take two grabs for the benthic
macroinvertebrate collections and composite the sediment into the sieve.
8. Process the grab sample for either benthic community analysis or
chemistry/toxicity testing as described below.
9. Repeat steps 4-8 until all samples are successfully collected. To minimize the
chance of sampling the exact same location twice, the boat engines can be
turned periodically to change the drift of the boat, or additional anchor line
can be let out.
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Acceptable grab
At leasl 7 cm deep with even surlace
Unacceptable grab Unacceptable grab
Sloping surface Insufficient volume
Unacceptable grab Ur>acceplable grab
Wash-put Overfilled
Figure 12.11llustration of acceptable & unacceptable grabs for benthic community analysis. An acceptable
grab is at least 7 cm in depth (using a 0.04m2 Van Veen sampler), but not oozing out of the top of the grab,
and has a relatively level surface.
12.4 PROCESSING PROCEDURE - BENTHIC MACROINVERTEBRATE [BENT]
COMPOSITION AND ABUNDANCE
Grab samples obtained to assess the benthic macroinvertebrate community are processed
as outlined below.
1. Measure the depth of the sediment at the middle of the sampler and record the
value on the Sample Collection (Back) form. The depth should be >7 cm if
possible (see previous section).
• Record descriptive information about the grab, such as the presence or
absence of a surface floe, color and smell of surface sediments, and visible
fauna.
2. Dump the sediment into a clean basin (plastic tub or bucket) and then into a
0.5 mm mesh sieve. Place the sieve into a table (sieve box) containing water
from the sampling station, a larger bucket, or place the sieve over the side of
the boat.
• Gently agitate the sieve to wash away sediments and leave organisms,
detritus, sand particles, and pebbles larger than 0.5 mm. This method
minimizes mechanical damage to fauna that is common when forceful jets
of water are used to break up sediments.
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• A gentle flow of water over the sample is acceptable. Extreme care must
be taken to assure that no sample is lost over the side of the sieve.
3. Drain the water from the sieve and gently rinse the contents of the tray to one
edge. Remove large non-living items such as rocks and sticks after inspecting
them and ensuring that all benthic organisms are included in the collection.
• Using either your fingers or a spoon, GENTLY scoop up the bulk of the
sample and place it in the 1 L HOPE bottle (which should be placed in the
sieve or a bucket in case some of the sample spills over).
4. Complete the BENT sample label with Site ID, date collected, visit number, and
jar number. At marine sites, also write the salinity (in ppt) on the label.
5. Attach the completed label to the 1 L wide mouth sample bottle and cover
with clear plastic tape.
6. Rinse the outside of the sample jar into the sieve, then, using a funnel, rinse
the contents into the jar. The jar should be filled no more than one-half full.
• If the quantity of sample exceeds 500 ml, place the remainder of the
sample in a second container with a "2 of 2" label. For samples with a large
amount of benthos, additional jars may be needed.
7. Use a pencil to fill out waterproof benthic infauna (BENT) label(s) with the
pertinent sample information and place it inside the bottle(s). Be sure to
include the sample ID and jar number.
8. Record sample collection location and the total number of jars on the Sample
Collection (Back) form.
9. Carefully inspect the sieve to ensure that all organisms are removed. Use fine
forceps (if necessary) to transfer fauna from the sieve to the bottle containing
the proper sample number.
10. A stained 100% percent buffered formalin solution is used to fix and preserve
benthic samples. The solution should be mixed according to the directions in
Table 4.1. 100 ml of the formalin should be added to each sample jar along
with an additional teaspoon-full of borax to ensure saturation of the buffer.
Rose Bengal stain is added to the stock formalin solution for use at all sites.
• Make sure that there is sufficient preservative to ensure everything gets
preserved properly, then FILL THE JAR TO THE RIM WITH
SEAWATER/LAKEWATER TO ELIMINATE ANY AIR SPACE. This eliminates the
problem of organisms sticking to the cap because of sloshing during
shipment.
• Crews may choose to use a more dilute formalin solution in larger
quantities as long as the end concentration of the preservative is at least 6
percent.
11. After preservation, replace the bottle lid(s) and seal tightly with electrical
tape. Gently rotate the bottle to mix the contents and place in the dark.
• If the sample occupies more than one container, label all the sample
bottles containing material from that grab together. All benthos jars from a
single site will have the same sample ID number.
12. Prior to sieving the sample at the next site, use copious amounts of forceful
water and a stiff brush to clean the sieve, thereby minimizing cross-
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contamination of samples. Be sure to rinse the brush between each sieve
cleaning.
12.5 PROCESSING PROCEDURE - SEDIMENT COMPOSITION, CHEMISTRY, AND
TOXICITY
In addition to grab samples collected for benthic community analysis, additional grabs are
collected for chemical analyses (organics/metals and TOC), grain size determination, and
for use in acute toxicity tests. The top two centimeters of these grabs are removed,
homogenized, and split into these four sample types.
The samples are removed and processed in the order described below.
1. As each grab is retrieved, carefully examine it to determine acceptability. The
grab is considered acceptable as long as the surface layer is intact. The grab
need not be greater than 7 cm in depth for chemistry samples, but the other
criteria outlined above apply (see Section 12.3 and Figure 12.1 above).
• Carefully drain off, or siphon, any overlying water, and remove and
discard large, non-living surface items such as rocks or pieces of wood.
Remove any submerged aquatic vegetation (SAV) after recording its
presence on the Sample Collection (Back) form.
Note: Great care must be taken to avoid contamination of this sample
from atmospheric contaminants. The boat engine should be turned off or
the boat maneuvered to ensure the exhaust is downwind. All containers,
including the grab sampler, should be kept closed except when opening is
necessary to remove or add samples.
2. A clean stainless steel or Teflon spoon that has been washed with Alconox and
rinsed with ambient sitewater is used to remove sediments from grab samples
for these analyses.
3. Remove the top 2 cm of sediment using the stainless steel or Teflon spoon.
Sediment which is in direct contact with the sides of the sampler should be
excluded as they may be contaminated from the device.
• Place the sediment into a pre-cleaned (washed with Alconox and rinsed
with ambient sitewater) stainless steel pot or bowl and place the pot in
a cooler on wet ice (NOT dry ice). The sample must be stored at 4°C,
and MUST NOT BE FROZEN.
4. Repeat obtaining sediment samples from the grab and compositing the
sediment in the same stainless pot/bowl until a sufficient quantity of sediment
has been collected for all samples (approximately 3L at marine sites and 2L at
Great Lakes sites).
• Stir sediment homogenate after every addition to the composite to
ensure adequate mixing. Keep the container covered and in the cooler
between grabs.
5. Record the location (zone) of the sediment collection on the Sample Collection
(Back) form. If sediment was collected from more than one zone, fill in the
bubble of the zone where the majority of the sediment was collected and
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describe the proportions of sediment collected from each zone in the
comments section.
6. Homogenize the sediment by stirring with a Teflon paddle or stainless steel
spoon for 10 minutes. Divide the composite into the four sample types listed
below. In the case of limited sediment, prioritize sample distribution in the
order listed.
a) ORGANICS and METALS [SEDO]:
• Complete the SEDO sample label with Site ID, date collected, and
visit number.
• Attach the completed label to the 120 ml (4 oz) glass sample jar
and cover with clear plastic tape.
• Using a clean stainless steel spoon, carefully place approximately
100 ml of sediment into the jar. CARE MUST BE TAKEN TO ENSURE
THAT THE INSIDE OF THE JAR, CAP, AND THE SAMPLE IS NOT
CONTAMINATED. Be sure that you leave Vi inch headspace to avoid
breakage due to possible sample expansion from freezing.
• Replace the lid and seal tightly with electrical tape, wrap the jar in
the provided foam sleeve to protect it from breakage, and place the
sample in a cooler with dry ice.
• Record sample ID along with any comments on the Sample
Collection (Back) form.
• Fill in the "frozen" bubble on the sample collection form to confirm
that the sample has been frozen.
b) SEDIMENT TOXICITY [SEDX]:
• Complete the SEDX sample label with Site ID, date collected, and
visit number.
• Attach the completed label to the 0.6 gallon plastic sample bucket
and cover with clear plastic tape.
• Using the stainless steel spoon, fill the bucket with sediment to
within about 1 inch from the rim (Preferred volume for marine sites
is 1800 mL; if that is not possible, minimum volume required is 900
mL; for Great Lakes sites, preferred volume is 900 mL, minimum
required is 400 mL).
• Replace the lid and tighten so that the locking mechanism engages
and holds the lid tightly closed.
• Record sample ID along with any comments on the Sample
Collection (Back) form.
• Place the sample on wet ice (NOT dry ice). The sample must be
stored at 4°C, and MUST NOT BE FROZEN.
• Fill in the "chilled" bubble on the sample collection form to confirm
that the sample has been chilled.
c) TOTAL ORGANIC CARBON [SEDC]:
• Complete the SEDC sample label with Site ID, date collected, and
visit number.
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• Attach the completed label to the 60 ml glass sample jar and cover
with clear plastic tape.
• Using a clean stainless steel spoon, place approximately 50 ml of
sediment into the jar. Be sure that you leave 1/2 inch headspace to
avoid breakage due to possible sample expansion from freezing.
• Replace the lid and seal tightly with electrical tape, wrap the jar in
the provided foam sleeve to protect it from breakage, and place the
sample in a cooler with dry ice.
• Record sample ID along with any comments on the Sample
Collection (Back) form.
• Fill in the "frozen" bubble on the sample collection form to confirm
that the sample has been frozen.
d) SEDIMENT GRAIN SIZE [SEDG]:
• Complete the SEDG sample label with Site ID, date collected, and
visit number.
• Attach the completed label to the inner quart sized plastic sample
bag and cover with clear plastic tape.
• Using a clean stainless steel spoon, place approximately 100 ml of
sediment into the pre-labeled bag. Double bag the sample into a
second quart sized plastic bag, ensuring that the tops of both bags
are sealed tightly.
• Record sample ID along with any comments on the Sample
Collection (Back) form.
• Place the sample on wet ice (NOT dry ice). The sample must be
stored at 4°C, and MUST NOT BE FROZEN.
• Fill in the "chilled" bubble on the sample collection form to confirm
that the sample has been chilled.
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13 FISH TISSUE COLLECTION
Crews collect fish at all NCCA sites. At revisit sites, ecofish and fish plugs are only
collected during visit 1. At Great Lakes revisit sites that are also human health fish tissue
sites, crews that are unsuccessful at collecting the human health fish tissue sample during
visit 1 are expected to attempt the collection of that sample during visit 2. Labs analyze
whole body (also known as "ecological fish" or "ecofish") tissue samples for
concentrations of organic and inorganic contaminants. The results provide information
about the ecological risks to wildlife associated with fish consumption. Refer to Section
13.1 for detailed information regarding ecofish collections.
In addition to whole fish samples collected at all sites for ecological risk purposes, crews
will also collect fish tissue plugs at all sites. These plugs can be taken from fish collected
for the ecofish sample or crews can allow the fish to be released after the tissue plug
sample is collected. The sample is analyzed for mercury concentrations and used to
provide a measure of human health risk at all sites. Refer to Section 13.2 for a detailed
discussion of fish tissue plug collection.
Finally, crews at 150 Great Lakes sites collect a fish tissue sample for human health
contaminant analysis. Refer to Section 13.3 for detailed information regarding samples
collected for human health fish tissue contaminant analysis.
When target fish are plentiful, crews in the Great Lakes will be able to submit specimens
for both the ecofish and human health fish tissue collections. If specimens are less
plentiful, crews may be able to split the sample between the two whole fish collection
types and still meet the minimum criteria for each sample. In rare cases where only
enough fish are collected to fulfill the requirements of one of the samples, crews should
submit the fish as the ecofish sample and mark the human health fish tissue sample as not
collected.
13.1 ECOLOGICAL CONTAMINATION FISH TISSUE COLLECTION [FTIS]
13.1.1 SUMMARY OF METHOD
Ecological Fish Tissue collection protocols require crews to collect at least five individuals
of the target species, yielding a minimum of 300 g total mass from each site. These fish
are to be collected within a 500 meter radius of the X-site (may expand to 1000 meters if
needed - see below and Figure 5.2). Crews may collect these samples using any
reasonable method (e.g., otter trawl, hook and line, gill net, seine, etc.) that is most
efficient and the best use of available time on station.
For each attempted fish collection method, record equipment details, start and stop
times, and fishing location(s) on the Eco Fish Collection (Front) form. Record sample ID,
species retained, and specimen lengths on the Eco Fish Collection (Back) form.
Secondary fish tissue collection zones for ecofish and/or fish plugs may be selected up to
an additional 500 m beyond the original 500 m radius at all estuarine and Great Lakes
sites. Please observe the following guidelines:
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1. In order to move to a secondary fish tissue collection zone, crews must be
unsuccessful at obtaining target fish during a reasonable portion of the three
hours allotted to fishing (at least 30 minutes and no more than two hours)
within the original 500 m radius.
2. The crew must have attempted several sampling locations within the 500 m
radius without success.
3. Crews must observe signs of fish presence such as schools of bait fish just
below the surface, predator activity or prey escape behavior on the surface of
the water, overhead shading or favorable underwater habitat structure or
bathymetric features within an additional 500 m from the X-site.
4. When relocating outside of the original 500 m radius from the X-site, but inside
of the 1000 m radius of the X-site, crews must document:
a) The amount of time spent fishing within the original 500-meter radius.
b) The direction of travel from the X-site.
c) The coordinates of the site where fish were ultimately caught.
5. For collection of the human health fish tissue sample ONLY (if applicable),
crews may move out to a maximum of 1500 meters from the X-site in an effort
to collect this sample.
Crews working in each of the regional areas— Northeast, Southeast, Gulf, West Coast, and
Great Lakes —collect different target fish species based on biogeographically specific lists.
Recommended Primary and Secondary target species are given by region in the following
tables:
• Northeast-Table 13.2
• Southeast-Table 13.3
• Gulf of Mexico - Table 13.4
• West-Table 13.5
• Great Lakes - Table 13.6
If a full composite sample is not collected after 3 hours of effort, crews may terminate
the sampling, record the details of the sample, and submit as many fish as possible. If the
target species are unavailable, the fisheries biologist selects an alternative available
species (i.e., a species that is commonly present in the study area and in sufficient
numbers to yield a composite) to obtain a fish composite sample. However, all attempts
should be made to collect the targeted species if at all possible. Regardless of the species
that is ultimately collected, all fish in the composite MUST be of the same species.
Crews may spend additional time fishing (i.e. more than three hours) if desired. It is not
recommended that crews purchase fish specimens dockside unless they can document that
the purchased fish came from an area in close proximity to the X-site (i.e. within 1000
meters).
Crews identify specimens to species and measure the total length to the nearest
millimeter. They record the taxonomic name (genus-species) and the length of each fish
on the Eco Fish Collection (Back) form. The preferred minimum length for a specimen for
ecological risk purposes is 100 mm with a preferred length range of 100 - 400 mm. All
individuals must be of similar size, such that the smallest individual in the composite is no
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less than 75% of the total length of the largest individual. Up to 20 individuals (a total of
300 g of whole body tissue is needed) should be collected and retained for analysis. If it is
suspected that 20 individuals will yield less than 300 g total weight, additional specimens
should be collected. The lengths of any additional fish should be recorded in the blank
space provided.
13.1.2 EQUIPMENT AND SUPPLIES
Table 13.1 Equipment ft supplies: eco fish tissue collection
For collecting fish
composite sample
scientific collection permit
Otter trawl (or other device to collect sufficient sample)
sampling vessel (including boat, motor, trailer, oars, gas, and all required safety
equipment) Coast Guard-approved personal flotation devices
Global Positioning System (GPS) unit
nitrile gloves
livewell and/or buckets
measuring board (millimeter scale)
scale (in grams)
wooden bat
For storing and
preserving fish
composite sample
For documenting the
fish composite
sample
For shipping the fish
composite samples
self-sealing bag(s) (plastic, 2 gallon)
large plastic (composite) bags
self-sealing bag(s) (sandwich size) — for labels cooler
plastic cable tie
dry ice or wet ice (for temporary transport)
side cutter (cleaned with Alconox between sites)
Eco Fish Collection form
fish tissue sample labels
pencils (for data forms)
fine-tipped indelible markers (for labels)
Tyvek label tag with grommet
clear tape strips
Tracking: Eco Fish Tissue — Overnight (Dry Ice) form
FedEx airbill (pre-addressed)
cooler
dry ice (~20 Ibs per cooler)
packing/strapping tape
13.1.3 SAMPLING PROCEDURE
The procedures for collecting and processing ecological fish composite samples are
presented below. If fish plugs are to be collected from specimens in the ecofish
collection, complete the steps in Section 13.2 before packaging the ecofish collection.
Note: Do not handle any food, drink, sunscreen, or insect repellant until after the
composite sample has been collected, measured and bagged (or implement measures to
reduce contamination by such chemicals if applied such as washing, wearing long gloves,
etc.)
1. Put on clean nitrile gloves before handling the fish.
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2. Rinse potential target species/individuals in ambient water to remove foreign
material from the external surface and place them in clean holding containers
(e.g., livewells, buckets).
3. Select at least five fish, with a minimum total weight of 300 grams, to include
in the eco fish composite. If needed, 20 or more fish may be composited to
reach the minimum weight of 300 grams. The selected fish must meet the
following criteria:
• all fish are of the same species.
• the preferred specimen length is between 100 and 400 mm; if after
sufficient fishing only smaller or larger fish of the target species are
available, those will be accepted.
• all fish are of similar size, so that the smallest individual in a composite
is no less than 75% of the total length of the largest individual.
• all fish for one site visit are collected as close to the same time as
possible, but no more than one week apart.
Note: Individual fish may have to be frozen until all fish to be included in
the composite are available for delivery to the designated laboratory.
4. Identify the fish to species and record the scientific name on the Eco Fish
Collection (Back) form.
Note: Accurate taxonomic identification is essential in assuring and
defining the composited organisms submitted for analysis. Individuals from
different species may not be composited in a single sample. Submit only
one species per site.
5. Measure each individual fish from the anterior-most part of the fish to the tip
of the longest caudal fin ray (when the lobes of the caudal fin are depressed
dorsoventrally) to determine total body length in millimeters.
6. Record collection method and equipment details, start and stop times, and
fishing location(s) on the Eco Fish Collection (Back) form. Record sample ID,
species name and specimen lengths on the Eco Fish Collection (Back) form.
Make sure the sample ID recorded on the collection form match those on the
sample labels.
7. While wearing clean nitrile gloves, remove each fish retained for analysis from
the clean holding container(s). Dispatch larger fish using a clean wooden bat
(or equivalent wooden device).
8. Place all fish from the composite in a two-gallon self-sealing bag. Take care to
prevent fish spines from piercing the bag. If spines are likely to puncture the
bag, break off or clip the spines with a side-cutter or other appropriate tool
(cleaned with Alconox and rinsed with ambient sitewater before use at each
site) and place the spine in the bag with the fish. Use additional bags if all the
fish collected for a composite will not fit in a single two-gallon bag.
9. Weigh the composite bag(s) to determine if enough fish have been collected to
reach a minimum weight of 300 grams.
10. Prepare interior and exterior FTIS sample labels for the two-gallon bag(s),
ensuring that the label information matches the information recorded on the
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Eco Fish Collection (Back) form. Be sure to record scientific name and
minimum and maximum lengths on the labels.
• Place the interior label inside a small (sandwich-size) self-sealing bag
and place the bag inside the two-gallon bag with the fish composite.
• Affix the exterior label to the two-gallon bag and cover with clear
plastic tape. If additional two-gallon bags are used, fill out extra labels
with the same sample ID and information for each bag and label
accordingly (i.e. bag 2 of 2).
11. Double-bag all specimens in the composite by placing all two-gallon bag(s) from
the site inside a large plastic bag.
12. Prepare a sample label for the outer bag, ensuring that the label information
matches the information recorded on the Eco Fish Collection (Back) form. Be
sure to record scientific name and minimum and maximum lengths on the
sample label.
13. Affix the sample label to a Tyvek tag and cover with clear plastic tape. Thread
a cable tie through the grommet in the Tyvek tag and seal the outer bag with
the cable tie.
13.1.4 SAMPLE STORAGE AND SHIPPING PREPARATION
1. After the sample is packaged, immediately place it on dry ice for shipment.
• Fill in the "frozen" bubble on the Eco Fish Collection (Back) form to
confirm that the sample has been frozen.
• Packaged samples may be placed on wet ice in coolers if they will be
transported to a laboratory or other interim facility to be frozen before
shipment.
• Samples may be stored on wet ice for a maximum of 24 hours.
• Freeze the samples within 24 hours of collection at <-20°C and store the
frozen samples until shipment within 2 weeks of sample collection. Crews
may ship the frozen fish sample along with the other frozen samples from
the site using a cooler with a dry ice insert or may ship the ecofish
separately. Frozen samples should be packed on at least 20 pounds of
layered dry ice and shipped to the batched sample lab via priority overnight
delivery service.
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Table 13.2 Northeast region primary and secondary marine target species - whole body fish tissue collection
(Ecofish)
NC
FAMILY
Moronidae
Paralichthyidae
Pleuronectidae
•
Sparidae
NOR
• JHIII
Achiridae
Anguillidae
Atherinopsidae
Batrachoididae
Ephippidae
Moronidae
Mugulidae
Pomatomidae
•
Serranidae
Triakidae
Triglidae
IRTHEAST REGION PRIMARY E
SCIENTIFIC NAME
Ameiurus catus
Ictalurus punctatus
Morone americana
Paralichthys dentatus
Pseudopleuronectes americanus
Cynoscion regalis
Sciaenops ocellatus
Stenotomus chrysops
THEAST REGION SECONDARY
SCIENTIFIC NAME
Trinectes maculatus
Anguilla rostrata
Menidia menidia
Opsanus tau
Chaeto dip terus faber
Morone saxatilis
Mugil cephalus
Pomatomus saltatrix
Bairdiella chrysoura
Menticirrhus saxatilis
Centropristis striata
Mustelus canis
Prionotus carolinus
Prionotus evolans
COFISH TARGET SPECIES
COMMON NAME
White catfish
Channel catfish
White perch
Summer flounder
Winter flounder
Gray weakfish
Red drum
Scup
ECOFISH TARGET SPECIES
COMMON NAME
Hogchoaker
American eel
Atlantic silverside
Oyster toadfish
Atlantic spadefish
Rock fish
Black mullet
Bluefish
Silver perch
Northern kingfish
Black sea bass
Smooth dogfish
Northern searobin
Striped searobin
FISH PLUG LIST*
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Secondary
Secondary
Secondary
' Indicates whether species also occurs in the primary or secondary fish plug list (see Table 13.8).
Table 13.3 Southeast region primary and secondary marine target species - whole body fish tissue collection
(Ecofish)
axmvm
' '
Ariidae
Paralichthyidae
Sciaenidae
Sparidae
UTHEAST REGION PRIMARY ECOFISH TARGET SPECIES
SCIENTIFIC NAME
BSrtTAyAitifUnTT^I
FISH PLUG LIST*
Ariopsisfelis I Hardhead sea catfish | Primary
Bagre marinus
Paralichthys albigutta
Paralichthys dentatus
Paralichthys lethostigma
Cynoscion arenarius
Cynoscion nebulosus
Cynoscion regalis
Leiostomus xanthurus
Lagodon rhomboides
Gafftopsail sea catfish
Gulf flounder
Summer flounder
Southern flounder
Sand weakfish (or seatrout)
Speckled trout
Gray weakfish
Spot croaker
Pinfish
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
THEAST REGION SECONDARY ECOFISH TARGET SPECIES
SCIENTIFIC NAME
^^Jyfy^^ 1 L riTMl ^^H
FISH PLUG LIST*
Cichlidae Tilapia mariae \ Spotted tilapia
Haemulidae
Sciaenidae
Serranidae
Haemulon aurolineatum
Bairdiella chrysoura
Menticirrhus americanus
Centropristis striata
Tomtate
Silver perch
Southern kingfish
Black sea bass
' Indicates whether species also occurs in the primary or secondary fish plug list (see Table 13.8).
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Table 13.4 Gulf region primary and secondary marine target species - whole body fish tissue collection (Ecofish)
GULF REGION PRIMARY ECOFISH TARGET SPECIES
Ariidae
Paralichthyidae
Sciaenidae
Sparidae
SCIENTIFIC NAME
Ariopsisfelis
Bagre marinus
Paralichthys albigutta
Pamlichthys dentatus
Paralichthys lethostigma
Cynoscion arenarius
Cynoscion nebulosus
Cynoscion regalis
Leiostomus xanthurus
Micropogonias undulatus
Sciaenops ocellatus
Lagodon rhomboides
COMMON NAME FISH PLUG LIST*
Hardhead sea catfish
Gafftopsail sea catfish
Gulf flounder
Summer flounder
Southern flounder
Sand weakfish (orseatrout)
Speckled trout
Gray weakfish
Spot croaker
Atlantic croaker
Red drum
Pinfish
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
GULF REGION SECONDARY ECOFISH TARGET SPECIES
Carangidae
Diodontidae
Gerreidae
Haemulidae
Ictaluridae
Lepisosteidae
Lutjanidae
Sciaenidae
Serranidae
Triglidae
SCIENTIFIC NAME
COMMON NAME FISH PLUG LIST*
Caranx hippos Crevallejack
Chloroscombrus chrysurus
Chilomycterus schoepfii
Eucinostomus gula
Orthopristis chrysoptera
Ictalurus furcatus
Lepisosteus oculatus
Lutjanus griseus
Pogonias cromis
Di plectrum formosum
Prionotus scitulus
Atlantic bumper
Burrfish
Silver jenny
Pigfish
Blue catfish
Spotted gar
Gray snapper
Black drum
Sand perch
Leopard searobin
' Indicates whether species also occurs in the primary or secondary fish plug list (see Table 13.8).
