United States Environmental Protection Agency
Office of Water
Washington, DC
EPA# 841 -F-19-005
National Coastal Condition Assessment
2020
Field Operations Manual
April 2020
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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 2020: Quality Assurance Project Plan
(EPA # 841-F-19-003)
• National Coastal Condition Assessment 2020: Site Evaluation Guidelines (EPA #
841-B-20-001)
• National Coastal Condition Assessment 2020: Field Operations Manual (EPA #
841-F-19-005)
• National Coastal Condition Assessment 2020: Laboratory Operations Manual
(EPA # 841-F-19-004)
This Field Operations Manual contains a brief introduction and base and site location
procedures for in situ measurements, sampling water (grabs for chemistry, pathogen analysis,
and algal toxin analysis), benthic macroinvertebrates, sediment (for composition,
contamination and toxicity), and fish tissue (for human health and ecological indicators).
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:
U.S. EPA. National Coastal Condition Assessment 2020 Field Operations Manual. United States
Environmental Protection Agency, Office of Water, Office of Wetlands, Oceans and
Watersheds. Washington, D.C. EPA- 841-F-19-005. 2020.
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Version History
FOM Version
Date
Changes Made
1.0
April 10, 2020
Not Applicable
1.1
April 23, 2020
Change only to update
Version to 1.1. and maintain
version alignment with QAPP
VI.1 and LOM VI.1
1.2
February 18, 2021
Section 6.4, step 3, added text
to take another reading at
0.5m from bottom as you
start upcast. This is not a
change in procedure, just a
clarification.
1.2
February 18, 2