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Table 13.5 Western region primary and secondary marine target species - whole body fish tissue collection
(Ecofish)
v\
Atherinopsidae
Cynoglossidae
Gasterosteidae
Paralichthyidae
Sciaenidae
Echinodermata/
Toxopneustidae
Chimaeridae
Embiotocidae
Paralichthyidae
Sciaenidae
/ESTERN REGION PRIMARY EC
SCIENTIFIC NAME
Atherinops affinis
Leptocottus armatus
Oligocottus rimensis
Symphurus atricaudus
Cymatogaster aggregate!
Embiotoca lateralis
Gasterosteus aculeatus
Paralichthys californicus
Citharichthys sordidus
Citharichthys stigmaeus
Isopsetta isolepis
Parophrys vetulus
Psettichthys melanostictus
Platichthys stellatus
Genyonemus lineatus
Paralabrax nebulifer
Paralabrax maculatofasciatus
SCIENTIFIC NAME
Tripneustes gratilla
(Hawaii ONLYJ
Porichthys notatus
Porichthys myriaster
Hydrolagus colliei
Amphistichus argenteus
Xystreurys liolepis
Pleuronichthys guttulatus
Micmstomus pacificus
Lepidopsetta bilineata
Lyopsetta exilis
Umbrina roncador
OFISH TARGET SPECIES
COMMON NAME
Topsmelt silverside
Pacific staghorn sculpin
Saddleback sculpin
California tonguefish
Shiner perch
Striped seaperch
Three-spined stickleback
California flounder
Pacific sanddab
Speckled sanddab
Butter sole
English sole
Pacific sand sole
Starry flounder
White croaker
Barred sand bass
Spotted sand bass
COFISH TARGET SPECIES
Collector urchin
Plainfin midshipman
Specklefin midshipman
Spotted ratfish
Barred surfperch
Fantail sole
Diamond turbot
Dover sole
Rock sole
Slender sole
Yellowfin croaker
FISH PLUG LIST*
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
^m i
FISH PLUG LIST*
Secondary
Secondary
Secondary
* Indicates whether species also occurs in the primary or secondary fish plug list (see Table 13.8).
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Table 13.6 Great Lakes primary and secondary target species - whole body fish tissue collection (Ecofish)
FAMILY SCIENTIFIC NAME COMMON NAME
Catostomidae
Centrarchidae
Cottidae
Cyprinidae
Esocidae
Gasterosteidae
Gobiidae
Ictaluridae
Gadidae
Moronidae
Osmeridae
Percidae
Percopsidae
Salmonidae
Sciaenidae
Moxostoma macrolepidotum Shorthead redhorse
Ambloplites rupestris
Lepomis qibbosus
Lepomis macrochirus
Micropterus dolomieu
Pomoxis annularis
Pomoxis niqromaculatus
Cottus bairdii
Cottus coqnatus
Couesius plumbeus
Cyprinus carpio
Pimephales notatus
Esox lucius
Esox masquinonqy
Gasterosteus aculeatus
Neoqobius melanostomus
Proterorhinus marmoratus
Ameiurus nebulosus
Ictalurus punctatus
Noturus flavus
Lota lota
Morone americana
Morone chrysops
Osmerus mordax
Gymnocephalus cernuus
Perca flavescens
Percina caprodes
Sander canadensis
Sander vitreus
Percopsis omiscomaycus
Coreqonus artedi
Coreqonus dupeaformis
Oncorhynchus qorbuscha
Oncorhynchus kisutch
Oncorhynchus mykiss
Oncorhynchus tshawytscha
Salvelinus namaycush
Aplodinotus qrunniens
Rock bass
Pumpkinseed
Bluegill
Smallmouth bass
White crappie
Black crappie
Mottled sculpin
Slimy sculpin
Lake chub
Common carp
Bluntnose minnow
Northern pike
Muskellunge
Three-spined stickleback
Round goby
Tubenose goby
Brown bullhead
Channel catfish
Stonecat
Burbot
White perch
White bass
American/ rainbow smelt
Ruffe
Yellow perch
Logperch
Sauger
Walleye
Trout-perch
Cisco/ lake herring
Lake whitefish
Pink salmon
Coho salmon
Rainbow trout
Chinook salmon
Lake trout
FISH PLUG LIST*
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Primary
Freshwater drum Primary
GREAT LAKES SECONDARY ECOFISH TARGET SPECIES
FAMILY SCIENTIFIC NAME
Catostomidae
Centrarchidae
Clupeidae
Cyprinidae
Esocidae
Fundulidae
Ictaluridae
Salmonidae
WSSSSSSmSSSSSSSII!^^^^^
Catostomus commersonii
Moxostoma anisurum
Micropterus salmoides
Alosa pseudoharenqus
Dorosoma cepedianum
Cyprinella spiloptera
Luxilus cornutus
Notropis stramineus
Esox niqer
Fundulus diaphanus
Fundulus maialis
Ameiurus melas
Prosopium cylindraceum
Sal mo trutta
Salvelinus fontinalis
Salvelinus fontinalis x namaycush
COMMON NAME FISH PLUG LIST*
IBm!E!!|Sffl£!^^^^H
White sucker
Silver redhorse
Largemouth bass
Alewife
American gizzard shad
Spotfin shiner
Common shiner
Sand shiner
Chain pickerel
Banded killifish
Striped killifish
Black bullhead
Round whitefish
Brown trout
Brook trout
Splake
Secondary
Secondary
' Indicates whether species also occurs in the primary or secondary fish plug list (see Table 13.9).
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13.2 FISH TISSUE PLUG [FPLG]
13.2.1 SUMMARY OF METHOD
Because many fish spend their entire life in a particular water body, they can be
important indicators of water quality, especially for toxic pollutants (e.g., pesticides and
trace elements). Toxic pollutants, which may be present in the water column or
sediments at concentrations below our analytical detection limits, can be found in fish
tissue above detection limits due to bioaccumulation.
Typical fish tissue collection methods require the fish to be sacrificed, whether it be a
whole fish or a skin-on fillet tissue sample. This can be problematic when there is a need
to collect large trophy-sized fish for contaminant analysis or when a large sample size is
necessary for statistical analysis. The following method collects fish tissue plugs instead of
a skin-on fillet. One fish tissue plug for mercury analysis will be collected from each of
two fish of the same species (one plug per fish) from the target list (below) at every site.
These fish are collected during the ecological fish tissue collection effort (Sections 13.1
and 13.3). In order of preference, fish tissue plugs should be collected from 1) an
ecological fish specimen that will be sent to the lab (when size and species requirements
overlap), or 2) a live fish that will be released after the plug has been collected. When
possible, select larger individuals from which to collect the fish plugs. Do not collect fish
plugs from specimens that are part of the human health fish tissue sample collection. A
tissue plug sample is collected by inserting a biopsy punch into a de-scaled area of dorsal
muscle section of a fish. After the plug has been collected, ecofish specimens are frozen
according to the protocol in Section 13.1; if a plug is collected from a live fish, antibiotic
salve is placed over the wound and the fish is released.
13.2.2 EQUIPMENT AND SUPPLIES
Table 13.7 lists the equipment and supplies necessary for field crews to collect fish
tissue plug samples. Record the fish tissue plug sampling data in the Fish Tissue Plug
Samples section of the Eco Fish Collection (Back) form.
Table 13,7 Equipment & supplies: fish tissue plugs
For fish tissue plug antibiotic salve
samples cooler with dry ice
cooler with wet ice
dip net
biopsy punch (sterile, disposable)
fish collection gear (trawl, nets, livewell, etc.)
disposable forceps (sterile)
glass scintillation vial (20 niL)
nitrile gloves
measuring board
aspirator bulb
scale (in grams)
scalpel (disposable, sterile)
For recording Eco Fish Collection (Back) form
measurements fish tissue plug sample labels
pencils (for data forms)
fine-tipped indelible markers (for labels)
clear tape strips
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13.2.3 SAMPLING PROCEDURE
The fish tissue plug indicator samples will be collected using the same gear and
procedures used to collect the ecological and/or human health fish tissue samples, and
collection occurs within the same area as other fish collections. Samples should be taken
from the species listed in the target list (primary and secondary species) found in Table
13.8 and Table 13.9. When ecofish specimens meet the size and species requirements for
fish plug samples, the plugs should be taken from the ecofish prior to placing on ice. If
ecofish specimens do not meet the size and species requirement for fish plugs, fish plugs
should be taken from live fish and the fish are released with antibiotic salve on the
wound, as in step 14 below. If the recommended primary and secondary species are
unavailable, the fisheries biologist will select an alternative species (i.e., a species that is
commonly consumed in the study area, with specimens of harvestable or consumable size)
to obtain a sample from the species that are available. If a listed species is unavailable,
aim to collect fish in the following order: 1) those that are consumed by humans; 2)
predatory fish; and 3) other available fish species. In no instance should fish plugs be
removed from specimens submitted for the human health fish tissue sample.
In order of preference, crews should try to submit species from 1) the Primary Target List;
2) the Secondary Target List; and 3) any other available fish. It is recognized that there
are species not on these lists that may be culturally or regionally important food sources,
essential to subsistence fishers or increasingly popular among food trends. For these
reasons, the guidance for selecting species for fish plug samples is purposefully inclusive.
Please note: There are no invertebrate organisms on this list with the exception of sea
urchins for Hawaii. Crab, shrimp, molluscs, lobsters, etc., will not be used in assessment
of mercury content in fish plugs. If invertebrate species are submitted for FPLG samples,
those data will be reported as MISSING for the associated sites.
The procedures for collecting and processing fish plug samples are presented below.
1. Spread out a cooler liner bag on a flat surface for your workspace.
2. Prepare the FPLG sample label with Site ID, date collected, and visit number.
3. Attach the completed label to the 20 milliliter scintillation vial and cover with
clear tape.
4. Put on clean nitrile gloves before handling the fish.
Note: Do not handle any food, drink, sunscreen, or insect repellent until
after the plug samples have been collected (or implement measures to
reduce contamination by such chemicals if applied such as washing,
wearing long gloves, etc.).
5. Rinse potential target species/individuals in ambient water to remove any
foreign material from the external surface and place in clean holding
containers (e.g., livewells, buckets). Return non-target fishes or small
specimens to the water.
6. Retain two individuals of the same target species from each site. The fish
should be:
• large enough to collect a fish plug yielding ~ 0.5 grams (wet weight) of
tissue,
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• on the recommended primary or secondary target list (if not available
select an alternative species present),
• both the same species,
• both satisfy legal requirements of harvestable size (or weight) for the
sampled water body, or at least be of consumable size and
• of similar size, so that the smaller individual is no less than 75% of the total
length of the larger individual.
Note: Whenever possible, larger specimens should be selected over smaller
specimens.
7. Remove one fish retained for analysis from the clean holding container(s) (e.g.,
livewell) using clean nitrile gloves.
8. Measure the fish to determine total body length. Measure total length of the
specimen in millimeters from the anterior-most part of the fish to the tip of
the longest caudal fin ray (when the lobes of the caudal fin are depressed
dorsoventrally).
9. Weigh the fish in grams using the fish weigh scale.
10. Note any anomalies (e.g., lesions, cuts, sores, tumors, fin erosion) observed on
the fish.
11. Record sample ID, species, and specimen length and weight in the Fish Tissue
Plug Samples section of the Eco Fish Collection (Back) form. Make sure the
sample ID numbers and specimen numbers/lengths that are recorded on the
collection form match those on the sample tracking form and labels, where
applicable.
12. On a meaty portion of the left side, dorsal area of the fish between the dorsal
fin and the lateral line, clear a small area of scales with a sterile disposable
scalpel.
13. Wearing clean nitrile gloves, insert the 8 millimeter biopsy punch into the
dorsal muscle of the fish through the scale-free area. The punch is inserted
with a slight twisting motion cutting the skin and muscle tissue. Once full depth
of the punch is achieved, a slight bending or tilting of the punch is needed to
break off the end of the sample. Remove biopsy punch taking care to ensure
sample remains in the punch.
Note: The full depth of the punch should be filled with muscle tissue,
which should result in collecting a minimum of 0.25 to 0.35 grams of fish
tissue for mercury analysis.
14. If the fish is to be released, apply a generous amount of antibiotic salve to the
plug area and gently return the fish to the water. If the fish is part of the
ecofish collection, return the fish to the ecofish holding area without the
application of antibiotic.
15. Using an aspirator bulb placed on the end of the biopsy punch, give a quick
squeeze, blowing the tissue sample into the 20 milliliter scintillation vial.
16. Place the vial with sample immediately on dry ice for temporary storage.
17. Repeat steps 2-15 for the second fish, to collect a second fish plug sample.
Place the second plug in the same scintillation vial as the first. The two plugs
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should provide at least 0.5 grams of tissue. NOTE: If two qualifying fish cannot
be caught, both plugs may be taken from the same fish.
18. Replace the lid and seal tightly with electrical tape, insert the vial into the
"bubble bag" to protect it from breakage, and then place it into the 4 by 4 self-
sealing bag. Place the sample in a cooler with dry ice
19. Dispose of gloves, scalpel, and biopsy punch.
13.2.4 SAMPLE STORAGE
1. Keep the samples frozen on dry ice or in a freezer at <-20°C until shipment.
2. Frozen samples will subsequently be packed on dry ice and shipped to the
batched sample laboratory via priority overnight delivery service within 1
week. Please see Appendix C: Shipping and Tracking Guidelines for next
steps.
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Table 13.8 Primary and secondary marine target species for fish plug collection
F
Cottidae
Moronidae
Pleuronectidae
Sciaenidae
Sparidae
SE
^MTn^^^l
Anguillidae
Moronidae
Paralichthyidae
Pomatomidae
Sciaenidae
Scorpaenidae
'RIMARY MARINE FISH PLUG TARGI
SCIENTIFIC NAME
Ariopsisfelis
Bagre marinus
Leptocottus armatus
Cymatogaster aggregate!
Embiotoca lateralis
Ameiurus catus
Ictalurus punctatus
Morone americana
Citharichthys sordidus
Citharichthys stigmaeus
Paralichthys albigutta
Paralichthys californicus
Paralichthys dentatus
Paralichthys lethostigma
Parophrys vetulus
Platichthys stellatus
Pseudopleuronectes americanus
Cynoscion arenarius
Cynoscion nebulosus
Cynoscion regalis
Genyonemus lineatus
Leiostomus xanthurus
Micropogonias undulatus
Sciaenops ocellatus
Paralabrax maculatofasciatus
Paralabrax nebulifer
Stenotomus chrysops
CONDARY MARINE FISH PLUG TAR
SCIENTIFIC NAME
An gu ill a rostrata
Amphistichus argenteus
Amphistichus rhodoterus
Embiotoca jacksoni
Hyperprosopon argenteum
Morone saxatilis
Hippoglossina oblonga
Hippoglossoides platessoides
Limandaferruginea
Microstomus pacificus
Pleuronichthys guttulatus
Pomatomus saltatrix
Menticirrhus undulatus
Scorpaena guttata
Sebastes caurinus
Sebastes entomelas
Sebastes flavidus
Sebastes melanops
Sebastes mystinus
Sebastes paucispinis
ET SPECIES
COMMON NAME
Hardhead sea catfish
Gafftopsail sea catfish
Pacific staghorn sculpin
Shiner perch
Striped seaperch
White catfish
Channel catfish
White perch
Pacific sanddab
Speckled sanddab
Gulf flounder
California flounder
Summer flounder
Southern flounder
English sole
Starry flounder
Winter flounder
Sand weakfish (orseatrout)
Speckled trout
Gray weakfish
White croaker
Spot croaker
Atlantic croaker
Red drum
Spotted sand bass
Barred sand bass
Scup
GET SPECIES
COMMON NAME
American eel
Barred surfperch
Redtail surfperch
Black perch
Walleye surfperch
Rock fish
Fourspot flounder
American dab
Yellowtail flounder
Dover sole
Diamond turbot
Blue fish
California whiting
California scorpionfish
Copper rockfish
Widow rockfish
Yellowtail rockfish
Black rockfish
Blue rockfish
Bocaccio
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Serranidae
Triakidae
Paralabrax clathratus
Triakis semifasciata
Kelp bass
Leopard shark
Table 13.9 Primary and secondary Great Lakes target species for fish plug collection
PRIM,
Catostomidae
Cyprinidae
Gadidae
Salmonidae
Sciaenidae
Catostomidae
Ictaluridae
Salmonidae
ARY GREAT LAKES FISH PLUG TAR
SCIENTIFIC NAME
Moxostoma macrolepidotum
Ambloplites rupestris
Lepomis gibbosus
Lepomis macrochirus
Micropterus dolomieu
Cyprinus carpio
Esox lucius
Esox masquinongy
Ameiurus nebulosus
Ictalurus punctatus
Lota lota
Mo rone americana
Morone chrysops
Perca flavescens
Sander vitreus
Coregonus dupeaformis
Oncorhynchus kisutch
Oncorhynchus mykiss
Oncorhynchus tshawytscha
Salvelinus namaycush
Aplodinotus grunniens
DARY GREAT LAKES FISH PLUG TA
SCIENTIFIC NAME
Catostomus commersonii
Ictalurus furcatus
Sal mo trutta
GET SPECIES
COMMON NAME
Shorthead redhorse
Rock bass
Pumpkinseed
Bluegill
Smallmouth bass
Common carp
Northern pike
Muskellunge
Brown bullhead
Channel catfish
Burbot
White perch
White bass
Yellow perch
Walleye
Lake whitefish
Coho salmon
Rainbow trout
Chinook salmon
Lake trout
Freshwater drum
RGET SPECIES
COMMON NAME
White sucker
Blue catfish
Brown trout
13.3 HUMAN HEALTH FISH TISSUE COLLECTION [HTIS] (SELECT GREAT LAKES
SITES ONLY)
13.3.1 SUMMARY OF METHOD
Field crews collect human health fish tissue composites at a subset of 150 of the Great
Lakes sites (30 sites per lake). These sites are designated with "FT" in the panel code. If
a site has been designated as a human health fish tissue site and is dropped, the
replacement site will take on the FT designation and human health fish tissue should be
collected. At revisit sites that are also human health fish tissue sites, crews that are
unsuccessful at collecting the human health fish tissue sample during visit 1 are expected
to attempt the collection of that sample during visit 2.
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Labs analyze fillet tissue for mercury, polychlorinated biphenyls (PCBs), fatty acids,
perfluorinated compounds (PFCs), and additional contaminants of emerging concern (e.g.,
polybrominated diphenyl ethers or PBDEs).
This section contains the sampling procedures and target species for human health fish
tissue collection. Note that the human health fish species table (Table 13.11) includes 25
primary target species and 18 secondary fish species. Field crews must attempt to collect
a primary target species wherever possible. If primary target species are not available at a
particular site, then the field crew collects a composite of one of the secondary fish
species. In the event that a crew is unable to collect fish which are on the human health
species list, then the field crew should contact the Great Lakes Human Health Fish Tissue
Manager.
As with the ecological fish tissue samples, crews collect human health fish tissue samples
using any reasonable method that represents the most efficient or best use of the
available time on station (e.g., gill net, otter trawl, or hook and line). However, in
contrast to the allowable procedures for ecological fish tissue samples, crews may not
purchase fish for human health fish tissue collection. Record sample collection
information on the Human Health Fish Collection (Front) form.
For each attempted fish collection method, record equipment details, start and stop
times, and fishing location(s) on the Human Health Fish Collection (Front) form. Record
sample ID, species retained, and specimen lengths on the Human Health Fish Collection
(Back) form.
Identify and measure the specimens collected for each composite. Record the scientific
name (genus and species) and total length for each specimen on the Human Health Fish
Collection (Back) form. Human health fish composites should consist of 5 similarly sized
(i.e., the total length of the smallest specimen is no less than 75% of the total length of
the largest specimen) adult fish of the same species that will collectively yield about 500
g of fillet tissue. This translates to a total of about 20 ounces, or about 4 ounces of fillets
per fish (assuming collection of a 5-fish composite). Field crews should make every effort
to consistently obtain 5 fish for the human health fish tissue sample; however, a sample of
fewer than 5 fish is acceptable if it provides sufficient fillet tissue to meet the
requirement (500 g). Conversely, for the exceptions where field crews collect 5 fish that
are too small to collectively meet the fillet tissue requirement, they should collect
additional fish as necessary to provide adequate tissue.
Fish submitted as part of the human health fish tissue sample should remain intact and be
submitted as whole specimens. Crews should not take fish plugs from human health fish
tissue specimens.
13.3.2 EQUIPMENT AND SUPPLIES
Table 13.10 lists the equipment and supplies necessary for field crews to collect human
health fish tissue samples. Additional human health fish collection supplies can be ordered
through the Supply Request Form. A list of frequently asked questions and responses will
be provided with the fish sampling supplies to clarify situations that field crews may
encounter while collecting human health fish composites. Detailed procedures for
collecting and processing fish composite samples are presented below.
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Table 13.10 Equipment ft supplies: human health fish tissue collection
For collecting fish
composite sample
For storing and
preserving fish
composite sample
For documenting the
fish composite
sample
For shipping the fish
composite samples
scientific collection permit
gill net, otter trawl, hook and line (or other device to collect sufficient sample)
sampling vessel (including boat, motor, trailer, oars, gas, and safety equipment)
nitrile gloves
Coast Guard-approved personal floatation devices
Global Positioning System (GPS)
livewell and/or buckets
measuring board (millimeters)
wooden bat
aluminum foil (solvent rinsed)
polyethylene tubing (food-grade)
large plastic (composite) bags
coolers
plastic cable ties
dry ice (for preservation) or wet ice (for temporary transport)
Human Health Fish Collection form
human health fish tissue sample labels
pencils (for data forms)
fine-tipped indelible markers (for labels)
Tyvek label tag with grommet
clear tape strips
Tracking: Human Health Whole Fish Sample — Overnight (Dry Ice) form
FedEx airbill (pre-addressed)
cooler
dry ice (50 Ibs per cooler)
packing/strapping tape
13.3.3 SAMPLING PROCEDURE
Note: Do not handle any food, drink, sunscreen, or insect repellent until after the
composite sample has been collected, measured, and wrapped (or implement measures to
reduce contamination by such chemicals if applied such as washing, wearing long gloves,
etc.).
1. Put on clean nitrile gloves before handling the fish.
2. Rinse potential target species/individuals in ambient water to remove foreign
material from the external surface and place them in clean holding containers
(e.g., livewells, buckets).
3. For each human health fish tissue sample composite, select five whole fish of
adequate size to provide a total of 500 grams of fillet tissue. Criteria for
inclusion in the human health fish tissue composite:
a) All fish are of the same primary target species or secondary fish species
(See Table 13.11)
Note: It is essential that field crews accurately identify the organisms
submitted for analysis. Do not submit organisms from different species in a
single sample.
b) All fish are adult fish; and
c) All fish are of similar size, so that the smallest individual in a composite is
no less than 75% of the total length of the largest individual.
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4. Measure each fish selected for the composite from the anterior-most part of
the fish to the tip of the longest caudal fin ray (when the lobes of the caudal
fin are depressed dorsoventrally) to determine total body length in millimeters.
5. On the Human Health Fish Collection (Back) form:
• Record the sample identification number.
• Fill in the circles verifying that all samples are of similar length and the
same species.
• Below the header, record species selected for analysis, specimen length
(total length in mm), and any relevant comments. Extra rows are provided
on the form in the event that additional specimens are collected to meet
the 500 gram fillet tissue requirement (refer to Frequently Asked
Questions for further clarification).
• Make sure the sample ID and specimen numbers recorded on the form
match those on the sample labels.
6. Wearing clean nitrile gloves, remove each fish selected for analysis from the
clean holding container(s). Dispatch each fish using a clean wooden bat (or
equivalent wooden device).
7. Wrap each whole fish in extra heavy-duty aluminum foil, with the dull side in
contact with the fish (foil will be provided by EPA as solvent-rinsed, oven-
baked sheets).
8. Prepare a sample label for each sample specimen, ensuring that the label
information matches the information recorded on the Human Health Fish
Collection (Back) form. Be sure to record the fish species and specimen
length on each label.
9. Cut separate lengths of food grade tubing (provided by EPA) long enough to
contain each individual fish, allowing extra length on each end to seal with
cable ties. Place each foil-wrapped specimen into the appropriate length of
tubing. Seal the ends of each tube with a plastic cable tie. Attach the
appropriate sample label to the plastic tubing by wrapping clear tape around
the label and then completely around the wrapped fish (so that the clear tape
wraps over itself).
10. Double-bag the entire set of specimens in the composite by placing all fish
composited from the site inside a large plastic bag (provided by EPA). If
additional bags are required for large fish specimens or fish samples, please use
plastic bags of similar thickness as those provided by EPA.
11. Prepare a Sample Identification Label for the outer bag, ensuring that the label
information matches the information recorded on the Human Health Fish
Collection (Back) form. Be sure to record fish species and specimen length
range on the label.
12. Affix the sample label to a composite bag tag (Tyvek tag) and cover with clear
plastic tape. Thread a cable tie through the grommet in the tag and seal the
outer bag with the cable tie.
13.3.4 SAMPLE STORAGE AND SHIPPING PREPARATION
1. After the sample is packaged, immediately place it on dry ice for shipment.
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• Packaged samples may be placed on wet ice in coolers if they will be
transported to a laboratory or other interim facility to be frozen before
shipment.
• Samples may be stored on dry ice for a maximum of 24 hours.
• If possible, keep all specimens designated for a particular composite in the
same cooler for transport.
2. Crews have two options for freezing and shipping fish tissue samples,
depending on site logistics:
a) Ship the samples via priority overnight delivery service (e.g., Federal
Express), packed on dry ice, so that they arrive at the sample preparation
laboratory within 24 hours from the time of sample collection. Do NOT ship
on Fridays, Saturdays, or the day before federal holidays. Samples must be
packed on sufficient dry ice (50 pounds minimum, layered to ensure direct
contact between fish and dry ice) to keep them frozen for up to 48 hours.
Remember to record the tracking number on the sample tracking form
before submitting it to the Information Management Center.
b) Freeze the samples within 24 hours of collection at <-20°C, and store the
frozen samples until shipment within 2 weeks of sample collection. Frozen
samples will subsequently be packed on at least 50 pounds of layered dry
ice and shipped to the sample preparation laboratory via priority overnight
delivery service. Refer to reminders in option 2a (above) about not shipping
on Fridays, Saturdays, or the day before federal holidays and about
including sample tracking numbers on tracking forms.
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Table 13.11 Primary and secondary Great Lakes target species for human health fish tissue collection
PRIMAF
FAMILY
Centrarchidae
Cyprinidae
Esocidae
Ictaluridae
Gadidae
Percidae
Sciaenidae
SECOND;
Centrarchidae
Ictaluridae
IY HUMAN HEALTH FISH TISSUE T/
SCIENTIFIC NAME
Ambloplites rupestris
Micropterus dolomieu
Micropterus salmoides
Pomoxis annularis
Pomoxis nigromaculatus
Cyprinus carpi o
Esox lucius
Esox masquinongy
Esox niger
Ictalurus punctatus
Lota lota
Morone americana
Morone chrysops
Perca flavescens
Sander canadensis
Sander vitreus
Coregonus clupeaformis
Oncorhynchus gorbuscha
Oncorhynchus kisutch
Oncorhynchus tshawytscha
Oncorhynchus mykiss
Salmo salar
Sal mo trutta
Salvelinus namaycush
Aplodinotus grunniens
\RY HUMAN HEALTH FISH TISSUE 1
SCIENTIFIC NAME
Carpiodes cyprinus
Catostomus catostomus
Catostomus commersonii
Hypentelium nigracans
Ictiobus cyprinellus
Ictiobus niger
Lepomis cyanellus
Lepomis gibbosus
Lepomis gulosus
Lepomis macrochirus
Lepomis megalotis
Ameiurus melas
Ameiurus natalis
Ameiurus nebulosus
Coregonus artedi
Coregonus hoyi
Prosopium cylindraceum
Salvelin us font in alis
^RGET SPECIES
COMMON NAME
Rock bass
Smallmouth bass
Largemouth bass
White crappie
Black crappie
Common carp
Northern pike
Muskellunge
Chain pickerel
Channel catfish
Burbot
White perch
White bass
Yellow perch
Sauger
Walleye
Lake whitefish
Pink salmon
Coho salmon
Chinook salmon
Rainbow trout
Atlantic salmon
Brown trout
Lake trout
Freshwater drum
FARGET SPECIES
COMMON NAME
Quillback
Longnose sucker
White sucker
Northern hogsucker
Bigmouth buffalo
Black buffalo
Green Sunfish
Pumpkinseed
Warmouth
Bluegill
Longear Sunfish
Black bullhead
Yellow bullhead
Brown bullhead
Cisco/ lake herring
Bloater
Round whitefish
Brook trout
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14 FINAL SITE ACTIVITIES
After sampling, crews complete a visual site assessment and, upon return to the launching
location, the field crew must perform a post-measurement calibration check of the multi-
parameter sonde, review all data forms and labels, inspect samples, complete tracking
forms, ship or store samples, submit tracking forms, submit data forms, clean sampling
equipment, and inventory supplies. Activities described in this section are summarized in
Figure 14.1.
COMPLETE SITE
ASSESSMENT
REVIEW DATA FORMS
(Crew Leader)
• Completeness
• Accuracy
• Legibility
• Flags/Comments
REVIEW SAMPLE LABELS
(Crew Leader)
• Completeness
• Accuracy
• Legibility
• Cross-check with forms
PACK EQUIPMENT AND
SUPPLIES FOR TRANSPORl
FILTER, PRESERVE, &
INSPECT SAMPLES
Complete
Sealed
Ice packs
Packed for transport
INSPECT BOAT, MOTOR,
TRAILER, AND NETS FOR
PRESENCE OF PLANT AND
ANIMAL MATERIAL, AND
CLEAN THOROUGHLY
LOAD BOAT ONTO TRAILED
CLEAN UP LAUNCH SITE
AND STAGING AREA
> LEAVE SITE *
COMMUNICATIONS
SHIP SAMPLES
Figure 14.1 Final site activities summary
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14.1 GENERAL SITE ASSESSMENT
After sampling, complete the Site Assessment (Front) form which is a template for
recording pertinent field observations. Record all observations from the site that were
noted during the course of the visit. The Site Assessment (Front) form is by no means
comprehensive, and crews are encouraged to record any additional pertinent observations
in the General Assessment section.
14.1.1 SHORELINE ACTIVITIES AND DISTURBANCES
Rank shoreline activities and disturbances at the site. Consider only the shoreline that is
ecologically significant to, adjacent to, and visible from the X-site. Do not consider the
shoreline that is not in the same estuary, waterbody and/or embayment as the X-site. If
the shore cannot be seen from the X-site (due to weather conditions or distance), note in
the comments section the reason that the shoreline assessment was not possible. If an
activity or disturbance is present, fill in the appropriate bubble: "L" for low, "M" for
medium or "H" for high indicating the level of each.
Note: If an activity or disturbance is not observed, do not fill in any bubble. Also be sure
to fill in the 'super bubble' at the top the activities and disturbances section to verify
that blank fields indicate absence of the specific type of activity or disturbance.
14.1.2 SITE CHARACTERISTICS
Record the general characteristics of the site. When assessing site characteristics, look at
a 200 m radius around the X-site. Rank the site on a scale of 1 to 5, with 1 indicating
"pristine" or "appealing" and 5 indicating "highly disturbed" or "unappealing." As with
other aspects of the general visual assessment, all crew members contribute to the final
ranking. Observations of site characteristics will be understandably subjective, but
provide valuable information on crew impressions of the overall character of the site. The
NCCA analysts use crew observations to help explain data and results. The assessment of
visible trash in water (aquatic trash) will provide data for the U.S. EPA's Trash Free
Waters Program. If any items listed are visible in the water from the X-site, fill in a
bubble estimating the amount each type of trash. If none are visible, leave the bubbles
empty. If possible, list "Other plastic items", types of "Fishing gear" and "Other" items
not accounted for above. Additional information on aquatic trash may be written in the
General Assessment area at the crew's discretion. Document dominant land use. If
dominant land use is "forest," estimate the age class. Document the weather conditions
on the day of sampling, as well as any extreme weather conditions just prior to sampling.
Note: If there is no land within 200 meters of the X-site, leave the dominant land use
section blank.
14.1.3 GENERAL ASSESSMENT
Record any additional information and observations in this narrative section. Include
observations on biotic integrity, presence of SAV, presence and abundance of endangered
and/or exotic species, local anecdotal information, or any other pertinent information
about the site or its adjacent areas. Record any observations that may be useful for future
data interpretation.
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14.2 PROCESSING THE FECAL INDICATOR
14.2.1 SUMMARY OF METHOD
At each site, crews collect and filter water samples for fecal indicator analyses. Upon
receipt of the filters, the lab uses quantitative polymerase chain reaction (qPCR) analysis
to quantify Enterococci bacteria trapped on the filter.
14.2.2 EQUIPMENT AND SUPPLIES
Table 14.1 provides the equipment and supplies needed for field crews to filter the fecal
indicator sample. The filtering apparatus for this indicator MUST be sterile (i.e. a new
unused filter funnel with pre-loaded filter is used for each filtration). Because some
implements (forceps, centrifuge tube, etc.) will be reused for the filtering of the
chlorophyll sample, Enterococci must be filtered before filtering chlorophyll-a samples.
Table 14.1 & supplies: Enterococci processing
For processing samples nitrile gloves
sterile screw-cap graduated 50 mL centrifuge tube (for measuring sample)
filter flask (500 mL with side arm, labeled for ENTE only)
rubber stopper (#8 white, with 10mm hole) and small filter funnel adapter
2 filtration units (white base, sterile 100 mL units, includes pre-loaded filter for
ENTE) + 1 extra for revisit sites
vacuum pump (electric or hand)
sterile phosphate buffer solution
2 sterile disposable forceps
2 sterile microcentrifuge tubes containing sterile glass beads (chilled on dry ice
during pre-sampling activities) + 1 extra for blank filter (at revisit sites)
bubble bag (3 microcentrifuge tubes at revisit sites; 2 at all other sites)
dry ice
cooler
For recording Sample Collection form
measurements pencils (for data forms)
fine-tipped indelible markers (for labels)
fecal indicator sample labels (2 vial labels and 1 bag label)
clear tape strips
14.2.3 PROCESSING PROCEDURE - FECAL INDICATOR FILTER BLANK
At revisit sites (sites that will be visited twice in the index period for quality assurance
purposes), not only do crews filter the Enterococci samples, but they also prepare a filter
blank to be sent to the lab for analysis during both Visit 1 and Visit 2. A filter blank is
prepared prior to filtering the Enterococci sample. See below for filter blank field
processing procedure.
1. Put on nitrile gloves.
2. Set up the sample filtration apparatus on a flat surface and attach the vacuum
pump (Figure 14.2). Set out:
a. 50 ml sterile centrifuge tube,
b. 1 bottle of chilled phosphate buffer solution (PBS),
c. 2 sterile forceps.
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3. Attach the filter funnel with pre-loaded sterile filter to the filtering flask with
reusable rubber stopper and adapter.
4. Measure 20 ml of the chilled PBS with the sterile graduated centrifuge tube
and pour into the filter funnel.
5. Replace the cover on the filter funnel and use the vacuum pump to generate a
vacuum of no more than 7 inches of Hg (or -3.4 psig). Keep pumping until all
liquid is in filtrate collection flask.
6. Remove the filter funnel from the base without disturbing the filter. Using
sterile disposable forceps remove the filter (touching only the filter edges) and
fold it in half, in quarters, in eighths, and then in sixteenths (filter will be
folded 4 times).
7. Insert the filter into the chilled microcentrifuge tube (with beads) open end
first (pointed end up). Replace and tighten the screw cap.
8. Record the filter blank information on the Sample Collection (Front) form.
9. Prepare a sample label [Filter: Blank] by recording the volume of PBS filtered.
10. Affix the sample label to the microcentrifuge tube. Do NOT place tape on
either the label or the cap of the microcentrifuge tube.
11. Insert the tube into the bubble envelope. Place the bubble envelope on dry ice
while waiting to process the remaining filters.
12. Proceed to Section 14.2.4 for processing the water sample collected for
Enterococci.
14.2.4 PROCESSING PROCEDURE - FECAL INDICATOR SAMPLE
The filtering apparatus must be sterile when filtering the fecal indicator sample. A
separate, sterile, filter funnel pre-loaded with a filter will be provided for each sample
collected and processed. Crews must filter and freeze the fecal indicator sample within 6
hours of collection. See below for field processing procedures.
1. Put on nitrile gloves.
2. Set up the sample filtration apparatus on a flat surface and attach the vacuum
pump (Figure 14.2). Set out:
a) 50 ml sterile centrifuge tube,
b) 1 bottle of chilled PBS,
c) 2 sterile forceps.
3. Attach the filter funnel with pre-loaded sterile filter onto the filtering flask
with reusable rubber stopper and adapter.
4. Shake the sample bottle 25 times to mix well.
5. Using the 50 ml sterile graduated centrifuge tube, measure 25 ml of the mixed
water sample and pour into the filter funnel.
6. Replace the cover on the filter funnel. Use the vacuum pump to generate a
vacuum of no more than 7 inches of Hg (or -3.4 psig). Keep pumping until all
liquid is in the filtrate collection flask.
7. If the first 25 ml volume passes readily through the filter, add another 25 ml
and continue filtration. If the filter clogs before completely filtering the first or
second 25 ml volume, discard the filter and, using a new sterile filter funnel
with pre-loaded filter, repeat the filtration using a lesser volume.
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8.
9.
10
11
Pour approx. 10 ml of the chilled PBS into the same graduated centrifuge tube
used for measuring the water sample. Cap the tube and shake 5 times. Remove
the cap and pour the rinse into the filter funnel to rinse the filter.
Filter the rinsate and repeat with another 10 ml of chilled PBS.
Remove the filter funnel from the base without disturbing the filter. Using
sterile disposable forceps remove the filter (touching only the filter edges) and
fold it in half, in quarters, in eighths, and then in sixteenths (filter will be
folded 4 times).
Insert the filter into the chilled microcentrifuge tube (with beads)—open end
first (pointed end up). Replace and tighten the screw cap.
12. Record the volume of water sample filtered through the filter (minimum is 25
ml, target is 50 ml) and the volume of PBS used to rinse each filter on the
Sample Collection (Front) form. Record the filtration start time (beginning of
first filter) and finish time (end of second filter) for the sample.
13. Prepare a corresponding sample label (Filter:1 or Filter:2), ensuring that the
volume filtered on the label matches the information recorded on the Sample
Collection (Front) form.
14. Affix the sample label to the microcentrifuge tube. Do NOT place tape on
either the label or the cap of the microcentrifuge tube.
15. Insert the tube into the bubble envelope. Place the bubble envelope on dry ice
while processing the second filter.
16. Repeat steps 1 to 15 for the second filter, using a new sterile filter funnel with
pre-loaded filter. It is important that the same sample volume be filtered
through each filter.
17. Prepare an exterior label for the bubble envelope [ENTEROCOCCI (ENTE) -
BAG], ensuring that the label information (site ID, date, visit #, volume
filtered, sample ID) matches the information recorded on the Sample
Collection (Front) form. Affix the exterior label on the outside of the bubble
envelope and cover with clear plastic tape.
18. Place the bubble envelope in a 4 by 4 self-sealing bag and then on dry ice for
preservation during transport and shipping.
Sterile 100mL _
filter funnel with
pre-loaded filter
(single use)
Reusable
filtering
funnel
Rubber stopper (white)
and small funnel adapter
Vacuum pump
(hand or electric)
Figure 14.2 Filtering set-up for Enterococci filtering
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14.3 PROCESSING THE CHLOROPHYLL-^ & DISSOLVED NUTRIENTS INDICATORS
14.3.1 SUMMARY OF METHOD
At each site, crews collect and filter water samples for chlorophyll-a and dissolved
nutrient analyses. The chlorophyll-a sample is submitted to the lab as residue on a
Whatman GF/F filter. Upon receipt of the filters, the lab extracts the pigment from the
filter and quantifies it using flourometry. A portion of the filtrate produced from
collecting the chlorophyll-a sample is submitted to the laboratory and processed for
dissolved nutrients. In order to avoid cross-contamination, a new filter funnel will be used
at each site. This filter funnel is provided in each site kit.
14.3.2 EQUIPMENT AND SUPPLIES
Table 14,2 Equipment fi supplies: chlorophyll-a ft dissolved nutrients processing
For filtering \X1iatman GF/F 47mm 0.7 micron filter
chlorophyll-a Nutrients filtering chamber OR 500 mL side-arm filter flask, labeled for
sample CHLA/NUTS only)
Filtration unit (blue base filter funnel, 250 mL unit)
rubber stopper (#8 blue, with 15mm hole) and large filter funnel adapter vacuum
pump (electric or hand)
DI water
nitrile gloves
forceps
graduated cylinder (250 mL)
For recording Sample Collection form
measurements chlorophyll-^ & dissolved nutrients sample labels
pencils (for data forms)
fine-tipped indelible markers (for labels)
clear tape strips
For sample centrifuge tube (50 mL, screw-top)
collection and aluminum foil square
preservation HDPE bottle (250 mL, white)
cooler with dry ice
electrical tape
plastic bag (sandwich size)
14.3.3 PROCESSING PROCEDURE
Below presents the field procedures for processing chlorophyll-a and dissolved nutrient
samples. The steps below describe using the nutrients filtering chamber supplied in the
base kit. Crews have the option of using a side-arm filtering flask or other filtrate
collection device in place of the nutrients chamber. If a flask or other device is used, it is
important to NOT use the same flask/device as is used for the filtering of Enterococci.
Doing so will lead to potential contamination of the nutrients sample with phosphate
buffer used to rinse the Enterococci filter. If a flask or other filtrate collection device is
used to collect the filtered nutrients sample (as opposed to collecting the sample directly
into the nutrients bottle with a chamber), the collection device must be rinsed three
times with filtered sample water before allowing any sample to enter the bottle.
Note: Crews must make every attempt to process chlorophyll-a samples in subdued light,
out of direct sunlight.
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1. Complete the NUTS sample label with Site ID, date collected, and visit number.
2. Attach the completed label to the 250 ml clear HOPE sample bottle and cover
with clear plastic tape.
3. Set up the nutrients filtering chamber on a flat surface, insert the sample
bottle into the chamber and attach the vacuum pump (Figure 14.3)
4. Put on nitrile gloves.
5. Crews will use a 250 ml filter funnel (with blue bottom), rubber stopper, and
adapter that are specifically designated for chlorophyll filtering (i.e. not the
same ones used for the Enterococci filtering). A new filter funnel will be
provided in each site kit and should not be reused. The stopper and adapter are
to be cleaned between sampling events. Prior to filtration of the sample, rinse
the filter funnel adapter three times with Dl water. Rinse graduated cylinders
with Dl water. After assembling the filtering apparatus and attaching the filter
funnel to the nutrients chamber with the correct stopper and adapter, remove
the cup portion of the filter funnel from the blue base. Remove the pre-loaded
filter (which has a faint grid pattern on it) but leave the white support pad in
place.
6. Use clean forceps to place a Whatman GF/F 47 mm 0.7 micron filter on the
support pad with the gridded/pressed side of the filter facing down, making
sure both the support pad and filter are centered on the base.
7. Reattach the funnel portion of the filter funnel to the base by pressing it
straight down firmly until it snaps into place. This will firmly hold the filter in
place.
8. Remove the 2 L amber chlorophyll-a collection bottle from cooler and shake to
mix the sample. Using the graduated cylinder, measure and pour 250 ml of
water into the filter holder, replace the cover, and use the vacuum pump to
draw a small portion of the sample through the filter. Do not exceed 7 inches
of Hg of vacuum -3.4 psig or a filtration duration of more than 5 minutes for a
single sample volume, to avoid cell damage or loss of contents during filtering.
9. Use the first 10-20 ml of filtrate to rinse the 250 ml sample bottle and discard
the rinsate. Be sure to cap the bottle and rotate it so that the filtered water
contacts all the surfaces. Replace the bottle and chamber cap and continue
filtering. Repeat the rinse of the sample bottle with an additional two rinses of
filtered site water then discard the rinsate.
10. If the filter clogs before 250 ml of site water will pass through the filter,
discard the filter and water remaining in the filter funnel, rinse the filter
funnel with Dl water, install a new filter, and repeat the procedures using 100
ml of site water.
11. Observe the filter for readily visible color. If there is visible color, proceed to
the next step; if not, filter additional aliquots until color is visible on the filter
or until a maximum of 2,000 ml have been filtered.
12. After collecting 250 ml of filtered site water in the dissolved nutrients sample
bottle, remove the 250 ml HOPE bottle. Replace the lid and seal tightly with
electrical tape. Submit this filtrate for dissolved nutrient analyses.
13. Move the filter funnel and adapter to a side-arm filter flask to complete the
filtering process. Additional filtrate will be discarded.
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14. Record the dissolved nutrients sample information on the Sample Collection
(Front) form. Place the sample on wet ice.
15. After achieving a readily visible stain on the filter and collecting the filtrate for
dissolved nutrient analyses, record the actual sample volume filtered in the
Chlorophyll-a section on the Sample Collection (Front) form and on the sample
label.
16. Attach the completed label to the 50 ml centrifuge tube and cover with clear
plastic tape.
17. Rinse the graduated cylinder and upper portion of the filter funnel thoroughly
with Dl water to include any remaining cells adhering to the sides and pump
through the filter. Monitor the level of water in the lower chamber to ensure
that it does not contact the filter or flow into the pump.
18. Remove the filter from the holder with clean forceps. Avoid touching the
colored portion of the filter. Fold the filter in half, with the colored side folded
in on itself. Place the folded filter into the 50 ml screw-top centrifuge tube
used previously for measuring the Enterococci sample and replace the cap.
19. Tighten the cap as tightly as possible. The cap will seal tightly after an
additional 1/4 turn past the point at which initial resistance is met. Failure to
tighten the lid completely could allow water to infiltrate into the sample and
may compromise its integrity. Seal the cap of the centrifuge tube with
electrical tape.
20. Wrap the 50 ml tube in a foil square and place in the provided self-sealing
plastic bag.
21. Close the plastic bag and place it on dry ice.
Note: if the chlorophyll filtering process did not yield at least 250 mL of filtered site
water, install a new GF/F filter and continue filtering site water until 250 mL of
filtrate has been collected for the dissolved nutrients sample. Be sure to collect the
filtrate prior to any rinsing of the filter funnel with Dl water as directed in Step 17.
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Filter funnel
(250 ml with blue base)
Rubber stopper (Blue) and '
Large funnel adapter
Chamber Cap
Vacuum Pump
(Hand or Electric)
250 ml Nutrients Bottle
Nutrients Filtering Chamber
Figure 14.3 Filtering set-up for chlorophyll-a and nutrients filtering
14.4 POST-MEASUREMENT CALIBRATION CHECK OF MULTI-PARAMETER SONDE
After all in situ measurements have been completed for the sampling day, the crew must
perform a post-measurement calibration check of the multi-parameter sonde. To do this,
measure the pH and conductivity of one of each of the respective calibration standards
that were used earlier in the day to calibrate the instrument. Record these values in the
Post-Measurement Calibration Check section on the Field Measurement (Front) form. If
significant drift is detected as defined the manufacturer, the meter may need service and
data collected since the last successful calibration and post-measurement calibration
check should be flagged. Discontinue use of any meter that is not functioning properly.
14.5 FIELD DATA & TRACKING FORM REVIEW
The Field Crew Leader is ultimately responsible for reviewing the App submission and/or
all data forms for completeness, legibility, accuracy, and consistency. The following are
some checks to perform on the data forms:
• Ensure that all required data forms for the site have been completed.
• Confirm that the Site ID, visit number, and date of visit are correct on all
forms.
• Verify the accuracy and legibility of all recorded information.
• Ensure that any flags are explained in the respective comments sections.
• Ensure that written comments are clear, with no "shorthand" or
abbreviations.
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• Make sure there are no stray markings on the forms. (Field forms are
scanned and read by optical character recognition (OCR) software. Stray
marks often lead to erroneous data recording and must be hand checked
and corrected when discrepancies occur.)
• Make sure the header information is completed on all pages of each form.
• After reviewing each form, initial the upper right corner of each page of
the form.
If information is missing from the forms, the Field Crew Leader must complete the missing
sections. If utilizing paper forms, upon completing the review, the Field Crew Leader must
initial the field forms, indicating that they are complete, legible, accurate, consistent,
and ready to be sent to NARS IM. If utilizing the NARS App, the Field Crew Leader must
submit the data. The receipt of a submission is a confirmation that the data has been
reviewed by the Field Crew Leader.
14.6 SAMPLE PACKAGING AND LABEL REVIEW
All samples must be appropriately preserved and packaged for transport. The following
are some checks to perform on the labels:
• All samples are collected. If obtainable samples are missing, the crew must
reschedule a site visit or return to the site that same day to complete
collection of the missing samples.
• All samples are labeled.
• All labels are complete, legible, accurate, and consistent.
• Although the data forms, tracking forms, and labels are preprinted with the
sample IDs, review the labels and forms to ensure consistent sample ID
information was utilized.
• Each label is covered with clear plastic tape (except those on the ENTE
sample vials).
• Inspect the integrity of each sample container; be sure there are no leaks.
Make sure that all sample containers are properly sealed.
• Verify that all sample containers are properly preserved for storage or
immediate shipment.
If information is missing from the labels, the Field Crew Leader must complete the missing
sections. The Field Crew Leader must also verify the integrity of all samples. The Field
Crew Leader must reconcile any disagreements between sample IDs on the data
forms/NCCA App and labels before tracking forms are transmitted to NARS IM and samples
are packaged and sent to the labs.
14.7 SAMPLE SHIPMENT & TRACKING FORM SUBMITTAL
Refer to Appendix C: Shipping and Tracking Guidelines for additional details on preparing
samples for shipping.
14.7.1 TIME-SENSITIVE SAMPLES
The field crew must ship or deliver time-sensitive samples (i.e., water chemistry (CHEM),
chlorophyll-a (CHLA), and dissolved nutrients (NUTS)) to the appropriate analytical
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laboratory (WRS Corvallis or approved state lab) so that the samples will arrive within 48
hours of collection. Therefore, crews must send them via Priority Overnight shipping,
preferably the same day as collection, but no later than the following day. Reminder:
FedEx does not deliver shipments on Sunday, so you must ensure samples are shipped
by Friday afternoon to allow for a Saturday delivery. Be sure to verify the last
EXPRESS drop off time at the FedEx facility you plan to use.
The Field Crew Leader will complete a Site and Sample Status/Water Chemistry lab
tracking form for the samples and will email the form to sampletracking@epa.gov or
submit tracking via the NCCA App (other submittal options are provided in Section 15.3).
Please name these files in the following format: NCCA15_T#_Tracking_SitelD_V#, where
T#' is the number of the tracking form and W is the visit number (i.e.
NCCA15_T1_Tracking_NCCA15-1061_V1). If scanning paper forms, be sure that the file
scanned is clear and legible. Genius Scan or Cam Scanner are great apps that are available
for free that will help to ensure that the scan is clear and legible.
The Field Crew Leader will place the samples and Site and Sample Status/Water Chemistry
lab tracking form (in a waterproof bag or plastic sleeve) in the cooler provided with the
site kit. The Field Crew Leader will attach the appropriate pre-addressed FedEx airbill
from the site kit marked for the WRS lab. The field crew will either drop off the cooler for
shipment at a local FedEx location or arrange for a pick up at the hotel or other
appropriate facility. If the field crew has chosen a pick up, they must follow up with the
facility at which it has been left to ensure its actual pick up.
14.7.2 OTHER SAMPLES
Samples that are less time sensitive will be shipped in batches, according to the chart in
Appendix C: Shipping and Tracking Guidelines. See Section 15: Post-Sampling Activities
for further guidance.
14.8 EQUIPMENT CLEANUP & CHECK
Field crews must take appropriate precautions to avoid transfer of national and regional
invasive species of concern. Nuisance species of concern in the U.S. include zebra mussels
(Dreissena polymorpha), mitten crabs (Eriocheir sinensis) and Eurasian ruffe
(Gymnocephalus ceinuus). In the Great Lakes, Viral Hemorrhagic Septicemia (VMS) is an
invasive and deadly fish virus that is threatening Great Lakes fish. VMS was identified as
the cause of large fish kills in lakes Huron, St. Clair, Erie, Ontario and the St. Lawrence
River in 2005 and 2006. To reduce the risk of transferring nuisance species and pathogens,
all equipment and gear must be cleaned and disinfected prior to traveling over land from
one field site to another. For specific techniques to disinfect boats and gear in the Great
Lakes, please see Section 14.8.3.
Online resources regarding invasive species:
• Aquatic Nuisance Species Task Force (http://www.anstaskforce.gov)
• U.S. Geological Survey Nonindigenous Aquatic Species website
(http://nas.er.usgs.gov)
• Protect Your Waters website, co-sponsored by the U.S. Fish and Wildlife
Service (http://www.protectyourwaters.net/hitchhikers)
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• Sea Grant Program (http://www.sgnis.org)
• USDA Animal and Plant Health Inspection Service (http://aphis.usda.gov)
14.8.1 BOAT & TRAILER CLEANUP
While your organizations likely have protocols in place to account for these precautions,
the following are some procedures and checks to perform on your equipment:
1. Load the boat on the trailer.
2. Drain all bilge water from the boat.
3. Inspect the boat, motor, and trailer for evidence of weeds and other
macrophytes.
4. Clean the boat, motor, and trailer as completely as possible before leaving the
launch site.
• Follow any state or other requirements associated with nuisance species,
pathogens and/or viruses.
14.8.2 POST SAMPLING EQUIPMENT CARE
1. Inspect sampling gear (seines, dip nets, sieves, foul weather gear, boots, etc.)
for evidence of mud, snails, plant fragments, algae, animal remains or debris.
Rinse and remove using brushes or other tools. Use one of the procedures
below to disinfect gear if necessary. Let dry.
2. Pack all equipment and supplies in the vehicle and trailer for transport.
3. Keep equipment and supplies organized so they can be inventoried using the
equipment and supply checklists (Appendix A: Equipment and Supplies Lists).
4. Clean up all waste material at the launch site and dispose of or transport it out
of the site if a trash can is not available.
14.8.3 ADDITIONAL DECONTAMINATION INFORMATION
Additional precautions to prevent transfer of Whirling Disease spores, New Zealand
mudsnails, and amphibian chytrid fungus are important for Great Lakes sites. Before
visiting the site, research the site and determine if it is in an area where one of these
organisms are known to exist. Contact the local or State fishery biologist to confirm the
presence or absence of these organisms.
If the site is listed as "positive" for any of the organisms, or no information is available,
avoid using felt-soled wading boots. After sampling, disinfect all fish and benthos
sampling gear and all other equipment that came into contact with water or sediments
(i.e., waders, boots, etc.) by one of the following procedures:
Option A:
1. Soak gear in a 10% household bleach solution for at least 10 minutes, or wipe or
spray on a 50% household bleach solution and let stand for 5 minutes.
2. Rinse with tap water (do not use sea or lake water) and remove remaining
debris.
3. Place gear in a freezer overnight, soak in a 50% solution of Formula 409®
antibacterial cleaner for at least 10 minutes or soak gear in 120°F (49°C) water
for at least 1 minute.
4. Dry gear in direct sunlight (at least 84 °F) for at least 4 hours.
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Option B:
1. Soak gear in a solution of Sparquat® (4-6 oz. per gallon of water) for at least 10
minutes (Sparquat is especially effective at inactivating whirling disease
spores).
2. Place gear in a freezer overnight or soak in 120°F (49° C) water for at least 1
min.
3. Dry gear in direct sunlight (at least 84 °F) for at least 4 hours.
Clean and dry other equipment prior to storage.
• Rinse coolers with clean water to remove any dirt or debris on the outside
and inside.
• Make sure water quality meter probes are rinsed with deionized water and
stored moist.
• Rinse all equipment used to collect water samples three times with
deionized water. Place sampling equipment in a clean location for use at
the next site.
• Check nets for holes and repair or locate replacements.
• Inventory equipment and supply needs and relay orders through the fillable
PDF Supply Request form.
• Remove GPS and multi-parameter sonde, and set up for pre-departure
checks and calibration. Examine the oxygen membranes for cracks,
wrinkles, or bubbles. Replace if necessary, allowing sufficient time for
equilibration.
• Recharge/replace batteries as necessary.
• Replenish fuel and oil.
• If a commercial car wash facility is available, thoroughly clean vehicle and
boat (hot water pressurized rinse—no soap).
Note: Handle and dispose of disinfectant solutions properly, and take care to avoid
damage to lawns or other property.
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15 POST-SAMPLING ACTIVITIES
15.1 SAMPLE SHIPPING
Samples that are less time sensitive will be shipped in batches, according to the chart in
Appendix C: Shipping and Tracking Guidelines. The Field Crew Leader will complete the
appropriate batch tracking form(s) for the samples and will email electronic copies of the
form(s) to sampletracking@epa.gov or submit tracking via the NCCA App (other submittal
options are provided in Section 15.3). Please name these files in the following format:
NCCA15_T#_Tracking_SitelD_V#, where T#' is the number of the tracking form and W is
the visit number (i.e. NCCA15_T1_Tracking_NCCA15-1061_V1).
The Field Crew Leader will place the samples and the correct batch tracking form (in a
waterproof bag or plastic sleeve) in a requested batch shipment cooler. The Field Crew
Leader will attach the appropriate pre-addressed FedEx airbill from the site kit marked
for the appropriate lab. The field crew will either drop off the cooler for shipment at a
local FedEx location or arrange for a pick up at the hotel or other appropriate facility. If
the field crew has chosen a pick up, they must follow up with the facility at which it has
been left and/or track the package through FedEx tracking tools to ensure its actual pick
up. Once the package is in the possession of FedEx, the IM Team and FLC will track the
package to its destination and take steps necessary to ensure its timely delivery.
15.2 TRACKING FORM SUBMITTAL
Each tracking form has been assigned a "T" page number to help crews identify the
correct tracking form to use when sending samples. This "T" number is located on the
bottom right corner of each tracking form. Crews will also find reference to the same "T"
numbers on the individual samples labels and on the top of the pre-printed FedEx return
labels provided in the site kits.
Crews include copies of all tracking forms in the coolers when they send samples to the
labs. They have several different options for electronically submitting sample and tracking
information. A hard copy of the sample tracking form must be submitted to the lab in the
cooler and an electronic copy must be submitted to NARS IM using one of four options. If
a cooler contains samples from more than one site, then multiple forms must be placed in
the cooler and submitted to NARS IM.
In order of preference, the options are:
1. Using the NCCA mobile App, enter data and tracking information into the NCCA
App and submit the tracking information. An email will pop up on your device
with an attachment and the NARSFieldData@epa.gov address. Copy yourself,
any other crew members or managers, and click send. This form will be
returned to you via email after a few minutes in a portable document format
(PDF). It may be printed and used as the form for the cooler shipment.
2. Using a handheld device or portable computer, enter data into fillable portable
document format (PDF) forms and submit. Please name these files in the
following format: NCCA15_T#_Tracking_SitelD_V#, where T#' is the number
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of the tracking form and W is the visit number (i.e.
NCCA15_T1_Tracking_NCCA15-1061_V1) before emailing or using the SUBMIT
button. Send the file via email to sampletracking@epa.gov. It may be printed
and used as the form for the cooler shipment.
3. Hand-enter data on a paper form. Photograph the form with a handheld device
or office scanner. Attach the file (in PDF version) to an email and address to
sampletracking@epa.gov. Please name these files in the following format:
NCCA15_T#_Tracking_SitelD_V#. Be sure that the file scanned is clear and
legible. Genius Scan or Cam Scanner are great apps that are available for free
that will help to ensure that the scan is clear and legible. Copy yourself any
other crew members or managers, and click send. After scanning, include this
form in the cooler.
4. Hand-enter data on a paper form. Fax the form to the number printed on the
form. After faxing include this form in the cooler.
If the crew visits a site with the intention of sampling, but determines the site to be
unsampleable (either temporarily or permanently), the site status portion of the Site and
Sample Status/Water Chemistry Lab Tracking form needs to be completed and
submitted, but the water chemistry and batch sample status tracking portion of the form
can remain blank. The Field Crew Leader must also submit the Site Verification (Front)
form for the site, which contains additional information about the site that is not
captured on the tracking form. This can be submitted with the packet of field forms that
gets sent out every two weeks. For ease of use, these two forms are available in fillable
PDF form on the EPA SharePoint site.
Regardless of the type of sample being shipped, a completed tracking form must be
placed inside the cooler with the samples (typically sealed in a plastic bag or pouch and
affixed to the inside of the cooler lid). Again, crews may choose to complete either the
digital or manual forms. In addition to sending the tracking forms with the shipment, a
copy of the tracking form must be submitted to the NARS IM staff at
sampletracking@epa.gov before the samples are due to arrive at the lab. The various
tracking forms are listed in Appendix C: Shipping and Tracking Guidelines.
15.3 DATA SUBMITTAL
15.3.1 APP USERS
For crews utilizing the mobile App, after the Field Crew Leader has reviewed form content
at the end of your sampling day, click the submit button. An email will pop up on your
device addressed to NARSFieldData@epa.gov. Copy yourself, any other crew members or
managers and click send.
15.3.2 PAPER FORM USERS
Every two weeks, the Field Crew Leader will batch the field data forms together and send
them to NARS IM. After checking the field data forms for completeness, legibility,
accuracy, and consistency, the Field Crew Leader will make scans or copies of them. The
Field Crew Leader will complete a Tracking: Packs form for the data packets and will
email that form to sampletracking@epa.gov. The Field Crew Leader will place the original
field data forms and batch tracking form in the FedEx envelope provided in the site kit.
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The Field Crew Leader will attach the pre-addressed FedEx airbill from the site kit, and
send the original forms to NARS IM via FedEx.
Note: The original forms are specially printed to be used in an optical scanner for
automated data entry. Copies of forms will not scan properly and are not acceptable for
entering field data. All field forms must be turned in within 2 weeks of completing
sampling. A tracking form will be submitted with each shipment of data forms and the
data forms will be tracked in the same manner as all other samples.
15.4 TRACKING REMINDERS
It is very important to submit the Site and Sample Status/Water Chemistry Lab Tracking
form immediately after every sampling event. Prompt status reports allow the FLC to
closely track sampling progress. More importantly, it enables NARS IM to track samples
that were collected at each site versus those that were not, and to immediately track the
shipment of the time-sensitive samples after each sampling event.
The field crews must promptly report any field sampling problems to the FLC and report
sample tracking or data reporting problems to NARS IM. They will follow up with the EPA
NCCA 2015 Lead throughout the sampling period.
The EPA Logistics Coordinator serves as the central point of contact for information
exchange among field crews, the management and QA staff, the NARS IM staff, and the
public. The EPA Logistics Coordinator and Contractor Field Logistics Coordinator contact
information can be found on Table 1.1 of this manual.
15.5 SITE EVALUATION SPREADSHEET SUBMITTAL
Throughout the field season or at the end of the field season, EPA HQ needs field crews to
submit their completed Site Evaluation Spreadsheets. These are critical to determining
site weights used in data analysis. Please submit these forms to the FLC and EPA Logistics
Coordinator within two weeks of completion of your last site.
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16 FIELD QUALITY CONTROL
The NCCA program requires that all cooperators and field crews follow strict quality
assurance and quality control guidelines. Standardized training and data forms set the
foundation to help ensure that data quality standards for field sampling are met. In
addition, repeat sampling and field evaluation and assistance visits address specific
aspects of the data quality standards for the NCCA.
16.1 STANDARDIZED TRAINING
All Field Crew Leaders must attend a formal three day NCCA training prior to participating
in field sampling for the NCCA and all field crew members are encouraged to attend. The
training, which is divided into classroom and hands-on field sessions, is designed to reduce
sampling variability, and subsequently ensure data comparability from crew to crew and
site to site. Standardized training allows the EPA to collect field crew input that will help
to identify potential sampling pitfalls and troubleshoot solutions. The entire three day
training session is required to qualify a crew for sampling activities.
16.2 STANDARDIZED FIELD DATA FORMS
All field crews, with the exception of crews collecting samples in the Great Lakes, collect
and record data using identical field forms. The Great Lakes has one additional two-sided
data form (D11 & D12) and two additional tracking forms to complete (T7 & T8). These
identical forms serve several purposes. First, they ensure that all crews measure and
record the same parameters. Second, use of identical field forms promotes efficient data
entry and minimizes the opportunities for data transcription errors. Finally, the use of
identical forms facilitates field form quality control reviews when data are received at
NARS IM.
Paper field forms and the NARS App have been developed for data collection and contain
the same data.
16.3 REPEAT SAMPLING
The NCCA collects temporal repeat samples in order to estimate site measurement and
index period variance. Repeat sampling provides data that can be used to evaluate the
potential for the NCCA design to estimate status and detect trends in the target site
population.
During the field season, crews will revisit approximately 10% of the target sites as
designated in the EPA site list with "RVT2" in the panel code. In order to ensure that
sampling procedures are as comparable as possible from the first visit to the second visit,
the same field crew who initially sampled the site also conducts the revisit. During site
revisits, crews collect the full set of samples and in situ measurement parameters (except
all fish tissue samples, which are targeted only on the first visit). At Great Lakes revisit
sites that are also human health fish tissue sites, crews that are unsuccessful at collecting
the human health fish tissue sample during visit 1 are expected to attempt the collection
of that sample during visit 2. When sampling sites are identified as revisit sites, crews
collect Enterococci filter blanks during both the initial visit and the revisit. The crews
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must always collect the filter blanks before the sample is filtered. See Section 14.2.3 for
the procedure for collecting filter blanks.
The NCCA identifies sites targeted for repeat visits in the state's site draw. The number of
repeat visit sites varies from state to state, depending on the number of base sites drawn
within the state. If a site selected for repeat sampling is dropped, then the alternate site
assigned to replace it becomes the revisit site. The time elapsed between the initial and
repeat site visits should be as long as possible within the index period, but not shorter
than two weeks.
16.4 FIELD EVALUATION AND ASSISTANCE VISITS
A rigorous program of field and laboratory evaluation and assistance visits supports the
quality assurance and control for the NARS. The following sections focus only on the field
evaluation and assistance visits.
By coupling assistance visits conducted early in the data collection process with uniform
training, sampling variability associated with specific implementation or interpretation of
the protocols will be significantly reduced. Field evaluation and assistance visits provide
an opportunity to ensure that crews follow field procedures and meet minimum quality
control requirements. In addition, assistance visits allow for uniform evaluation of the
standard NCCA data collection methods. When widespread problems or confusion surround
a given method, the information from assistance visits contributes to refining the method
for sites that are yet to be sampled and in future field manuals.
The field evaluators observe and review the information listed on the Field Evaluation and
Assistance Visit Checklist. An assistance visit has been scheduled to evaluate each unique
crew collecting and contributing data under this program. If unforeseen events prevent
the EPA from evaluating every crew, the NCCA Quality Assurance Coordinator (QAC) will
rely on the data review and validation process to identify unacceptable data that will not
be included in the final database. If inconsistencies cannot be resolved, the QAC may
contact the Field Crew Leader for clarification.
16.4.1 SPECIFICATIONS FOR QC ASSURANCE
Field evaluation and assistance personnel are trained in the specific data collection
methods detailed in this FOM. A plan and checklist for field evaluation and assistance
detail the methods and procedures that will be evaluated. The plan and checklist are
included as Attachment D in the QAPP and will be posted on the SharePoint site for crews
to access. Table 16.1 summarizes the plan, the checklist, and corrective action
procedures.
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Table 16,1 General information during field evaluation
Field
Evaluation
Plan
Field
Evaluation
and
Assistance
Visit
Checklist
Corrective
Action
Procedures
Regional Coordinators or another assigned trained individual arrange the field assistance visit
with each field crew, ideally within the first two weeks of sampling.
The Evaluator observes the performance of a crew through one complete set of sampling
activities.
If the crew misses or incorrectly performs a procedure, the Evaluator notes it on the checklist
and immediately points it out so the mistake can be corrected on the spot.
The Evaluator reviews the results of the evaluation with the field crew before leaving the site,
noting positive practices as well as problems.
The Evaluator observes all pre-sampling activities and verifies that equipment is properly
calibrated and in good working order, and that NCCA protocols are followed.
The Evaluator checks the sample containers to verify that they are the correct type and size, and
checks the labels to be sure they are correctly and completely filled out.
The Evaluator confirms that the field crew has followed NCCA protocols for locating the site.
The Evaluator observes the complete set of sampling activities, confirming that all protocols are
followed.
The Evaluator will record responses or concerns, if any, on the Field Evaluation and Assistance
Visit Checklist.
If the Evaluator's findings indicate that the field crew is not performing the procedures
correctly, safely, or thoroughly, the Evaluator must continue working with this field crew until
certain of the crew's ability to conduct the sampling properly and minimize adverse effects on
data quality.
If the Evaluator finds major deficiencies in the field crew operations, the Evaluator must
contact the NCCA QA Coordinator immediately (e.g., within 24-48 hours) so that additional
correction actions can be taken.
The EPA anticipates that evaluation and assistance visits will be conducted with each
Field Crew early in the sampling and data collection process, and that corrective actions
will be conducted in real time. The role of the Evaluator is to provide additional training
and guidance so that the procedures are being performed in a manner consistent with the
Field Operations Manual, all data are recorded correctly, and paperwork is properly
completed at the site. If the field crew misses or incorrectly performs a procedure, the
Evaluator will note the error on the checklist, immediately point it out and direct the
crew to correct it on the spot.
16.4.2 REPORTING
Upon completion of the sampling operations, the Evaluator will review the results of the
evaluation with the Field Crew before leaving the site (if practicable). The evaluator will
note positive practices and problems (termed weaknesses if they might affect data quality
or deficiencies if they would adversely affect data quality). The Evaluator ensures that all
crew members understand the findings and can perform the procedures properly in the
future. The Evaluator will record field crew responses or concerns, if any, on the Field
Evaluation and Assistance Visit Checklist. After the Evaluator completes the Field
Evaluation and Assistance Visit Checklist, including a brief summary of findings, all field
crew members must read and sign off on the evaluation.
If after directing the crew to correct problems, findings indicate that the field crew is not
performing the procedures correctly, safely or thoroughly, the Evaluator must continue
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working with this field crew until certain of the crew's ability to conduct the sampling
properly. If the Evaluator finds major deficiencies in the field crew operations (e.g.,
major misinterpretation of protocols, equipment or performance problems that will
adversely affect data quality), they must be reported to the following QA official:
• Hugh Sullivan, EPA NCCA QA Coordinator
The QAC official will contact the Project Manager to determine the appropriate course of
action. Data records from sampling sites previously visited by this field crew will be
checked to determine whether any sites must be resampled.
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17 LITERATURE CITED
American Red Cross. 2006. First Aid/CPR/AED for schools and the community. Third edition.
210 pgs.
Klemm, D. J., P. A. Lewis, F. Fulk, and J. M. Lazorchak. 1990. Macroinvertebrate Field and
Laboratory Methods for Evaluating the Biological Integrity of Surface Waters. EPA
600/4-90/030. U.S. Environmental Protection Agency, Cincinnati, Ohio.
National Institute for Occupational Safety and Health. 1981. Occupational Health Guidelines
for Chemical Hazards (Two Volumes). NIOSH/OSHA Publication No. 81-123.
U.S. Government Printing Office, Washington, D.C.
Occupational Safety & Health Administration (OSHA). 2006. Regulations (Standards - 29 CFR).
Substance technical guidelines for formalin - 1910.1048 App A. Occupational Safety &
Health Administration. Washington, DC 20210.
Schriver et al. 1995. Impact of Submerged Macrophytes on Fish-Zooplankton- Phytoplankton
Interactions - Large-Scale Enclosure Experiments in a Shallow Eutrophic Lake.
Freshwater Biology 33, no. 2: 255-70.
U.S. Coast Guard. 1989. Federal Requirements for Recreational Boats. U.S. Department
of Transportation, United States Coast Guard, Washington, D.C. 27 pgs.
USEPA. 2001. National Coastal Assessment: Field Operation Manual. EPA-620-R-01-003.
U.S. Environmental Protection Agency., Office of Research and Development, National
Health and Environmental Effects Research Laboratory.
USEPA. 2000a. EPA Quality Manual for Environmental Programs 5360A1. May 2000.
http://www.epa.gov/quality/qs-docs/5360.pdf
USEPA. 2000b. EPA Order 5360.1 A2 CHG2, Policy and Program Requirements for Mandatory
Agency-wide Quality System, May 5, 2000. http://www.epa.gov/quality/qs-docs/5360-
l.pdf
USEPA. 2001. Methods for Collection, Storage, and Manipulation of Sediments for Chemical
and Toxicological Analyses: Technical Manual. EPA-823-B-01-002. U. S. Environmental
Protection Agency, Office of Water, Washington, D.C.
USEPA. 2015. National Coastal Condition Assessment Quality Assurance Project Plan. EPA-841-
R-14-005. U.S. Environmental Protection Agency. Office of Water, Washington, DC.
Li-COR. 2006. LI-COR Underwater Radiation Sensors Instruction Manual: LI-192 Underwter
Quantum Sensor LI-193 Spherical Quantum Sensor. LI-COR, Inc., Lincoln, Nebraska.
Web Pages:
Aquatic Nuisance Species Task Force (http://www.anstaskforce.gov)
U.S. Geological Survey Nonindigenous Aquatic Species website (http://nas.er.usgs.gov)
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Protect Your Waters website, co-sponsored by the U.S. Fish and Wildlife Service
(http://www.protectyourwaters. net/hitchhikers)
Sea Grant Program (http://www.sgnis.org)
USDA Animal and Plant Health Inspection Service (http://aphis.usda.gov)
The Code of Federal Regulations (49 CFR Section 173.150)
National Coastal Condition Assessment 2015: Quality Assurance Project Plan (EPA-841-
R-14-005)
National Coastal Condition Assessment 2015: Site Evaluation Guidelines (EPA-841-R-14-
006)
National Coastal Condition Assessment 2015: Field Operations Manual (EPA-841-R-14-
007)
National Coastal Condition Assessment 2015: Laboratory Operations Manual (EPA-841-
R-14-008)
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APPENDIX A: EQUIPMENT AND SUPPLIES LISTS
BASE KIT
A base kit will be provided to the field crews for all sampling sites. Some items are sent in the
base kit as extra supplies to be used as needed.
Note: Sodium thiosulfate tablets, filters, 1 Liter HOPE bottles, aluminum foil squares, and
disposable nitrile gloves will be provided in the base kit; you may order more throughout the
field season if needed.
Item Quantity Protocol
B
w
%
m
Aluminum foil squares
Antibiotic salve
Aspirator bulb
Centrifuge tube (50 mL, sterile) - spares
Centrifuge tube stand
Clear tape strips
Electrical tape
FedEx Saturday delivery stickers
Filters (Whatman 47mm GF/F glass fiber 0.7 micron)
Filter flask (500 mL, with side arm) labeled for ENTE filtering
Nutrients filtering chamber
Filtration unit (white base, sterile 100 mL unit, includes pre-loaded
filter for ENTE) — spares
Filtration unit (blue base, 250 mL unit) - spares
Filter funnel adapter (small)
Filter funnel adapter (large)
Forceps (fine-tipped, watchmakers type)
Forceps (sterile, disposable) - spares
Funnel (wide-mouth)
Graduated cylinder (250 mL)
HDPE bottle (2 L, amber)
HDPE bottle (1 L, wide mouth)
Micro centrifuge tube (with sterile glass beads) - spares
Nitrile gloves
Plastic cable tie — spares
Plastic storage tub (for small items in base kit)
Packing tape
Rubber bands (spares)
Rubber stopper (#8 blue, with 15mm hole )
Bag of 50
1 spray tube
1
5
1
3 packs
Iroll
100
Ibox
1
1
5
5
3
3
1
2
1
1
1
12
5
2 boxes
20
1
3 rolls
20
1
Chlorophyll A
Fish Tissue Plug
Fish Tissue Plug
Chlorophyll A
Chlorophyll A
General
Packaging
General
Chlorophyll A
Enterococci
Chlorophyll A and
Dissolved Nutrients
Enterococci
Chlorophyll A
Dissolved Nutrients
Enterococci
Chlorophyll A
Dissolved Nutrients
Benthic
Macroinvertebrates
Enterococci
Sediment Collection
Chlorophyll A
Chlorophyll A
Benthic
Macroinvertebrates
Enterococci
General
Eco Fish Tissue (FTIS)
General
General
Sediment Collection
Chlorophyll A
Dissolved Nutrients
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Item Quantity Protocol
Rubber stopper (#8 white, with 10 mm hole)
Sodium thiosulfate tablets
Scale (in grams)
Secchi disk (20 cm diameter, weighted)
100' of 1/4 inch nylon line for Secchi disk and PAR meter (crews to
mark in 0.5 m intervals)
Self-sealing bags (2 gallon) - spares
Self-sealing bags (sandwich size) - for labels - spares
Sieve box or bucket (stainless steel, 0.5 mm OR 1.0 mm for CA,
OR, & WA)
Spoon, stainless steel (15")
Squirt bottle (for ambient water)
Vacuum pump (hand)
Tyvek tag with grommet - spares
1
Vial of 25
1
1
1
12
100
1
1
1
1
20
Enterococci
Enterococci
Fish Tissue Plug
Water Profile
Water Profile
Eco Fish Tissue
Eco Fish Tissue
Benthic
Macroinvertebrates
Sediment Collection
Sediment Collection
Chlorophyll A
Enterococci
Dissolved Nutrients
Eco Fish Tissue (FTIS)
ADDITIONAL BASE KIT ITEMS - GREAT LAKES CREWS
Item Quantity Protocol
GL BASE KIT
HDPE bottle (1 L, white, narrow mouth)
LugoPs
Pipet (10 mL)
Pipet bulb
Seaviewer underwater camera system (with DVR, GPS, cables,
case)
I/ site
1
2
1
1
Phytoplankton
Phytoplankton
Phytoplankton
Phytoplankton
Underwater video
SITE KIT
A site kit will be provided to the field crews for each sampling site. Please submit an
electronic request form well in advance of field sampling. Kits must be requested at least
three weeks before sampling is to take place. Each site kit will also include necessary coolers
and shipping supplies for all samples collected. Prior to sampling, inspect each site kit to
ensure all supplies are included. Some items may not be used at all sites and should be held
until the end of the field season and shipped back.
The Field Crew Leader MUST provide a general schedule in order to receive the site kits.
These kits include:
Item Quantity Protocol
g
S
HH
C/3
Bubble bag (microcentrifuge tubes in this)
Bucket, screw top (0.6 gallon)
Centrifuge tube (50 mL, sterile)
Cooler(s)
FedEx air bills (pre-addressed) plus handle tags, zip ties, etc.
1
1
1
1
Enterococci (Shipping)
Sediment Toxicity
Chlorophyll A
Enterococci
Shipping
Shipping
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Item Quantity Protocol
Filtration unit (white base, sterile 100 mL units, includes pre-loaded
filter for ENTE)
Filtration unit (blue base, 250 mL unit)
Fish Tissue Plug Kit
Biopsy punch (sterile, disposable)
Disposable forceps (sterile)
Glass scintillation vial (20 mL)
Scalpel (sterile, disposable)
Bubble bag for vial
Outer bag for vial
Forceps (sterile, disposable)
Glass jar (120 mL, amber)
Glass jar (60 mL, amber)
HDPE bottle (250 mL, white)
HDPE bottle (250 mL, amber)
HDPE bottle (250 mL, white, sterile)
HDPE bottle (500 mL, white, wide mouth)
HDPE bottle (1 L, white, wide mouth)
Plastic bag (large, composite)
Plastic bag (sandwich size) for CHLA tube
Plastic bag (quart)
Plastic cable ties
Self-sealing bags (2 gallon)
Self-sealing bags (sandwich size) - for eco fish labels
Sterile phosphate buffer solution (PBS)
Tyvek tags with grommets
2
1
1
1
1
1
1
1
1
2
1
1
1
1
1
2
1
1
1
2
1
2
2
1 jar
10
Enterococci
Chlorophyll A
Dissolved Nutrients
Fish Tissue Plugs
Enterococci
Chlorophyll A
Sediment
Organics/Metals
Sediment TOC
Dissolved Nutrients
Water Chemistry
Enterococci
Algal Toxin
Microcystin
Benthic
Macroinvertebrates
Eco Fish Tissue
Chlorophyll A
Sediment Grain Size
Eco Fish Tissue
Eco Fish Tissue
Eco Fish Tissue
Enterococci
Eco Fish Tissue
FORM & LABEL PACKET
A form 6t label packet will be provided to the field crews for all sampling sites (separately
from site kits). Please submit an electronic request form well in advance of field sampling. A
packet must be requested at least three weeks before sampling is to take place. Prior to
sampling, inspect each packet to ensure all forms and labels are included. Depending on what
your crew is doing (type of sites and whether you are using e-forms), you may request:
• Great Lakes field forms, tracking forms & labels
• Marine field forms, tracking forms & labels
• Tracking forms & labels only (e-forms users)
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HUMAN HEALTH FISH TISSUE SAMPLING SITE KIT
A human health fish tissue kit will be provided to the field crews for selected sampling sites
(separately from site kits). Please submit an electronic request form well in advance of field
sampling. Kits must be requested at least three weeks before sampling is to take place. Prior
to sampling, inspect each human health fish tissue kit to ensure all supplies are included.
These kits include:
Item Quantity Protocol
HH FISH TISSUE KIT
Aluminum foil (solvent rinsed & baked)
Cooler (blue)
Dry ice (Class 9) shipping label
FedEx airbill (pre-addressed)
Nitrile gloves
Plastic bags (large, composite)
Plastic cable ties
Polyethylene tubing (heavy-duty, food grade)
Tyvek tags with grommets
5
1
1
1
5 pairs
1
12
Iroll
1
Packaging
Storage & Shipping
Shipping
Shipping
Packaging
Packaging
Packaging
Packaging
Packaging
CREW SUPPLIED EQUIPMENT
Item Quantity Protocol
GENERAL
Active/passive fish sampling device (e.g. trawl, seine, hook & line, etc.)
Alconox
Barometer (for calibration)
Batteries (AA)
Bleach (1-10% solution)
Borax
Buckets (large)
Calibration cups & standards
Cell phone, 2-way radios, walkie talkies
Clipboard(s)
De-ionized water (lab certified preferred, not required)
Digital camera (with extra memory card & batteries)
Dip net
Dry ice
Fine-tipped, indelible markers
Formalin (100% buffered) with stain
Fuses (10 amp)
GPS unit (with manual & reference card, extra battery pack);
Graduated cylinder (for measuring formalin)
Knife
Livewell/buckets with aerator
Maps & access instructions
Measuring board (mm scale)
1-2
1
-50 Ibs/site
1
1
Fish Collection
Sediment Collection
Water Profile
GPS, Water Profile,
Underwater Video
Decontamination
Sediment Collection
Sediment Collection
Profile
General
General
Water Profile
General
Fish Collection
Shipping
General
Sediment Collection
Underwater Video
General
Benthos
General
Fish Collection
General
Fish Collection
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Item Quantity Protocol
I
m
Multi-parameter probe water quality meter (with pH, DO, temperature,
and conductivity/ salinity probes - e.g. Hydrolab, YSI, etc.)
NCCA 2015 Fact Sheets (available on NARS SharePoint)
PAR meter (with LI-190 Quantum Sensor and LI-192 Underwater
Quantum Sensor & cables, independent datalogger)
Pencils (#2)
Plastic tub or bucket
QCS - quality check solution
Rose Bengal stain
Ruler (in cm)
Sampling permits /permission letters
Scissors
Scrub brush
Sieve box/frame (if necessary)
Spare parts
Stainless steel or Teflon spoons (large & small), spatulas, & scoops
Stainless steel mixing pot or bowl with lid
Stop watch
Thermometer
Water sampling device (e.g., Niskin) or pump system
Weights & pads for grabs
Wet ice
Wooden bat
Young-modified Van Veen grab sampler (0.04 m2) OR standard OR
Petite Ponar sampler with grab stand, plastic tub, drop line, pinch pin
Anchor (with 75 m line or sufficient to anchor in 50 m depth)
Boat horn
Bow/ Stern lights
Emergency tool kit
Extra boat plug
Fire extinguisher
First aid kit
Float (to attach to anchor)
Gas Can
Hand bilge pump
Motor
PFDs (I/person)
1
10
1
5
1
If needed
1 bottle
1
1
1
1
Various
1
1
1
1
-50
Ibs/site,
additional
for shipping
1
1
Water Profile
General, Outreach
Water Profile
General
Sediment Collection
Water Profile
Sediment Collection
General
General
General
Sediment Collection
Sediment Collection
Multi-probe
Mixing and
dispensing sediment
Sediment Collection
Underwater Video
Water Profile
Chlorophyll A
Dissolved Nutrients
Phytoplankton
Water Chemistry
Microcystin
Sediment Collection
Shipping
Fish Collection
Sediment Collection
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Item Quantity Protocol
Fingers
Sonar unit
Spare prop
Spare prop shear pin
Type IV PFD (throwable life saving device)
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APPENDIX B: FIELD FORMS, LABELS & TRACKING FORMS
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SITE VERIFICATION (FRONT)
g NCCA2015SITEVERIFI
Site ID: Visit: O 1 O z
Reviewed by Initial):
L.AI ION (hront) •
Date: / /
Site Name: State of Site Location: Field Crew:
DID YOU SAMPLE THIS SITE?
O YES If YES, check one below:
SAMP LEA BLE (Choose method used|
O Marine
O Great Lakes
ARRIVAL TIME: | : 4
HABITAT TYPE: O Tidal River OOpen Water O MarsNv^and OEmbayment O Inter-Tidal O Rivermouth
O Other, explain:
BOTTOM TYPE: O Coral Reef O Oyster Bed OfcTass Bed
O Other, explain:
O Sand O Rocky/Shell OHardpan O Mud
Debris Present?: ! If Yes, TYPE:
OYES O MO I O Glass O Plastic OWood QCans O Other, explain:
SAV Present?: O Yes Ol*t ABUNDANCE:
Macroalgae Present? :O Yes ONo ABUNDANCE:
(Sparse, dense, etc)
(Sparse, dense, etc)
GENERAL COMMENTS
DIRECTIONS TO SITE
H 5924322647 03/31/2015 NCCA2015 Verification D1 H
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SITE VERIFICATION (BACK)
r
Site ID:
NCCA 2015 SITE VERIFICATION (Back) R"^-1" ^
Date: / /
SKETCH MAP
Arrow Indicates North; Label Sketch:
NOTE: If an outline map is attached
with the outline map on it.
L=Launch; X=X-site; F=Fishing Area; S=Sediment Area; Y=Y Location
here, use a continuous strip of clear tape across the top edge. You can also attach a separate sheet
^f)
s
X"Vv
PERSONNEL
Crew Leader:
Fish Taxonomist:
Name:
Name:
^ 0300322640
Name:
Name:
Name:
Name:
03« 1 /20 1 5 NCC A 20 1 5 Verificali on p. ,., ^
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FIELD MEASUREMENT (FRONT)
Site ID:
NCCA 2015 FIELD MEASUREMENT (Front)
Date: / /
Reviewed by (Initial):
CALIBRATION INFORMATION
Instrument manufacturer and model:
Instrument ID number:
Operator:
O Mode exempt from field calibration protocols (e.g., Sea-Bird)
fEMFERATURE
Thermometer Reading (°C) Sensor Reading (°C) Comments
Barometric
Pressure 4mm Hg) Callbratian Value
Displayed Value
DO
Qmg/L
,0%
Qmg/L
PH
Cal. STD 1 Description Cal. STD 1 Value Cal. STD 2 Description
Cal. STD Z Value
Cal. STD 1 Description
Cal. STD 1 Value Cal. STD 2 Descr'ntlon
Cal. STD Z Value
CONDUCTIVITY
QUALITY CONTROL CHECK (Perform at least once per week)
O No QC Check perfomied at this visit
O Internal meter checks performed and passed
Date Prepared: 1 1
.ne:
(rh:rim)
ParametBr TEMP. (DC) COND (MS) pH
Expected:
Measured: I
Comments:
POST-MEASUREMENT CALIBRATION CHi/CK
pH COND(uS)
Expected:
SECCHI DEPTH (m| XX.X:
|iih nM:i|
DISAPPEARS: REAPPEARS: CLEAR TO BOTTOM? Yes Q No Q
i If yes, record station depth as both disappearance and reappearance depth for
Riding 1: Readjno ,
Reading 2:
Reading 3:
Secchi Comments:
Flag
Comments
Flag and Comment here for Hydrographic Profile
3536545517 Flag codes: K = Sample not collected; U = Suspect sample; F1, F2, etc. = flag assigned by field crew.
03/31/2015 NCCA2015 Field Measurement D3
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FIELD MEASUREMENT (BACK)
• NCCA 2015 FIELD MEASUREMENT (Back)
Site ID: Date: /
Hydrograptiic Prof
Interva s (m : 0.1 m b
STATION DEPTH (m
XX. X
Nes
•
D
O
W
N
C
A
S
T
r Bottom
U
P
C
A
S
T
Rtvi«wtd bvllnlttal): ^^_
/
•
le
el ow surface, 0.5 below the surface, every 1 meter from depths of 1 .0 to 10m, and every 5 meters thereafter if the site is greater
than 10m. Take thelastset of me asureme nt s at 0.5m from the bottom.
eompi.i..ith.rSALQreci«C: Submitted data via eFlle Q
S AL (%J SP CON D (uSJc m) U GHT| AMB) LIGHT(UW)
DEPTH(m) TEKP. (-C) pM DO(iriiJL) (Marine) (Great Lanes) uElmzrs uEfflZfs
XX.X XX.X XX.XX XK.K XX.X XXX.X XXX.X XXX.X FLAG
0.1
0.5
* ^^v
1
V
Flag codes: K = Sample not collected; U = Suspect sample; F1, F2, etc. = flag assigned by field crew.
Record any Profile flag and comments on front side of this field measurement form.
03/31/2015 NCCA 2015 Field Measurement
D4 —
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SAMPLE COLLECTION (FRONT)
Site ID:
NCCA 2015 SAMPLE COLLECTION - (Front) flnit!a™d "* •
Date: / /
WATER CHEMISTRY , CHLOROPHYLL and NUTRIENT COLLECTION (0.5m) (CHEM, CHLA, NUTS)
Water Cheml stry ID:
(Non-Filtered)
» * t i i i *
Chlorophyll-a ID:
i i i i i i i
Nutrients ID:
(Filtered)
t i i i i i i
Chilled
O
Frozen
O
Chilled
O
Comments: No Sample Collected Q
'" (ml)"™11 Comments: No SamPle Collected Q
No Sample CollectedO
Comments:
MICROCYSTIN (MICX) No Col,ected Q
(Target Volume = 500 ml)
Sample ID: Frozen Comments:
1 1 1 1 1 1 1
O
ALGAL TOXIN (ALGX) No Co|]ected Q
(Target Volume = 500 ml)
Sample ID:
Frozen
O
Comments:
ENTEROCOCCI (ENTE) Nc ^P'6 Co"ectBd O
(Target Volume = 250 mL) Blank collected O
Sample ID:
1 I 1 1 1 1 1
Time Depth Fitt. n^tart Volume Filtered Buffer Rinse Filt. End Time
Collected Collected T:.ne (Target = 50 ml) (Target = 2 rinses of 1Qml_) Time Frozen
(hhmm
(m) (hhmm) Filt. 1 Fift. 2 Filt. 1 Filt. 1 (hhmm) (hhmm)
Comments:
GREAT LAKES ONLY
PHYTOPLANKTON (1 L narrow mouth HOPE bottle) (PHYT) Nc Sample Collected Q
Sample ID:
i * i i i _j f
Preserved
O
Depth Time Time
Collected Collected Preserved
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SAMPLE COLLECTION (BACK)
Site ID:
NCCA 2015 SAMPLE COLLECTION - (Back)
Date: /
Reviewed tjy
(initial): —
BENTHIC INFAUNA COLLECTION (1L wide mouth HOPE bottle) (BENT)
No Sample Collected
BENTHIC COLLECTION LOCATION:
O Within 37m from X-site O Between 37-100m from X-site O Between 100-500m from X-site
O Van Veen
GRAB AREA (m2):
GRAB TYPE: ~ I™' "' O Other
O Standard Ponar
SIEVE SIZE: O 0-5 mm O 1-0 mm NUMBER OF GRABS: O 1 O2 NOTE: 2 Grabs are required for samplers less than 0.03 m*
Sample ID:
Depth (cm)
(Should be >7 cm)
O
SEDIMENT CHARACTERISTICS (BenthiC Grab)
COLOR: O Black O Brown O Light Brown O Dark Brown O Gray O Other
SUBSTRATE: O Sand OMuck O Gravel O Cobble O Shellhash O Other
SMELL: O Fishy O Chemical OSuIPnur O None O Other
SURFACE: O Film O Floe O Nothing Noted O Other
VISIBLE FAUNA: O Yes O No TYPE:
VISIBLE FLORA: O Yes O No TYPE:
SEDIMENT SAMPLE COLLECTION
SEDIMENT COLLECTION LOCATION;
O Within 37m from X-site O Between 37-100m from X-site fj Between 100-500m from X-site
SEDIMENT TOXICITY (0.6 gal Screw Top Bu.cf«»^r-DX) (Target = 900mL)
No Sample Collected
Sample ID:
O
SEDIMENT ORGANIC5/METALS (Gla?s jar 120 ml) (SEDO) (Target = 100 mL)
Sample ID:
Fro zan L irr. nents:
No Sample Collected Q
O
SEDIMENT TOC (Glass Jar 60ml) (SEDC) (Target = 50 mL)
No Sample Collected
Sample ID:
Frozen Comments:
O
SEDIMENT GRAIN SIZE(1 Qt. Ziplock)(SEDG) (Targets 100 mL)
No Sample Collected
Sample ID:
O
Use comment section to explain: No measurement, suspect measurement or observation made.
9 2*33462138 03/31/2015 NCCA 2015 Sample Collection
D6
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Eco FISH COLLECTION (FRONT)
NCCA 2015 ECO FISH COLLECTION (Front)
Site ID: Date: /
Trawl
Zone(s): O Within 500m from X-site O Between 500-1000m from X-site
Start Time: : End Time: : Fished as:
1 ' ' ' ' ' ' ' ' ' ' ' O Bottom Trawl
Gear Details: Opening size (m): h» Mesh size (cm):
v ' DV ' O Mid-Water Trawl
O Attempted and caught target fish O Attempted and failed to catch target fish O Attempted and failed to catch any fish
Seine
Zone(s): O Within 500m from X-site O Between 500-1000m from X-site
Start Time: : End Time: :
Gear Details: Length (m): Height (m): Mesh size (cm):
O Attempted and caught target fish O Attempted and failed to catch target fish C Attempted and failed to catch any fish
Gill Net
Zone(s): O Within 500m from X-site O Between 500-1000m from X-i:™
,C*2
Start Time: : End Time: :
i i i i i i i i i i i i
GearDefaiis: Length (m): Height (m): Mesh size (cm):
noted and i^il
O Attempted and caught target fish O Attempted and lulled to catch target fish O Attempted and failed to catch any fish
Comments:
Hook and Line
Zone(s): O Within 500m from X-site C Between 500-1000m from X-site
Start Time: : E'd"irne: :
[X
Gear Details:
O Attempted and caught target fish O Attempted and failed to catch target fish O Attempted and failed to catch any fish
Other:
Zone(s): O Within 500m from X-site O Between 500-1 OOOrn from X-site
Start Time: : End Time: :
Gear Details:
O Attempted and caught target fish O Attempted and failed to catch target fish O Attempted and failed to catch any fish
Comments:
4640348526 04AJ2/2015 NCCA2015 ECO Fish CoHeclion (Front) Q^
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 113
Eco FISH COLLECTION (BACK)
• NCCA 2015 ECO FISH COLLECTIO
Site ID: Date: /
Reviewed by (inltia : ^_
N (Back) * •
/
FISH TISSUE SAMPLE (FTIS) MO SAMPLE COLLECTED O
FISH ALL WITHIN 75% OF LARGEST SPECIMEN Q
FISH ARE ALL THE SAME SPECIES Q
FISH COMPOSITE TOTAL MASS IS AT LEAST 300 GRAMS Q
Sample ID Total Length (mm Total Length (mm Total Length (mm)
Frozen: Q 1
Scientific Name (Genus Species) 2
3
4
The eco fish composite must consist of at least 5 fish of 5
qr]e>qi |qtp QJJP tn prnyjrlp H fntal Weight nf 3flfl rjr^m« "f
whole-body tissue, 6
7
8
9
10
11 21
12 22
13 23
14 24
15 25
15 | 26
17 27
18 28
19 29
20 30
Comments:
FISH TISSUE PLUG SAMPLES (FPLG) NO SAMPLE COLLECTED Q
FISH PLUG SAMPLE COLLECTED FROM SAME SPF.CIMEN AS ECO FISH SAMPLE?
O* ON If No, answer the following:
O FISH ALL WITHIN 75% OF . AK^EST SPECIMEN
O FISH ARE ALL THE SPL'.F. k SECIES
COLLECTION MET.OL
O TRAWL ^ HOOK & LINE Q SEINE Q GILL NET O PURCHASED DOCKSIDE
O OTHER EXPLAIN:
Sample ID Scientific 'Name (Genus Spec es)
i i i i i i i
Length(mm) Weight(g)
Comments:
1 7610492266 OM1/2015 NCCA 201 5 ECO Fish Collection (Back) Qg I
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 114
SITE ASSESSMENT (FRONT)
Site ID:
NCCA 2015 SITE ASSESSMENT (Front)
Date: / /
SHORELINE ACTIVITIES AND DISTURBANCES
(Intensity: Blank=Nol observed, L=Low. NT=Moderate, H=Heaw)
BLANK FIELD INDICATES ABSENCE: f")
Residential
Recreational
Agricultural
Industrial
Management
O O O Residences
O O O Maintained Lawns
0 © ©Construction
O 0 O Pipes. Drains
O 0 O Dumping
O 0 O R°ads
O0© Bridges/Causeway
O 0 O Sewage Treatment
0 0 O Hiking Trails
0 0 O Parts, Campgrounds
O 0 © Primitive Parks, Camping
©0© Trash/Litter
©0© Surface Films
0 0 O Dim«s
O 0 © Beach
O0© Forested
000 Cropland
000 Pasture
00© Livestock USB
0 0 © Orchards
00© Poultry
O 0 0 Irrigation Equip.
0 0 © water Withdrawn
0 0 0 Industrial Plants
000 Mines/Quarries
O © O OilrtSas Wfelk
© 0 © Power Plants
O 0 0 Logging
O 0 O Evidence of File
© © © Odors
© © ©Commercial
000 Chemical Treatment
0 0 O Angling Pressure
© © 0 Dredging
© © © Channefization
© © © Water Level Fluctuations
0 0 0 Shoreline Hardening
© © © Dredge Material
SITE CHARACTERISTICS (200m radius)
WATERBODY CHARACTER
PRISTINE: OS O4 O3 O2 O1 Highly DisUirbad
APPEALING: ©5 ©4 O3 ©2 ©1 Unappealing
ASSESSMENT OF VISIBLE TRASH IN WATER (AQUATIC TRASH) B: AN.' FIELD INDICATES ABSENCE Q
Items Observed
Aluminum Cans
Plastic Bottles
Other Plastic Items
Tires
Fishing Gear
Other
Qty. Observed
<5 5-20 >20
O
O
O
O
O
O
O
O
O
O
O
O
O
C!
List:
DOMINANT LAND USE
Dominant Land Use Around '.<' Ol-orest ©Agriculture ©Range ©Urban © Suburbann~own
If Forest, Dominant Age Class O 0 - 25 yrs © 26 - 75 yrs. Q>75 yrs.
WEATHER
GENERAL ASSESSMENT [Biotic integrity, Vegetation diversity, Local anecdotal information)
6659424523
04/07/2015 NCCA 2015 Site Assessment
D9
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National Coastal Condition Assessment 2015 Field Operations Manual
Version 1.0 May 2015 Page 115
SITE ASSESSMENT (BACK)
, NCCA 2015 SITE ASSESSMENT (Back)
Site ID: Date: / /
GENERAL ASSESSMENT (continued)
___
1641424529 n.n
03/31/2015 NCCA2015SiteAssessment D10
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National Coastal Condition Assessment 2015 Field Operations Manual
Version 1.0 May 2015 Page 116
HUMAN HEALTH FISH COLLECTION (FRONT)
r NCCA 2015 HUMAN HEALTH FISH COLLECTION (Front) RMmat,t,MW:
Great Lakes Only
Site ID: Date: / /
Trawl
Zone(s): O Within 500m from X-site O Between 500-1 OOOm from X-site O Between 1000-1500m from X-site
Start Time: : End Time: : Fished as:
O Bottom Trawl
Gear Details: Opening size(m): hi/ Mesh size (cm): m,^**, 4. -r
1 ' DJ * ' O Mid-Water Trawl
O Attempted and caught target fish O Attempted and failed to catch target fish O Attempted and failed to catch any fish
Seine
Zone(s): O Within 500m from X-site O Between 500-1 OOOrn from X-site O Between 1000-1500m from X-site
Start Time: : End Time: :
GearDetaiis: Length (m): Height (m): Mesh size (cm):
,
O Attempted and caught target fish O Attempted and failed to catch target fish C A'.tempted and failed to catch any fish
Comments:
Gill Net
Zone(s): O Within 500m from X-site O Between 500-1 OOOrn from X Vf*> O Between 1000-1500m from X-site
V?
Start Time: : End Time:
GearDetaiis: Length (m): Height (m): Mesh size (cm):
O Attempted and caught target fish O Attempted and lulled to catch target fish O Attempted and failed to catch any fish
nments:
Hook and Line
Zone(s): O Within 500m from X-site C Between 500-1 OOOm from X-site O Between 1000-1500m from X-site
Start Time: : E'd"'rne: :
Gear Details:
O Attempted and caught target fish O Attempted and failed to catch target fish O Attempted and failed to catch any fish
Other:
Zone(s): O Within 500m from X-site O Between 500-1 OOOrn from X-site O Between 1000-1500m from X-site
Start Time: : End Time: :
GearDetaiis:
O Attempted and caught target fish O Attempted and failed to catch target fish O Attempted and failed to catch any fish
133727 6098 04/02/2015 NCCA 2015 Human Health Fish Collection (Front) Q^
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 117
HUMAN HEALTH FISH COLLECTION (BACK)
Site ID:
NCCA 2015 HUMAN HEALTH FISH COLLECTION (Back) «*«
Great Lakes Only
Date: / /
HUMAN HEALTH FISH TISSUE SAMPLE (HTIS)
NO SAMPLE COLLECTED Q
FISH ALL WITHIN 75% OF LARGEST SPECIMEN
FISH ARE ALL THE SAME SPECIES
Sample ID
Total Length (mm) Total Length (mm) Total Length (mm)
Frozen: Q
.01
.11
.21
Scientific Name (Genus Species)
.02
.12
.22
.03
.13
.23
.04
.14
The human health fish composite ideally consists of 5 fish of
adequate size to provide a total weight of 500 grams (equivalent to
about 13 02. J of fillet tissue. Fewer or more fish can be collected to
meet the fillet tissue weight requirement.
.05
.15
_|
.06
.15
.07
.17
.08
.09
.10
.18
.19
.20
.24
.25
.26
.27
.28
.29
.30
o
2835224626
03/31/2015 NCCA2015 Human Health Fish Collection (Bach)
D12
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Version 1.0 May 2015
Field Operations Manual
Page 118
SAMPLE I
.ABELS (MARINE)
WATER CHEMISTRY (CHEM) WATER COLUMN CHLOROPHYLL (CHLA)
Site ID: Site ID:
Date: / /?01 VhitfcOl O? D.-itr?: / /701_ Visit »:O1 O?
T4 999000 fa Volume Filtered: ml
999001
NUTRIENTS (NUTS) MICROCYSTIN (MICX)
Site ID: Site ID:
Date: / /201 Visit ft O1O2 Date: / /201 Visit #:OlO2
T1 9990D2 T, Salinity: (K.)
999003
BENTHIC INFAUNA (BENT) SEDII^tN i TOC (SEDC)
Site ID: S ;e P.
Date: / /201 Visit «: Ol O2 Date: i /201_ Visit »: Ol O2
Salinity: (#.) Jar 1 of „ 999005
999004
SEDIMENT GRAIN SIZE (SEDG) SEDIMENT ORGANICS/METAL (SEDO)
Site ID: Site ID:
Date: / /201 Visit »: Ol O2 Date: / /201_ Visit *: Ol O2
T2 999006 T3 999007
0°
SEDIMENT TOXICITY (SEDX) ALGAL TOXIN (ALGX)
Site ID: . n, ' Site ID:
Date: / /201 Visi' h- Cl O2 Dale: / /201 Visit »: Ol O2
_ 999008 Salinrtv: (M
999009
FISH TISSUE PLUG (FPLG) ECO FISH TISSUE - OUTER BAG
Site ID: T6 Site ID:
Date: / /201 Visit ft Ol O? Date: / /201 Visit #: Ol O?
T;j 999010 Genjs Species:
Lenjitn fimm] Min.: Max.:
999012
ECO FISH TISSUE - INNER BAG OF ECO FISH TISSUE - INNER BAG OF
Site ID: Site ID:
Date: / /201_ Visit »: Ol O2 Date: / /201_ Visit #: Ol O2
Genus Species: Genus Species:
Lensth (mm) Min.: Max.: Length (mm) Min.: Max.:
999012 999012
ENTEROCOCCI (ENTE) - BAG ^ E
Site ID: ^ >
_j
E
T-H
^H
O
. . 01
*J 01
~ 0!
"5
>
Benthic Infauna (BENT) ECO Fish Tissue (FTIS)
Site ID:
Site ID:
Date: / /201 Visit #: O1 O2
Date: / /201 Visit # O1 O2
Length (mm) Mm.: Max.:
Sampler Tvpe: Baa of
nnllprtnr/i^ SAMPLE ID:
Jar of
SAMPLE ID:
ECO FISH TISSUE (FTIS) - EXTRA
Site ID-
Date: / /201 Visit #: 01 02
Lenath: mm Baa of
SAMPLE ID
BENTHIC INFAUNA (BENT) - EXTRA JAR
Date: / /201 Visit #: O1 O2
Jar of
SAMPLE ID:
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 119
SAMPLE LABELS (GREAT LAKES)
WATER CHEMISTRY (CHEM) WATER COLUMN CHLOROPHYLL (CHLA)
Site ID: Site ID:
Date: / /201_ Visit tf: Ol O2 Date: / /201 Visit »: Ol O2
999200 T« Volume Filtered: ml
999201
NUTRIENTS (NUTS) MICROCYSTIN (MICX)
Site ID: Site ID:
Date: / /201_ Visit tf: Ol O2 Date: / /201_ Visit »: Ol O2
T1 999202 T3 999203
V
BENTHICINFAUNA(BENT) SEDIu^NrTOC (SEDC)
Site ID: Site r.
Date: / /201 Visit 8: Ol O2 Date: / /201 Visit 8: Ol O2
T4 999204 TD 999205
SEDIMENT GRAIN SIZE (SEDG) PEDIMENT ORGANICS/METAL (SEDO)
Site ID: Site ID:
Date- / /201 Visit tf- Ol O2 Date- / /201 Visit #• Ol O2
T2 999206 T3 999207
0°
SEDIMENT TOXICITY (SEDX) ALGALTOXIN (ALGX)
Site ID: Site ID:
Date: / /201 Visi'tt. Oi O2 Date: / /201 Visit #: Ol O2
T2 999208 T3
999209
FISH TISSUE PLUG (FPLG) ECO FISH TISSUE - OUTER BAG
Site ID: T5 Site ID:
Date: / /201 Visit »: Ol O2 Date: / /201 Visit »: Ol O?
( f^, 999210 Genus Species:
Length (mm) Min.: Max.:
999212
ECO FISH TISSUE - INNER BAG OF ECO FISH TISSUE - INNER BAG OF
Site ID: Site ID:
Date: / /201 Visit tf: Ol O2 Date: / /201 Visit ff: Ol O2
Genus Species: Genus Species:
Length (mm) Min,: Max.: Length (mm) Min.: Max.:
999212 999212
ENTEROCOCCI (ENTE) - BAG ^
Site ID: -H _
Date: / /201 Visit #:O1 O2 -3
±l ~ 3
Vol. Fill: 1 ml 2 ml "- iz 01
999011 1
PHYTOPLANKTON (PHYT)
Site ID:
Date: / /201 Visit S:O1 O2
T2 999013
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: / /201_ Visit S:Ol O2
Genus Species:
Length: mm
999014.01
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: / /201 Visit S: Ol O2
Gen us Species:
Length: mm
999014.02
HH FISH TISSUE WHOLE (HTIS)
Sitp ID:
Ddle: / /201 Vi-.il B:O1 O2
Gen us Species:
Length: mm
999014.03
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: / /201_ Visit ft. Ol O2
99901404
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: / /'201 Visit «:OlO2
Gen us Species:
Length: mm
999014.05
HH FISH TISSUE WHOLE (HTIS) - BAG
Site ID:
Date: / /201_ Visit S:O1 O2
Genus Species:
Length range: mm to mm
999014
_i _i
E E
_a*
c
(N ^H [B rH
•• «H m iH
i-O >
Benthic Infauna (BENT) ECO Fish Tissue (FTIS)
Site ID:
Site ID:
ECO FISH TISSUE (FTIS) - EXTRA
Site ID:
Date: / /2Q1 Visit #. O1 O2 Datp
Date / /201 Visit if: O1 O2
Length (mm) Min : Max
Sampler Type Bag ^
Ler
Colleotor(s) SAMPLE ID.
BEN
Jar of
SAMPLE ID: Hate
: / /201 Visit #: O1 O2
ath: mm Baa of
SAMPLE ID
THIC INFAUNA (BENT) - EXTFJA JAR
Site ID:
/ /201 Visit #: O1 O2
Jar of
SAMPLE ID:
-------
National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 120
SITE AND SAMPLE STATUS/WATER CHEMISTRY LAB TRACKING
£ NCCA 2015 SITE AND SAMPLE STATUS/WATER CHEMISTRY LAB TRACKING £
Site ID: Visit #:Q1 O2 Date Collected: / /
State of Site Location: Crew:
Sender: Sender Phone: - -
Shipped by: O FedEx O UPS O Hand Delivery O Other:
Airbill/Tracking Number: Date Sent: / /
Site Status - Is Site Sampleable?
O YES If Yes, check one below
SAMPLEABLE (Choose method used)
O Marine
O Great Lakes
Sample Status - Water Chemistry Lab Samples
Sent to State
Sample Sent to (Note in Not
Sample ID Type WRS Comments) Collected
CHEM O O O
CHLA O O O
t NUTS O O O
Sample Status - Batch Samples
Sample Not
Type Collected Collected Comments
ALGX O O
BENT O O
ENTE O O
FPLG O O
FTIS O O
wiicx O O
SEDC O O
SEDG O O
O NO If No, check one below
NON-SAMPLEABLE-PERMANENT-Replace Site
O M ap Error
O Site too shallow for navigation/sampling
Q Unsafe
O No Access
NON-SAMPL E/ e.. t-TEMPORARY-Re schedule
O Temporarily h ic*.ii-ii/a.i
Email:monaco,pr,il
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 121
TRACKING: BATCH SAMPLES - OVERNIGHT (CHILLED)
Y NCCA
State of Site Location:
Sender:
Shipped by: O FedEx O UPS
Site ID:
Sample
Sample ID Type
SEDG
SEDX
PHYT*
2015 TRACKING: BATCH SAMPLES - OVERNIGHT (CHILLED) ^
Crew: Date Sent: j }
Sender Phone: - —
O Hand Delivery Airbill/Tracking Number:
Visit: Q1 Q 2 Date Collected: / /
Containers Comments
* Great Lakes Only
O GLEC - Traverse City
O STATE LAB (provide details below)
State Lab Name:
State Lab address:
City:
V
jt /*?: Zip Code:
Save completed form as:
Tracking Related Inquiries:
NCCA15_T2_Tratking_ShelD_V» Marlys Cappaert
Phone: 541 -754-4467
Email to :
sampletraikiriE@epa.gov Michelle Cover
Phone: 541-754-4793
Or fax to: 541-754-4637
^. 0046154808 A
^ 03/31 CO 15 NCCA 2015 Tracking- Batch Overnight Chilled J2
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Version 1.0 May 2015
Field Operations Manual
Page 122
Ti
IACKING: BATCH SAMPLES - OVERNIGHT (DRY ICE)
Y NCCA 2015 TRACKING: BATCH SAMPLES - OVERNIGHT (DRY ICE) ^
State of Site Location: Crew: Date Sent: / /
Sender:
Sender Phone: - —
Shipped by: O FedEx O UPS O Hand Delivery Airbill/Tracking Number:
Sits ID:
Sample ID
Sample #
Type Cont*
ALGX
ENTE
MICX
FPLG
SEDC
SEDO
Visit; Q1 Q 2 Date Collected: f f
of
liners Comments
O GLEC - Traverse City
rr
O STATE LAB (provide details below)
State Lab Name:
State Lab address
City:
.
-_ X S,:.ie: Zip Code:
Save completed form as:
NCCA15_T3_Tracti ing_S[tel D_V»
Email to :
sa m p let rack imj@e pa .gov
Or fax to: 541-754-4637
^ 2882141152
Tracking Related Inquiries:
Marlys Cappaert
Phone: 541 -754-4467
Michelle Cover
Phone: 541-754-4793
03/31/2015 NCCA 201 5 Tracking- Batch Overnight T3 ^
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 123
TRACKING: BATCH SAMPLES - GROUND (No ICE)
NCCA 2015 TRACKING: BATCH SAMPLES - GROUND (NO ICE)
State of Site Location:
Sender:
Sender Phone:
Shipped by: O FedEx O UPS O Hand Delivery AirbilUTracking Number:
Site ID:
Visit: O 1 O2 Date Collected:
Sample ID
Sample Type
BENT
O GLEC - Traverse City
O STATE LAB (provide details below)
State Lab Name:
•$•
State Lab address:
je
City:
Save completed form as:
Tracking Related Inquiries:
NCCA15_T4_Tracking_SitelD_V8
Email to :
sa m pletracki ng@epa.gov
Or fax to: 541-754-4637
1747216659
Marlys Cappaert
Phone: 541 -754-4467
Michelle Gover
Phone: 541-754-4793
03/31/2015 NCCA 201S Tracking - Batch Ground Shipping
T4
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 124
TRACKING: Eco FISH TISSUE - OVERNIGHT (DRY ICE)
Y NCCA 2015 TRACKING: ECO FISH TISSUE - OVERNIGHT (DRY ICE) ^
State of Site Location: Crew: Date Sent: / /
Sender:
Sender Phone: _ _
Shipped by: O FedEx O UPS O Hand Delivery Airbill/Tracking Number:
Sits ID: Visit: Q 1
FISH ALL WITHIN 75%
Sample ID
Frozen: O
Scientific Name (Genus Species) '
:
i
The eca fish composite must consist of at least 5 fish of j
adequate size to provide a total weight of 300 grams of
whole- body tissue I
•
t
i
i
O 2 Date Collected: / /
OF LARGEST SPECIMEN Q FISH ARE ALL THE SAME SPECIES Q
FISH COMPOSITE TOTAL MASS IS AT LEAST 300 GRAMSQ
Total Length (mm Total Length (mm Total Length (mm)
11 21
! 12 22
I 13 23
I 14 24
S 1C 25
t ,6 26
f (?) 17 27
t 18 28
i 19 29
0 20 30
Fish crew, if different than site crew
Crew Leader:
Fish Taxonornist:
Name:
Name:
Name:
Name:
Name:
Name:
OGLEQ - Traverse City State Lab Name:
O STATE LAB (provide details)
State Lab address:
Save completed form as:
City: State: Zip Code:
Tracking Related Inquiries:
NCCAI5 T5 Tra<:king_SiteID Vfl MarlYs Cappaert
Phone:541-754-4467
E ma i I to : sa m p let ra ck inge pa .gov 0 r ; a x to : 54 1- 7 54-4637
Michelle Cover
Phone:541-754-4793
^. 4955494336 A
^ 03/31/2018 NCCA 2015 Tracking - ECO Fish Tissue T5
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 125
TRACKING: PACKS
A NCCA 201 5 TRACKING: PACKS A
Sender: Sender Phone: _ _
State of Site Location: Crew:
Shipped By: O FedEx O UPS Q Hand Delivery Date Sent: / /
Airbill/Tracking Number:
Site ID
Date Sample Collected
MMIDD/YYYY
Visit
O1
O2
O1
O2
O1
O2
O1
O2
O1
O2
O1
O2
O1
O2
O1
O2
CM
02
01
02
01
02
Oi
02
01
02
01
02
01
02
01
02
CM
O2
O1
O2
Comments
Packet Lab ' Completed by Lab Save completed form as: Tracking Related Inquiries:
Attn: Marlys Cappaert
C/Q USEPA - WED Divisit
200 SW 35th St
Corvallis, OR 97333
Email:
cappaert.marlys@epa.go
| Date Received: NCCA15_T6_TrarkiriL_SitelDVS Marlys Cappaert
>n i , Phone:541-754-4467
Email to : samp let rackine@epa.gov '
Received by: • E i
Michelle Gover
Or fax to1 541 754 46'7 Phort&1 541 -7e>4-47cl3
/
i i
• 1857175712 -r- A
03/31/2015 NCCA 2015 Tracking - PACKS I D ^f
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 126
TRACKING: HUMAN HEALTH WHOLE FISH SAMPLE - OVERNIGHT (DRY ICE)
m NCCA 2015 TRACKING: HUMAN HEALTH WHOLE FISH SAMPLE - OVERNIGHT (DRY ICE) "^
Great Lakes Only
State of Site Location: Crew Date Sent: / /
Sender:
Sender Phone: - —
Shipped by: O FedEx O Hand Delivery Airbill/Tracking Number:
Site ID:
Visit: Q1 Q 2 Date Collected: j
./
FISH ALL WITHIN 75% OF LARGEST SPECIMEN Q
FISH ARE ALL THE SAME SPECIES Q
Sample ID
Total Length (mm) Total Length (mm) Total Length |mm)
Frozen: Q .01
Scientific Name (Genus Species)
.03
The human health fish eomposil
adequate size to provide a total
about 18 oz.) of fillet tissue. Few
meet the fillet tissue weight requ
.04
weight of 500 grams (equivalent to Q^
fer or more fish can be collected to
Irement. ~~
,w
.07
.08
.11
.12
.13
.14
.15
I
.17
\^ .18
.OJ .19
.10
.20
^^^^^^
Comments:
.21
.22
.23
.24
.25
.26
.27
.28
.29
.30
s
Lab
Save completed form as: Tracking Related Inquiries:
Attar Michael Arbaugh NCCAl5_T7_Tracking_SrtelD_V« Marlys Cappaert
c/o Microbac Laboratories f*10"6 : 541 -754-4467
2101 Van Deman Street Email to : sampletracking@epa.gov
Baltimore, MD 21224 ^helle Gover
41 0-633-1 800 Or fax to: 541-754-4637
^ 7896567T83
03y31/2015 NCCA 2015 Tracking - Human Health Fish Tissue Fillet T7 ^
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 127
TRACKING: UVID
A NCCA 2015 TRACKING: UVID A
Sender: Sender Phone: _ _
State of Site Location: Crew:
Shipped By: Q FedEx O UPS O Hand Delivery Date Sent:
• *
Airbill/Tracking Number:
Site ID
Date Sample Collected
MMIDD/YYYY
Visit
O1
O2
O1
O2
O1
O2
O1
O2
O1
O2
O1
02
01
02
01
02
CM
O2
O1
O2
O1
O2
Oi
02
01
02
01
02
01
02
01
02
CM
O2
O1
O2
file Name
UVID Lab Completed by Lab Save completed forms as:
USEPA - MED
6201 Cong don Blvd
Duluth, MN 55804
Phone: 218-529-5122
Contact: Julie Lietz
Date Received:
/ ;
Received
NCCA15_T8_Trac ki ng_Site I D_VS
Email to : sampletracking@epa.gov
Or fax to: 541-754-4637
/ /
Comment?
Tracking Related Inquiries:
Marlys Cappaert
Phone: 541-754-4497
Michelle Gover
Phone: 541-754-4793
• 1025389514 xo A
03/31/2015 NCCA 2015 Tracking -UVID lo ^f
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National Coastal Condition Assessment 2015 Field Operations Manual
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APPENDIX C: SHIPPING AND TRACKING GUIDELINES
TRACKING FORMS
Each tracking form has been assigned a "T" page number to help crews identify the correct
tracking form to use when sending samples. This "T" number is located on the bottom right
corner of each tracking form. Crews will also find reference to the same "T" numbers on the
individual samples labels and on the top of the pre-printed FedEx return labels provided in
the site kits.
Crews include copies of all tracking forms in the coolers when they send samples to the labs.
They have several different options for electronically submitting sample and tracking
information. A hard copy of the sample tracking form must be submitted to the lab in the
cooler and an electronic copy must be submitted to NARS IM using one of four options. If a
cooler contains samples from more than one site, then multiple forms must be placed in the
cooler and submitted to NARS IM.
In order of preference, the options are:
1. Using the NCCA mobile App, enter data and tracking information into the NCCA App
and submit the tracking information. An email will pop up on your device with the
NARSFieldData@epa.gov address. Copy yourself, any other crew members or managers,
and click send. This form will be returned to you via email after a few minutes in a
portable document format (PDF). It may be printed and used as the form for the
cooler shipment.
2. Using a handheld device or portable computer, enter data into a fillable PDF form,
save, and submit it via email. Please name these files in the following format:
NCCA15_T#_Tracking_SitelD_V#, where T#' is the number of the tracking form and
W is the visit number (i.e. NCCA15_T1_Tracking_NCCA15-1061_V1) before emailing or
using the SUBMIT button. Send the file via email to sampletracking@epa.gov. It may be
printed and used as the form for the cooler shipment.
3. Hand-enter data on a paper form. Photograph or scan the form with a handheld device
or office scanner. Attach the file (in PDF version) to an email and address to
sampletracking@epa.gov. Please name these files in the following format:
NCCA15_T#_Tracking_SitelD_V#, where T#' is the number of the tracking form and
W is the visit number (i.e. NCCA15_T1_Tracking_NCCA15-1061_V1). Be sure that the
file scanned is clear and legible. Genius Scan or Cam Scanner are great apps that are
available for free that will help to ensure that the scan is clear and legible. After
scanning, include the form in the cooler.
4. Hand-enter data on a paper form. Fax the form to the number printed on the form.
After faxing, include the form in the cooler.
It is very important to submit the Site and Sample Status/Water Chemistry Lab Tracking
form immediately after every sampling event. Prompt status reports allow the FLC to closely
track sampling progress. More importantly, it enables NARS IM to track samples that were
collected at each site versus those that were not, and to immediately track the shipment of
the time-sensitive samples after each sampling event.
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National Coastal Condition Assessment 2015 Field Operations Manual
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If the crew visits a site with the intention of sampling, but determines the site to be
unsampleable (either temporarily or permanently), the site status portion of the Site and
Sample Status/Water Chemistry Lab Tracking form needs to be completed and submitted,
but the Water chemistry and Batch sample status tracking portion of the form can remain
blank.
Daily Form:
Tl - SITE & SAMPLE STATUS/WATER CHEMISTRY LAB TRACKING FORM
• Complete the Site and Sample Status/Water Chemistry Lab Tracking form for
the samples that are shipped immediately after each sampling event (water
chemistry (CHEM), chlorophyll A (CHLA), dissolved nutrients (NUTS)).
• Send an electronic copy of this form to NARS IM using one of the options listed
above. This serves as the "status report" for that sampling event.
• Ship all of the samples to the lab in the same cooler with a hard copy of this
form.
• Samples from two sites may be shipped together in a single cooler if they were
collected on the same day.
• Samples need to be shipped on fresh wet ice.
• Water chemistry samples should be shipped within 24 hours of collection.
Batch Forms:
• Crews may hold BATCHED samples and ship them within the designated time
frame.
• Electronically send the tracking form(s) to NARS IM when the samples are
SHIPPED using one of the options listed above.
• Use one form for each site's worth of samples in the cooler, (i.e. if you have
batched samples from 4 sites in the cooler, there should be 4 forms
completed).
• Include paper copies of the forms in the cooler.
• All samples in the cooler should be listed on one of the included tracking
forms.
T2 -TRACKING: BATCH SAMPLES - OVERNIGHT (CHILLED) FORM
• Use this form for shipping batches of chilled samples:
• Sediment toxicity
• Sediment grain size
• Phytoplankton (Great Lakes sites only)
• Up to 3 site's worth of samples may be shipped together in a single cooler.
• Samples need to be shipped on fresh wet ice
• Chilled batched samples should be shipped at least every week
T3 -TRACKING: BATCH SAMPLES - OVERNIGHT (FROZEN) FORM
• Use this form for shipping batches of frozen samples:
• Microcystins
• Algal Toxins
• Enterococci
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• Fish tissue plugs
• Sediment TOC
• Sediment Organics/Metals
• Ecofish samples (may be shipped separately)
• 2-3 site's worth of samples may be shipped together in a single cooler,
depending on whether the ecofish sample is included and the size of the fish
comprising that sample.
• Samples need to be shipped with approximately 20 pounds of dry ice
• Frozen batched samples should be shipped at least every 2 weeks
T4 -TRACKING: BATCH SAMPLES - GROUND (No ICE) FORM
• Use this form for shipping batches of non-chilled samples:
• Benthic Macroinvertebrates
• Up to 12 site's worth of samples may be shipped together in a single cooler,
depending on whether more than one bottle of sample was collected at a site.
• Samples need to be shipped with absorbent material and no ice.
• Non-chilled batched samples should be shipped every 2-3 weeks.
NOTE: Federal regulations and FedEx rules allow for ground shipping of certain quantities of
flammable liquids WITHOUT the need for special certifications and labeling. Flammable
liquids may NOT be shipped via air carrier unless shipper is trained and qualified to do so and
specific documentation and labeling requirements are met.
The Code of Federal Regulations (49 CFR Section 173.150) lists the exceptions which allow
shipping of flammable liquids via ground carrier without labeling or special certifications.
Ethanol and formalin can be considered to be in either Packaging Group 2 or 3, so we use the
more stringent PC 2 as our guideline. The limited quantity exclusion allows ground shipping
of PC 2 flammable liquids provided that the individual containers inside the package are not
over 1.0 liters each, that the gross weight of the package does not exceed 66 pounds, and
that the outer packaging is a sturdy container. Please ensure that your shipment meets these
criteria to ensure the legal ground shipment of these samples.
T5 -TRACKING: Eco FISH TISSUE - OVERNIGHT (DRY ICE) FORM
• Use this form for shipping batches of frozen eco fish samples:
• Eco Fish samples may be sent in the same cooler as the other frozen batched
samples listed above or may be sent separately.
• 2-4 site's worth of samples may be shipped together in a single cooler,
depending on whether eco fish are included and the size of the eco fish
sample.
• Samples need to be shipped with approximately 20 pounds of dry ice
• Frozen batched samples should be shipped at least every 2 weeks
T6 -TRACKING: PACKS FORM
• If utilizing paper field forms, review and ship all field forms in the envelope
provided in the site kit to NARS IM every 2 weeks.
• Before shipping, make copies or scans for your records and as a backup in the
event the forms are lost during shipping.
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Field Operations Manual
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T7 -TRACKING: HUMAN HEALTH WHOLE FISH SAMPLE - OVERNIGHT (DRY ICE) FORM [SELECT
GREAT LAKES SITES ONLY]
• Use this form for shipping frozen human health fish tissue samples.
• More than one site's worth of samples may be shipped together in a single
cooler, depending on the size of the fish.
• Samples need to be shipped with 50 pounds of dry ice.
• Human health fish tissue samples should be shipped within 2 weeks of
collection.
T8 -TRACKING: UVID FORM [GREAT LAKES ONLY]
• Use this form for shipping the EPA-provided USB flash drive containing all
underwater video recorded during the season.
• Before shipping, make copies of the video files for your records and as a
backup in the event the forms are lost during shipping.
SHIPPING GUIDELINES
Samples will be shipped according to the chart in Appendix C: Shipping and Tracking
Guidelines. The Field Crew Leader will complete the appropriate tracking form for the
samples and will submit tracking via one of the options listed in the tracking forms section
above. The Field Crew Leader will place the samples and the tracking form (in a waterproof
bag or plastic sleeve) in a shipment cooler. The Field Crew Leader will attach the appropriate
pre-addressed airbill from the site kit marked for the appropriate lab. The field crew will
either drop off the cooler for shipment at a local FedEx location or arrange for a pick up at
the hotel or other appropriate facility. If the field crew has chosen a pick up, they must
follow up with the facility at which it has been left and/or track the package through FedEx
tracking tools to ensure its actual pick up. Once the package is in the possession of FedEx, the
IM Team and FLC will track the package to its destination and take steps necessary to ensure
its timely delivery. Prior to shipping, there are a few other guidelines to be aware of:
Preservation
Holding Time
Shipping
• See chart for specific
preservation information
for each sample
• Note the holding time
window for each sample
• Ensure that samples will
be shipped in time for the
lab to be able to process
them within the
allowable holding time
frame
• Samples may be shipped
on wet ice, dry ice, or
with no ice
• Secure the cooler with
strapping tape
• See dry ice shipping
protocols
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Ensure that the ice is fresh immediately prior to shipment;
Line the cooler with a large plastic liner bag. Double bag the ice with enough
white or clear 1 gallon zippered plastic bags to pack the entire cooler.
To prevent misidentification of any water leakage as a possible hazardous
material spill, use an indelible marker to label all bags of ice as "ICE".
Place bagged samples and bags of ice inside the cooler liner and seal the liner.
Secure the cooler lid with strapping tape.
1 Note: Not all FedEx locations will accept shipments containing dry ice. Dry ice
shipments can be shipped from "FedEx staffed" locations. You can also arrange
for a pick-up from your lab or hotel. Dry ice shipments usually cannot be shipped
from FedEx Kinko's Office and Print Centers® or FedEx Authorized ShipCenter®
locations. These types of locations are differentiated on FedEx.com in the "Find
FedEx Locations" feature. Please be sure to call in advance to ensure your location
will accept the package for shipment.
> Attach the provided FedEx airbill:
• Ensure that the label indicates the amount of dry ice in the package.
> Label the cooler with a Class 9 Dangerous Goods label
• Place the label on the front side of the
cooler, not the top.
• If it is not already completed, fill out the
upper corners of the label with the same shipper
and recipient information as on the FedEx airbill.
• Declare the weight (in kg) of the dry ice in the lower
right hand corner of the label, ensuring it is the same
weight listed on the airbill.
> Secure the cooler lid with strapping tape. Do not completely seal the entire edge
of the cooler such that pressure inside the cooler could build.
> Place the provided FedEx airbill on the top of the cooler or on a handle tag
secured to one of the cooler's handles.
Surround the jars with crumpled newpaper, vermiculite or other absorbent
material
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National Coastal Condition Assessment 2015
Version 1.0 May 2015
Field Operations Manual
Page 133
Water Chemistry
[CHEM]
•Ship within 24 hours
•Ship 250 mL amber HOPE bottle
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler liner
•Ship on wet ice
Chlorophyll-a[CHLA]
•Ship with CHEM/NUTS samples
•Ship foil wrapped centrifuge tube
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler liner
•Ship on wet ice
Dissolved Nutrients
[NUTS]
•Ship within 24 hours
•Ship 250 mL HOPE bottle
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler liner
•Ship on wet ice
Sediment Grain Size
[SEDG]
•Ship within 1 week
•Ship in plastic bag (quart size, double bagged)
•Confirm label completed & taped
•Place in lined cooler with other chilled batched samples
•Ship on wet ice
Sediment Toxicity
[SEDX]
•Ship within 1 week
•Ship in screw-top bucket (0.6 gal)
•Confirm label completed & taped
•Tighten the lid securely making sure the ratcheting
mechanism engages
•Place in lined cooler with other batched samples.
•Ship on wet ice
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Version 1.0 May 2015
Field Operations Manual
Page 134
Phytoplankton [PHYT]
(GLonly)
•Ship within 1 week
•Ship in HOPE bottle (1 L, white, narrow mouth)
•Confirm preserved with 10ml Lugol's solution
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler lined cooler with other chilled batched samples
•Ship on wet ice
Algal toxin [ALGX] and
Microcystin [MICX]
•Ship at least every 2 weeks
•Freeze after collection
•Ship in HOPE bottle (500 mL, white, wide-mouth)
•Confirm labels completed & taped
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 Ibs of dry ice
Enterococci [ENTE]
•Ship at least every 2 weeks
•Ship in frozen, microcentrifuge tubes
•Confirm labels completed
•Place each tube in small bubble bag with label on outside
•Place bags in zip-top bag
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 Ibs of dry ice
Sediment TOC [SEDC]
•Ship at least every 2 weeks
•Ship in frozen, glass jar (60 mL) (leave headspace)
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place jar in foam sleeve
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 pounds of dry ice. Pack with fill material
such as newspaper if necessary to ensure no shifting
Sediment
Organics/Metals
[SEDO]
•Ship at least every 2 weeks
•Ship in frozen, glass jar (120 mL) (leave headspace)
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place jar in foam sleeve
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 pounds of dry ice. Pack with fill material
such as newspaper if necessary to ensure no shifting
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Version 1.0 May 2015
Field Operations Manual
Page 135
Ecological Whole Fish
Tissue [FTIS]
•Ship at least every 2 weeks
•Freeze after collection, as soon as possible (-20 cooler)
•Ship in bags
•Confirm label completed & taped
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 Ibs of dry ice
Fish Plugs [FPLG]
Benthic
Macroinvertebrates
[BENT]
•Ship at least every 2 weeks
•Freeze after collection
•Ship in glass scintillation vial
•Confirm label completed & taped
•Place vial in small bubble bag
•Place bubble bag in zip-top bag
•Wrap packing material around bag to prevent breakage
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 Ibs of dry ice
•Ship every 2-3 weeks
•Preserve benthos samples immediately upon collection
•Ship in HOPE bottle (1 L, white, wide mouth)
•Confirm label completed & taped
•Seal with plastic electrical tape
•Surround the jars with crumpled newpaper, vermiculite or
other absorbent material
•Place in cooler liner
•Ship with NO ice
Human Health Whole
Fish Tissue [HTIS]
(select sites GLonly)
•Ship at least every 2 weeks
•Freeze after collection, as soon as possible (-20 cooler)
•Ship in bags
•Confirm label completed & taped
•Pack cooler with 50 Ibs of dry ice
Underwater Video
[UVID] (GLonly)
•Ship at end of season
•Transfer files from DVR system to EPA-provided flash drive
•Back up files to computer hard drive
•Be sure files are named appropriately
•Package EPA-provided flash drive(s) in padded envelope
securely
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Version 1.0 May 2015
Field Operations Manual
Page 136
/^ ^\
[Tl]
"WRS" Samples
1 - HOPE bottle (250
mL, amber)
CHEM
1 - Filter in centrifuge
tube (50 mL), in zip-
top bag
CHLA
1 - HOPE bottle (250
mL, white)
NUTS
/" ^\ S^
[T2]
Chilled Samples
(Batched)
1 - Plastic bag (quart
size, double
bagged)
SEDG
1 - Screw-top bucket
(0.6 gal)
SEDX
1 - HOPE (1L, white
narrow mouth)
PHYT
^^ f^ ^^i f^
^^
[T3] 1
Frozen Samples (Batched)
1 - HOPE bottle (500 mL, white, 1 - Glass scintillation
wide-mouth) vial with 2 plugs
MICX FPLG
1 - HOPE bottle (500 mL, white, 1 - Glass jar (60 mL)
wide-mouth) SEDC
ALGX i _ Glass jar (120 mL)
2 - Filters in centrifuge tubes in SEDO
bag (plus 1 blank if revisit site}
ENTE
V
r
^S
~
[T5] Eco Fish Samples (Batched)
5-20+ fish in large plastic composite bag FTIS
May be shipped with other frozen samples (T3) if
desired, or can be shipped separately
i
\ i
^
r "N
[T4]
Non-Chitted
Samples
(Batched)
1 or more -
HOPE
bottle
(1 L, white,
wide-
mouth)
BENT
V J
r ^\
[T6]
Completed
Data Packs
(Batched)
All data forms
from a site,
reviewed for
completeness,
legibility and
accuracy
PACK
r ^\
[T7]
Frozen HH
Whole Fish
Samples
(Batched)
5 - Fish in large
plastic
composite
bags
HTIS
V J \^ J
r "x
[T8]
UW VIDEO
(Batched)
1 - File
UVID
V J
._4_ __4_ __4_ ._4_
Pack 1 day's worth Pack 2-3 sites' Pack 2-4 sites' worth of samples Pack up to 12 Make copies or Pack frozen Put onto SD
of samples (up to 2 worth of samples in (depending on whether Eco Fish are sites ' worth of soa"s as bac^uP- fish in cooler card- backuP to
sites) in lined cooler lined cooler with ice included and size of Eco Fish sample) in samples in lined form number. with 50 °pi^ein
with ice in baes in bags cooler with dry ice insert and dry ice cooler with Place in pounds of vadded
j^^^f
|b^ ICE
1 -^^^fc^^^"
ICE^ ^^"""^
I 1
r r
SHIP WITHIN 2 4
HOURS
(MON-FRI)
FedEx*
Express
PRIORITY
OVERNIGHT
SHIP WEEKLY
(MON-FRI)
FedEx®
Express
PRIORITY
OVERNIGHT ,
j\/O ICE envelope with dry ice envelope with
trackingform offy tracking form
DRY
^® %3 pf3- ^
L T i
SHIP WITHIN 2 WEEKS |
(MON-FRI)
Fe
rV^t^V®
Express
PRIORITY OVERNIGHT
J
SHIP EVERY
2-3 WEEKS
ANYTIME
E ^
rectx
Ground
GROUND
^ ^
f ~
SHIP
EVERY 2
WEEKS
Express
r ~
SHIP MO N
- THURS
(No
Saturday
Delivery)
fedEx
Express
PRIORITY
OVERNIGHT
r ~
SHIP AT
END OF
SEASON
FecEx
Express
L. ^
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Version 1.0 May 2015
Field Operations Manual
Page 137
SAMPLE SAMPLE CONTAINER PRESERVATIVE PACKAGING HOLDING TIME
TARGET FOR
VOLUME SHIPMENT
WRS
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SAMPLE SAMPLE CONTAINER PRESERVATIVE PACKAGING HOLDING TIME
TARGET FOR
VOLUME SHIPMENT
DATA
FORMS
(T6)
HH
WHOLE
FISH*
(TV)
UW VIDEO
(T8)
Data Packets
(PACK)
Human Health
Whole Fish Tissue
Sample
(HTIS)* - Great
Lakes only
Underwater Video
(UVID) - Great
Lakes only
1 completed field
form packet
5 whole fish (500 g
of fillet weight)
1 minute video
Organize in proper
order
Put in envelope
from site kit
Wrapped
individually in
solvent rinsed foil
Sealed in poly
tubing
Large outer plastic
bag
Download from
DVR to USB flash
drive via computer,
send flash drive
N/A
Dry ice in field; Hold
in freezer
N/A
Ship in envelope
Ship in cooler
provided with
50 pounds of
DRY ICE
Ship in padded
envelope
Batch, ship monthly
to NARS IM
Batch, ship weekly
(except on Fridays,
Saturdays, or the day
before Federal
holidays) to HTIS
lab
Batch, ship at end of
season to EPA
Duluth lab
' Human Health Fish Tissue is collected at select Great Lakes sites only
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Field Operations Manual
Page 139
APPENDIX D: FIELD EVALUATION AND ASSISTANCE VISIT
CHECKLIST
Evaluation Date(s):
NCCA2015SITEID:
Location:
Evaluation Team Member(s):
Name
Organization
Phone
Field Crew ID:
Name
Organization
Phone
Other Observers Present During Evaluation:
Name
Organization
Phone
Please send completed form to Colleen Mason:
o Email scanned document (pdf): mason.colleen@epa.gov (preferred)
o OR fax: 202-343.9641 (If faxing please leave a message at 202-566-0417 so Colleen
knows to check the fax machine)
o OR mail hardcopy (and keep a copy) to:
Colleen Mason
EPA
1200 Pennsylvania Ave., NW (4503T)
Washington, DC 20460
If major corrective actions are required, please email Colleen Mason and Hugh Sullivan
(Sullivan.hugh@epa.gov) and provide:
o brief summary of the areas of concern
o best dates/times for a teleconference to discuss the concerns and resolution.
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PREDEPARTURE ACTIVITIES
Equipment and Supply Preparation
Did the crew request a site kit at least two weeks prior to sampling? Was
the site kit available for the site?
Great Lakes Only: Did field crews request a Great Lakes human health
fish tissue supply kit for appropriate sites (if applicable) or know that
they must do so?
Was the supply kit available for the site?
Did the crew have phytoplankton bottle(s) available for sampling?
Refer to Appendix A of the Field Operations Manual. Does the crew have
all of the required equipment and supplies?
Did the crew have back-up site kit(s) available during the sampling?
Did the crew obtain sufficient wet and dry ice for sample preservation
and storage? Record the amount of ice in Ibs: Dry: Wet:
Are the meters, probes, and sampling gear packed in such a way as to
minimize physical shock and vibration during transport?
Are copies of the Field Operations Manual, the Quick Reference Guide,
equipment manuals, etc. available?
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Preservatives and other Solutions
Are the recipes for stock preservatives readily available for crew
reference?
Were stock preservatives prepared?
Did the crew pack stock solutions as described in FOM Table 4.1? (Bleach,
100% stained buffered formalin, etc.)?
Y
Y
Y
N
N
N
N/A
N/A
N/A
Site Information and Access
Did the crew verify that it had completed the site evaluation spreadsheet
for sites dropped based on desktop recon?
Did the crew verify that replacement sites were chosen from the same
stratum/base year combination
Was the Verification Form completed for sites visited with the intent to
sample but not sampled?
Were individual site packets, including directions to the site, available
and organized?
Was the site access information/permission letter available?
Was the landowner contacted prior to site visit, if applicable?
Were other key contact persons notified (e.g., Regional Coordinator,
State or Tribal contacts)? Identify who was notified:
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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Vehicle/Boat
Did the crew perform necessary checks to the vehicle before leaving for
the day?
Were the trailer and hitch inspected prior to departing to the site to
ensure that the trailer was securely fastened?
Was the boat(s) in good working order and inspected before departure?
Were PFDs available for all passengers?
Y
Y
Y
Y
N
N
N
N
N/A
N/A
N/A
N/A
Global Positioning System Receiver
Were the batteries checked? Are spare batteries available (if applicable)?
Did the crew verify that any additional tests/checks recommended by the
operating manual were performed?
Y
Y
N
N
N/A
N/A
Multi-Probe
Were the sensors stored properly to prevent damage and desiccation?
Was the multi-probe meter inspected according to the
manufacturer's specifications?
Did the crew confirm that the accuracy of the temperature sensor was
checked against a thermometer that is traceable to the National Institute
of Standards at least once per sampling season? Record the date of the
last test:
Y
Y
Y
N
N
N
N/A
N/A
N/A
Photosynthetically Active Radiation (PAR) meter
Were the batteries checked?
Was the PAR meter assembled as described in the FOM (check sensor
connections and positions)
Were the correct calibration factors entered for each probe? (These
factors are supplied by the manufacturer and are specific to each
individual probe.)
If the probe is not new, has it been returned to the manufacturer and
calibrated within the last two years? (simply ask the crew, we do not
need written proof). Date calibrated:
Was a weight attached to the underwater probe frame so it hung
vertically?
Was the sounding line to be used with the PAR marked at least every 0.5
meters? (These marks should indicate the distance to the underwater
sensor)
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
Containers/Labels
Were labels affixed to containers and covered with clear tape when
required (pre-labeling is recommended)?
Were labels completed where feasible and appropriate (before or after
collection) using a permanent marker (pencil for benthos inside jar label)
and covered with clear tape?
Y
Y
N
N
N/A
N/A
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Field Operations Manual
Page 142
PREDEPARTURE ACTIVITIES NOTES
BASE AND LAUNCH SITE ACTIVITIES
Instrument Calibration
Was the DO calibration done at the launch site or other on-site location
(in accordance with section 6.3.1)?
Were the calibration values recorded on the data sheet?
Was the pH and salinity/conductivity calibration conducted and the
values recorded on the data sheet?
Were manufacturer recommended internal diagnostic checks of the meter
performed within the last week to ensure correct meter function (e.g.
cell constants, millivolt output, or other readings)?
Is the instrument not able to be calibrated in the field, but was factory
calibrated before field measurements were taken (i.e. Seabird)?
Date calibrated:
Instrument number/Serial number:
Does documentation match instrument identification/serial number (AV
evaluator may need to contact office or lab separately if documentation
not carried into field)?
If the internal meter checks were not done, was a commercially available
Quality Check Solution (QCS) used within the last week to verify values of
pH and conductivity?
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Other Preparations
Were the Enterococci filter microcentrifuge tubes with beads placed on
dry ice before filtering commenced?
Was a cooler(s) prepared with wet ice for storing samples?
Y
Y
N
N
N/A
N/A
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BASE AND LAUNCH SITE ACTIVITIES NOTES
SITE VERIFICATION
Site Verification at the Launch Site
Are the location coordinates the same in the crew's paperwork and EPA's
spreadsheet of target sites?
Was a description of the final part of the route to the site recorded on the
site verification form?
Was the arrival time (and later the departure time) recorded on the site
verification form?
Were the names of the field crew recorded?
Y
Y
Y
Y
N
N
N
N
N/A
N/A
N/A
N/A
Site Verification at the Index Site Location
Was the site classified correctly (e.g., target vs. nontarget vs.
inaccessible)?
Were the target coordinates (X-site) from the site list entered into the
crew's GPS unit?
Did the crew navigate to within O.OZnm or 37 meters from the X site?
If the initial sampling location was not sampleable, did the crew use the
steps outlined in the FOM to attempt locate a sampleable location within
the 37 meter radius (see Section 5.1 .3)?
Was the GPS checked after anchoring the boat to ensure the location was
within 37 meters?
Were the GPS coordinates of the initial sampling location (Y-Location)
recorded on the verification form?
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
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Was the GPS consulted periodically to ensure the sampling vessel had not
drifted further away from the X-site due to anchor drag?
N
N/A
Was the habitat assessment information completed?
N
N/A
Were photographs of the site taken if site has unique characteristics
(optional)?
N
N/A
SITE VERIFICATION NOTES
Y-LOCATION SAMPLING
Did the crew collect the probe and water column data as close to the X-site as
possible, but no further than 37 m?
Y
N
N/A
Dissolved Oxygen, pH, Temperature, Salinity/Conductivity
Was the depth measured at the Y- location, and the intervals calculated before
probe was placed in the water?
Were all measurements allowed to stabilize before recording?
On the downcast, were the measurements at each depth interval conducted
and recorded according to the protocol (0.1m, 0.5m, every meter from 1.0 to
10.0 meters and every 5 meters thereafter)?
Were the recorded data entered on the Field Measurement Form or saved as an
electronic file in the instrument?
If the crew will be submitting the hydrographic profile via an electronic file,
was the "Submitted data via eFile" bubble filled in?
Was the last measurement taken at 0.5 meters from the bottom?
Did the probe touch the bottom?
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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On the upcast, were the measurements at each depth interval conducted and
recorded according to the protocol on the Field Measurement Form (using the
exact same depths as above)?
Did the crew flag any measurements that could not be made or that required
further comment?
Was the probe stored correctly after the measurement?
Y
Y
Y
N
N
N
N/A
N/A
N/A
Secchi Disk Transparency
Was the Secchi disk being used the black and white 20 cm patterned disk?
Was the calibrated sounding line visibly marked in at least 0.5 meter intervals?
Were Secchi depths recorded to the nearest 0.1 meter?
Was the measurement taken from the shady side of the boat?
Was the recorder wearing sunglasses or a hat? (should not have any on)
Was a viewscope used? (should not be used)
If the disk could be seen at the bottom, did the crew mark the "clear to
bottom" bubble and record the station depth as both the disappearance and
reappearance depth for Reading 1?
If any one of the three sets of measurements varied more than 0.5 meters from
the others, was the entire process repeated?
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Photosynthetically Active Radiation (PAR) Meter
Verify that the correct calibration factors were entered for the probe, unit is
set up correctly, and underwater sensor is plugged in correctly.
Was the deck sensor placed on the boat on a non-shaded location?
Was the underwater sensor lowered on the sunny (or least shaded) side of the
boat to a depth of 10cm?
Was the underwater probe lowered by means of the rope attached to the probe
frame, not the cord?
Were both the ambient and underwater sensor readings taken at the same
instant?
Indicate how (e.g., data logger, two operators, camera):
Were both the deck and underwater sensor readings recorded at each of the
other depths calculated for the hydrographic profile?
If the underwater sensor hit the bottom, did the crew wait 2-3 minutes before
taking the reading?
If the light measurements became negative, was the profile terminated at that
depth?
Was the process repeated on the upcast?
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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Fecal Indicator (Enterococci) Sample Collection
Was the enterococci sample collected at a time to most effectively meet the
needs of both protecting the sample from potential contamination and meeting
the 6 hour holding time (figures 3.1 and 3.2)?
Were new, clean gloves worn?
Was the sample collected by hand or pole? Circle one:
Was the 250 ml sample bottle lowered un-capped and inverted to a depth of
0.5 meters below the water surface, avoiding surface scum, vegetation, and
substrates?
Was the mouth of the container pointed away from the body or boat?
Was the bottle righted and raised through the water column, allowing the
bottle to fill completely?
If a pole was used (larger vessel) was the pole cleaned and rinsed prior to
sampling?
If a pole was used, was the bottle attached in such a way to avoid
contamination?
If a pole was used, was the bottle plunged quickly to a depth of 0.5 meters and
allowed to fill?
After removing the container from the water, was a small portion of the
sample discarded to allow for proper mixing before analysis?
Was the sodium thiosulfate tablet added along with the cap, and the bottle
shaken 25 times?
Did the crew check that the tablet was dissolved?
Was the sample stored in a cooler on ice to chill (not freeze)?
Was the collection time and depth collected recorded correctly on the field
form?
Was the sample chilled for at least 15 minutes before filtering?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Water Sample Collection and Preservation
Were clean gloves worn?
Did the crew avoid applying sunscreen or other chemicals until after the
sample was collected (or implement measures to reduce contamination by such
chemicals if applied such as washing, wearing long gloves, etc.)?
Was the pumped system or water sampling bottle rinsed three times?
Was water collected from 0.5 meters below the surface?
Were bottles rinsed three times with ambient water before collecting final
sample?
Was a 2 L HOPE bottle (brown) filled with sample water and placed in a cooler
on wet ice? (chlorophyll a)
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
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Fecal Indicator (Enterococci) Sample Collection
Was the 250 ml brown bottle filled with sample water, the correct label
affixed, label information correctly filled out and the label taped over with
clear tape? (water chemistry)
N
N/A
Were two 500 ml HOPE bottles filled with sample water, the correct labels
affixed, label information correctly filled out and the labels taped over with
clear tape? (microcystins and algal toxins)
N
N/A
Did the microcystins and algal toxins bottles have at least an inch of headspace
to allow for expansion during freezing?
N
N/A
Were the samples placed on wet ice in a dark cooler?
N
N/A
Were comments about anything that could influence the sample chemistry
(heavy rain, etc.) included in the comments section of the sample collection
form?
N
N/A
GREAT LAKES ONLY: Was a 1L narrow mouth, white HOPE bottle filled with
sample water for phytoplankton?
N
N/A
GREAT LAKES ONLY: Were approximately 10 ml of Lugol's added to the 1L
bottle for phytoplankton preservation within 2 hours of collection?
N
N/A
Y-LOCATION SAMPLING NOTES
UNDERWATER VIDEO FOR GREAT LAKES ONLY
Was the camera deployed during the same time period as the in situ
measurements and water collection activities occurred?
Was the camera deployed at the Y-Location?
Was the GPS unit powered up and displaying 'ready to navigate' prior to
starting?
Y
Y
Y
N
N
N
N/A
N/A
N/A
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Were the 12v batteries attached to the GPS overlay and DVR?
Was the GPS overlay turned on?
Was the camera deployed on the windward side of the boat?
Were two crew members used in the operation (one to lower the camera and
one to operate the DVR and instruct the first person on descent speed and
depth)?
Was at least one minute of good footage captured that provides a clear view of
the bottom and a 360 degree sweep of the bottom?
If the bottom was a 'low light' situation, were the camera lights activated?
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
Archiving Underwater Video FOR GREAT LAKES ONLY
Was the underwater video clip properly archived/saved and file name recorded
on the Sample Collection form (Section 11.2.4)?
Was the video reviewed for quality?
If the video was of poor quality or unviewable, was another video taken?
Was the system properly shut down (DVR, GPS overlay, camera, and GPS)?
Was the battery recharged (or will it be before the next day's use)?
Were the files downloaded to the EPA-provided USB flash drive following the
process in the field operations manual (Section 11.2.5)?
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
UNDERWATER VIDEO NOTES
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Sediment sampling
Relocating for Sediment Collections (if Required)
If no successful sediment grabs could be collected first at the Y-Location and
then at other locations within 37 m of the X-site, did the field crew properly
make attempts within 100 meters as described in the field operations manual?
If no successful sediment grabs were made in either the primary or secondary
sediment sampling locations, did the field crew properly move to and make
attempts within 500 meters of the X-site as described in the field operations
manual?
Did the crew correctly identify the distance at which the sample was collected
on the field form?
Y
Y
Y
N
N
N
N/A
N/A
N/A
Sediment Collections
Does the sampler have a hinged or otherwise removable top?
Was the dimension and sample area of the grab recorded on the field form?
Was the sampler washed with Alconox prior to sampling and then rinsed with
ambient water?
Was the sieve thoroughly cleaned between sites to ensure no cross-
contamination of samples?
Was the grab sampler lowered so that the last 5 meters is no faster than about
1 m/sec?
Was the cable allowed to go slack once the substrate is reached?
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
Benthic Macroinvertebrate Collection
Was the top of the sampler opened to determine whether the grab was
successful?
Was the sediment depth in the middle of the sampler recorded (should be
>7cm)
If grabs of 7 centimeters could not be obtained after several tries, was the
next successful grab used regardless of depth and was this flagged on the field
form?
Were notes on the condition of the sample recorded?
Was the overlying water carefully drained into the container that will receive
the sediment? Describe how this was done in the comments section.
If the grab sampler is less than 0.03 nV, were two grabs for benthics taken and
composited into the sieve?
Was the sediment dumped into a basin and then into a 0.5 mm mesh?
Was the sieve placed in a sieve box or other appropriate medium and the tray
agitated to wash away sediments?
Was care taken to avoid loss of sample over the side of the sieve?
Were large non-living items inspected for organisms and then removed?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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Was the bulk of the sample gently scooped up and placed in a 1 -L Nalgene
bottle?
Was the outside of the sample jar rinsed into the sieve, then the contents
rinsed into the sample jar using a funnel?
Was the sieve inspected to make sure all organisms are transferred to a
container?
Were all sample jars filled no more than Vi full with sample?
If more than one jar is needed are they appropriately labeled (e.g., 2 of 2)?
Is all information correctly recorded on sample labels and field form?
Was a sample label, completed in pencil, placed inside the sample jar?
Are all containers preserved with a minimum of 100 ml of stained buffered
formalin solution and an additional teaspoon of borax added? (End
concentration of the preservative should be at least 6 percent)
Was each jar filled to the rim with seawater/lakewater to eliminate any air
space?
Was the lid sealed with electrical tape?
Are sample labels covered with clear tape?
Were each of the jars gently rotated and then placed in the dark?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Sediment Composition, Chemistry and Toxicity
Was the top of the sampler opened to determine whether the grab was
successful (does not have to be greater than 7 cm for these indicators)?
Was any overlying water drained off and large, non-living surface items
removed? Describe how this was done in the comments section.
Was any SAV removed after recording its presence on the field form?
Was the boat engine turned off or was the boat maneuvered to keep the engine
downwind?
Was a stainless steel or Teflon spoon washed with Alconox, rinsed with ambient
water, and used to collect sediment?
Was only the top two cm of sediment removed and used for sample?
Were any sediment used that was in direct contact with the sides of the
sampler? (Should not be used)
Was sediment placed in a stainless steel pot or bowl, and the pot placed on
wet ice? Was the container covered and in the cooler or on ice between grabs?
Was the process repeated until approximately 3 L of sediment (2 L in Great
Lakes) was collected? Identify the approximate number of grabs and total
amount of time required:
Number of Grabs: Time required:
Was the sediment stirred to sufficiently homogenize ALL sediment with the
spoon for approximately 10 minutes?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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Using the stainless steel spoon, was the 0.6 gallon bucket filled with sediment?
For marine sites preferred maximum volume = 1800 ml; min volume needed is
900 ml. For GL sites, preferred volume is 900 ml, minimum volume is 400 ml.
(Sediment toxicity)
Was the lid tightened to ensure a tight seal?
Was the bucket labeled, placed on wet ice and the sample id recorded on the
field form?
Using the stainless steel spoon, was 100 ml of sediment placed in the 120 ml
glass jar?
Was appropriate care taken to make sure the inside of the jar, cap, and
the sample was not contaminated?
Were the cap threads wiped off before the cap was screwed on the
bottle?
And was the bottle sealed with electrical tape applied in the clockwise
direction? (Sediment organics and metals)?
Was the 120 ml glass jar labeled, and placed on dry ice? Was the sample id
recorded on the field form?
Using the stainless steel spoon, was 50 ml of sediment placed in the 60 ml
glass jar?
Were the cap threads wiped off before the cap was screwed on the
bottle?
And was the bottle sealed with electrical tape applied in the clockwise
direction? (Sediment TOC)
Was the 60 ml jar labeled and placed on dry ice? Was the sample id recorded
on the field form?
Using the stainless steel spoon, was 100 ml of sediment placed in a labeled
zip-top bag, sealed and then double bagged in a second zip-top bag?
(Sediment grain size)
Was the bag labeled and placed on wet ice? Was the sample ID recorded on
the field form?
If insufficient sediment was collected for all sediment analyses, were the
sediments used in the priority order identified in the field operations manual
(section 5.3): 1) Organics/metals; 2) Toxicity; 3) TOC; 4) Grain size?
If insufficient sediment was collected for all sediment analyses, was this
flagged on the field forms and the pertinent 'no sample collected' bubbles
filled in?
Y
Y
V
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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SEDIMENT SAMPLING NOTES
Describe the tools used for transferring sample into sample containers and how the crew
ensured the tools were not contaminated from prior sampling sites.
Other comments:
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FISH TISSUE COLLECTION (Performed at Visit 1 only at Revisit sites)
Was a reasonable method for fish collection used and the method recorded on
the field form along with required gear specifics?
Were fish tissue collections attempted from within 500 meters of the X-site?
If no fish were collected within 500 meters of the X-site, did the crew attempt
fish collections from areas between 500 and 1000 meters from the X-site?
For human health fish tissue collections ONLY, if no suitable fish were
collected within 1000 meters of the X-site, did the crew attempt to collect the
HTIS sample between 1000 and 1500 meters from the X-site?
For each gear type used, were the pertinent GPS readings recorded?
Were clean nitrile gloves worn for handling fish?
Were crew members handling fish careful to avoid handling food, drink,
sunscreen and insect repellant prior to collecting fish?
Were potential target species/individuals rinsed in ambient water and placed
in live well?
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Eco Fish Tissue Collection
If no species on either the primary or secondary lists were available, did the
crew select an appropriate alternate species?
Was a target species with at least 5 fish of adequate size to provide a total
weight of 300 grams identified?
Did the field crew judge that all of the identified fish were of the same
species?
Were all of the fish in the sample at least 75% of the total length of the largest
fish? Provide length of longest fish, L*0.75, and length of smallest fish in
comments: Longest fish: Smallest fish:
Did the field crew report that all fish were collected at the same time (or no
more than one week apart?)
If fewer than 5 fish were collected, do they still meet the total weight and
other criteria?
If fewer than 5 fish were collected, did the crew spend at least 3 hours
attempting to collect fish?
Were the fish identified to species and the scientific name recorded on the Eco
Fish Collection Form?
Were total lengths measured in mm from the anterior most of the fish to the
top of the longest caudal fin ray (when the lobes of the caudal fin are
depressed dorsoventrally)?
Was the sample number, species retained, specimen lengths, location
collected, and sampling date/time recorded on the fish collection form?
Did the crew make sure that the sample ID recorded on the collection form
matches those on the sample labels?
If necessary, was each fish to be used as part of the sample dispatched with a
clean wooden bat (or equivalent wooden device)?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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Were all fish from the composite sample placed in a single 2-gallon self-sealing
bag? (or if they will not all fit in bag, in more than one)?
If spines that might puncture the bag exist, were they clipped/broken? Were
clipped spines placed inside the sample bag?
Did the crew prepare interior and exterior Sample Labels for the 2-gallon
bag(s) making sure that the label information matches the information on the
Fish Collection Form?
Was the interior label placed inside of a small (sandwich sized) self-sealing bag
and then placed inside the 2-gallon bag?
Was the exterior label affixed to the 2-gallon bag and covered with clear
plastic tape?
If needed, were labels with the same sample ID and information included with
additional bags?
Were all 2-gallon bags double-bagged together as one composite in a large
plastic bag?
Was the composite bag weighed to verify the minimum weight of 300 grams of
fish tissue was achieved?
Was a sample identification label prepared (making sure to include fish species
and max/min lengths and that the label information matches the Fish
Collection Form), affixed to a Tyvek tag, and covered with clear plastic tape?
Was a cable tie threaded through the grommet in the Tyvek tag and the outer
bag sealed with a cable tie?
Was the sample placed on dry ice or in a freezer immediately?
Or were they placed on wet ice for temporary holding and will be
frozen within 24 hours? (If "N" is circled, explain in comments)
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Fish Plug Collection
Were clean nitrile gloves worn?
If two individual fish were used, were they the same species?
If possible, were specimens selected from the Eco Fish collection?
Were plugs taken from specimens listed on the primary targeted fish list (or
secondary list of no primary species were available)?
If no species on either the primary or secondary lists were available, did the
crew select an alternate species using the following criteria: 1) those that are
consumed by humans; 2) predatory fish: and 3) other?
Was the smallest individual fish no smaller than 75% of the larger fish?
Was each specimen's total length and weight measured?
Were specimens rinsed in ambient water prior to plug removal?
Were two plugs taken, (typically one plug per/fish for a 2-fish composite)?
Was target weight of 0.5-0.7 grams collected from at least two plugs (i.e. the
equivalent of two full-depth plugs)?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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If the specimens were not part of the Eco Fish collection, was antibiotic spray
applied to the plug sites and the fish released?
N
N/A
FISH TISSUE COLLECTION NOTES
Human Health Fish Tissue Collection (Great Lakes Only)
If no species on either the primary or secondary target lists were available, did
the crew select an appropriate alternate species?
Was a target species with at least 5 fish of adequate size to provide a total of
500 grams of fillet tissue identified (fewer large fish are acceptable)?
Did the field crew judge that all of the identified fish were of the same
species?
Were all fish in the sample at least 75% of the total length of the largest fish?
Did the field crew report that all fish collected at the same time (or no more
than one week apart?)
If fewer than 5 fish were collected, do they still meet the total weight and
other criteria?
Were the fish identified to species and this information recorded on the Human
Health Fish Collection Form?
Were total lengths measured in mm from the anterior most of the fish to the
top of the longest caudal fin ray (when the lobes of the caudal fin are
depressed dorsoventrally)?
Was the sample number, species retained, specimen lengths, location
collected, and sampling date/time recorded on the Human Health Fish
Collection Form?
Did the crew make sure that the sample ID recorded on the collection form
matches those on the sample labels?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
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Was each fish to be used as part of the sample dispatched with a clean wooden
bat (or equivalent wooden device)?
N
N/A
Was each fish left intact and no fish plugs removed from any of the specimens?
N
N/A
Was each fish wrapped in extra heavy-duty aluminum foil (with dull side in)
(foil provided in fish tissue kit as solvent-rinsed, oven baked sheets)
Did the crew prepare a Sample Identification Label for each fish including
species and length?
Did the crew place each foil-wrapped fish individually in food grade tubing,
seal each end with a plastic cable tie, and attach an appropriate Sample Label
using clear tape and wrapping the tape completely around the wrapped fish so
the tape wraps around itself?
Did the crew double-bag the entire set of specimens in the composite inside a
large plastic bag?
Was an outer bag sample label prepared (making sure to include fish species
and individual lengths and that the label information matches the Human
Health Fish Collection Form), affixed to a Tyvek tag, and covered with clear
plastic tape?
Was a cable tie threaded through the grommet in the Tyvek tag and the outer
bag sealed with a cable tie?
Was the sample placed on dry ice or in a freezer immediately? Y N N/A
Will the samples either be shipped on at least 50 pounds of dry ice or
placed in a freezer within 24 hours? Y N N/A
Are the fish layered with the dry ice, starting and ending with dry ice?
(If "N" is circled, explain in comments) Y N N/A
HUMAN HEALTH FISH TISSUE COLLECTION NOTES
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
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Field Operations Manual
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ENTEROCOCCI FILTERING
Processing the Fecal Indicator Filter Blank
Is the site a Revisit site? If yes, filter blanks should be prepared and the
following questions should be answered. If no, skip to next section.
Was the filter blank prepared before the filtration of the Enterococci sample?
Were the microcentrifuge tubes with beads chilled on dry ice?
(enough for both the blanks and the samples?)
Were clean gloves worn?
Was the sterile phosphate buffer chilled on wet ice?
Was a new sterile 100 ml filter funnel with pre-loaded filter used??
Was the filter funnel attached to the side arm filter flask using the correct
rubber stopper (white) and adapter? (or crew supplied manifold system was
assembled appropriately) See figure 14.2.
Was 20 ml of the sterile phosphate buffer measured in the sterile 50 ml
graduated centrifuge tube and poured into the filter funnel?
Was a hand or electric vacuum pump attached to the filtering apparatus
Circle which was used.
Was care taken to ensure the vacuum pressure did not exceed 7 inches of
mercury -3.4 psig?
Was it pumped until all liquid was in the filtrate collection flask/ reservoir?
Was the top of the filter funnel removed from the base without disturbing
filter?
Were sterile disposable forceps used to remove the filter (touching only the
filter edges)?
Was the filter folded it in half, in quarters, eighths, and then in sixteenths
(filter is folded 4 times)?
Was the filter inserted into chilled filter extraction tube (with beads) point
side up?
Was the screw cap replaced and tightened?
Was the volume of buffer filtered through the filter recorded on the filter
blank sample label?
Was the label attached to the microcentrifuge tube and not on cap?
Was any tape applied to the cap or elsewhere on the microcentrifuge tube
(SHOULD NOT BE)?
Was the tube(s) inserted into bubble wrap bag on dry ice for preservation
during transport and shipping?
Did the crew mark the "Blank Collected" bubble on the Sample Collection Form?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
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N/A
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N/A
N/A
N/A
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N/A
N/A
N/A
N/A
N/A
N/A
Processing the Fecal Indicator Sample
Were nitrile gloves worn?
Y
N
N/A
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Field Operations Manual
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Were the microcentrifuge tubes with beads chilled on dry ice?
Was the Enterococci sample chilled for at least 15 minutes before filtering?
Was the sterile phosphate buffer chilled on wet ice?
Was a new sterile 100 ml filter funnel with pre-loaded filter used?
Was the filter funnel attached to the side arm filter flask using the correct
rubber stopper (white) and adapter? (or crew supplied manifold system was
assembled appropriately) See figure 14.2.
Was the sample bottle shaken 25 times to mix well?
Was the 25 ml of the mixed water sample measured in the sterile graduated 50
ml centrifuge tube and poured into the filter funnel?
Was a hand or electric vacuum pump attached to the filtering apparatus
Circle which was used.
Was care taken to ensure the vacuum pressure did not exceed 7 inches of
mercury -3.4 psig?
Was it pumped until all liquid was in the filtrate collection flask?
If the first 25 ml volume passed readily through the filter, was another 25 ml
measured and added and the filtration continued?
If the filter clogged before completely filtering the first or second 25 ml
volume, was the filter discarded and the filtration repeated using a lesser
volume and a new sterile filter funnel?
Was approx. 10 ml of the chilled sterile phosphate buffer poured into the
graduated 50 ml centrifuge tube used for the sample?
Was the tube capped and shaken 5 times?
Was the cap removed and the rinsate poured into the filter funnel to rinse
filter?
Was the rinsate filtered and repeated with another 10 ml of sterile buffer?
Was the top of the filter funnel removed from the base without disturbing
filter?
Were sterile disposable forceps used to remove the filter (touching only the
filter edges)?
Was the filter folded it in half, in quarters, eighths, and then in sixteenths
(filter is folded 4 times)?
Was the filter inserted into chilled filter extraction tube (with beads) point
side up?
Was the screw cap replaced and tightened?
Was the volume of water sample filtered through the filter recorded on the
sample label?
Was the label attached to the microcentrifuge tube?
Was any tape applied to the cap or elsewhere on the microcentrifuge tube
(SHOULD NOT BE)?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
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N/A
N/A
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N/A
N/A
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Was the tube inserted into bubble wrap bag on dry ice for preservation during
transport and shipping?
N
N/A
Was the volume of the buffer rinse used for the filter recorded on the sample
collection form?
N
N/A
Was the filtration start time and finish time as well as the time frozen
recorded for on the sample collection form?
N
N/A
Were the steps repeated for the other 50 ml sub-sample volume to be filtered?
NOTE: A new sterile filter funnel with pre-loaded filter is used for each filter.
N
N/A
Was aseptic technique used to store the forceps between filter runs?
N
N/A
Were the volumes the same for each of the 2 filters?
N
N/A
ENTEROCOCCI FILTERING NOTES
CHLOROPHYLL-^ AND DISSOLVED NUTRIENTS SAMPLE
Filtered Nutrients collection device
Did the crew use a nutrients chamber that allows collection of the dissolved
nutrients sample directly into the sample bottle?
// no, continue to steps below, if yes, skip to next section.
Did the crew use a filtering flask that was labeled for CHLA/NUTS only and is
DIFFERENT than the flask used for Enterococci?
Was the flask thoroughly cleaned and rinsed with Dl water prior to use for
collecting nutrients sample (i.e. between sites)?
Y
Y
Y
N
N
N
N/A
N/A
Processing the CHLA/NUTS Sample
Did the crew use a new sterile blue-bottom filter funnel (as opposed to reusing
a filter funnel that was previously used at a different site)? MUST USE NEW
Y
N
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Version 1.0 May 2015
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Were Nitrile gloves worn?
Was the filter funnel attached to the chamber or side arm filter flask using the
correct rubber stopper (blue) and adapter? See figure 14.3.
Was the originally loaded filter (patterned) removed from the base, but the
filter pad left in place?
Was a glass fiber filter placed in the filter funnel, pressed (gridded) side down
(i.e. rough side up)?
Was the filter handled with clean forceps?
Was 250 ml of site water measured with a clean graduated cylinder and
poured into the filter funnel, the cap replaced, and the sample pumped
through the filter?
Was a hand or electric vacuum pump attached to the filtering apparatus
Circle which was used.
Was care taken to ensure the vacuum pressure did not exceed 7 inches of
mercury (~ 3.4 psig) and that no single sample volume was filtered for longer
than 5 minutes?
// a filter flask was used to collect filtrate, was 10-20 ml of filtrate used to
rinse the filter flask and then discarded and was this process performed a total
of 3 times?
Was 10-20 ml of filtrate used to rinse the sample bottle and then discarded
and was this process performed a total of 3 times?
If 250 ml of water did not pass through the filter, was the filter changed, the
apparatus rinsed with Dl water, and the procedures (including rinses) repeated
using 100 ml of site water?
// a filter flask was used to collect filtrate, was 250 ml of filtrate poured into
a 250 ml HOPE bottle? (if no flask used, was 250 ml of filtered water
collected directly into the sample bottle)?
Was the dissolved nutrients label affixed to the bottle and then covered with
clear plastic tape?
Was the sample information recorded on the sample collection form and the
sample placed on wet ice?
Was the filter observed for visible color?
If there was no visible color, did the process proceed until color was visible on
the filter or until a maximum of 2,000 ml was filtered?
NOTE, if the crew is using a filter chamber, they should switch to a flask or
manifold setup once the nutrients sample is collected.
Was the level of water monitored in the lower chamber to ensure that it did
not contact the filter or flow into the pump?
After readily visible color was seen on the filter, was the actual sample volume
filtered recorded on the Sample Collection Form and on the CHLA sample
label?
Was the graduated cylinder and the upper portion of the filter funnel rinsed
thoroughly with Dl water to include any remaining cells adhering to the sides
and pumped through the filter?
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
N
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N/A
N/A
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Was the filter removed from the holder with clean forceps?
Was the filter folded in half, with the colored side folded inward?
Was the folded filter placed into a 50 ml centrifuge tube and capped?
Was the cap sealed tightly by turning an additional 14 turn past the point at
which initial resistance is met and then taped with electrical tape?
Was the sample volume filtered recorded on a chlorophyll label and attached
to the centrifuge tube?
Was the label covered with a strip of clear tape?
Does the "total volume of water filtered" on the Sample Collection Form
match the total volume recorded on the sample label?
Was the 50-mL centrifuge tube wrapped in aluminum foil and placed in a self-
sealing plastic bag?
Was this bag placed on dry ice?
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
CHLOROPHYLL-^ AND DISSOLVED NUTRIENTS SAMPLE NOTES
FINAL SITE ACTIVITIES
General site Assessment
Were any of the shoreline activities and disturbances recorded that were
observed on the shoreline adjacent to the sampling site visible from the X-site?
For shoreline activities and disturbances that the crew observed, was the
rating of the abundance or influence marked as low (L), moderate (M), or
heavy (H) on the line next to the listed disturbance?
If shoreline activities were not noted, did the crew leave the bubbles blank?
Y
Y
Y
N
N
N
N/A
N/A
N/A
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Were observations regarding the general characteristics of the site recorded on
the Site Assessment Form (in a 200 m circle around the X-site) using the scale
of 1 -5?
Did all field crew members contribute to the evaluation?
Were other items such as signs of pipe outflows, extreme weather, etc.
recorded?
Was the comments section used on the Site Assessment Form to note any other
pertinent information about the site?
Y
Y
Y
Y
N
N
N
N
N/A
N/A
N/A
N/A
Data Forms and Sample Inspection
After the Site Assessment Form was completed, did the Field Crew Leader
review all of the data forms and sample labels for accuracy, completeness, and
legibility?
Did the other crew member(s) inspect all sample containers and packages in
preparation for transport, storage, or shipment?
Did the crew ensure that all required data forms for the site were completed?
Did the crew confirm that the SITE-ID and date of visit are correct on all forms?
On each form, did the crew verify that all information was recorded
accurately, the recorded information was legible, and any flags were explained
in the comments section?
Did the crew ensure that comments were clear and legible, with no "shorthand"
or abbreviations?
After reviewing each form (if using paper forms), was the upper right corner of
each page of the form initialed?
Did the crew ensure that all samples were labeled, all labels are completely
filled in, and each label was covered with clear plastic tape?
If any samples were not collected, was the pertinent "no sample collected"
bubble(s) filled in on the data form(s)
Were all sample containers checked to ensure that they were properly sealed?
Will the coolers be shipped with fresh bags of ice in cooler?
Verify that the coolers will be shipped by overnight courier ASAP after
collection (e.g. the same or next day). Identify shipping date: / 72015
If samples will be held after collection, will they be kept cold and in darkness?
Identify where they will be stored:
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
N
N
N
N
N
N
N
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Launch Site Cleanup
Were the boat, motor, and trailer inspected for evidence of weeds and other
macrophytes?
Were the boat, motor, and trailer cleaned as completely as possible before
leaving the launch site?
Were all nets/sieves etc. inspected for pieces of macrophyte or other
organisms and as much as possible was removed before packing for transport?
Were all equipment and supplies packed in the vehicle and trailer for transport
and kept organized as presented in the equipment checklists?
Y
Y
Y
Y
N
N
N
N
N/A
N/A
N/A
N/A
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Was all waste material at the launch site cleaned up and disposed of or
transported out of the site if a trash can is not available?
N
N/A
Were equipment needs identified and those needs will be conveyed to the FLC
or Requested via the Request Form?
N
N/A
Miscellaneous
Do the crew members know the phone numbers for pertinent points of
communication (FLC, IMTeam, RMC, and/or HQ), is the number saved in cell
phone, or do they know the location of numbers in Field Ops Manual?
N
N/A
Do the crew members have suggestions/problems concerning the sampling
Procedures, forms, lodging, logistics, etc.?
N
N/A
FINAL SITE ACTIVITIES NOTES
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FINAL EVALUATION ACTIVITIES
Areas of Strength
Areas of Concern
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FINAL EVALUATION ACTIVITIES
Was the crew debriefed on the results of the evaluation by the evaluator?
N
N/A
COMMENTS OF THE CREW BEING EVALUATED
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SIGNATURES
Evaluator Date Field Crew Leader Date
Field QC Officer (if assigned by site) Date Field Crew Member Date
Field Crew Member Date Field Crew Member Date
Field Crew Member Date Field Crew Member Date
Field Crew Member Date Field Crew Member Date
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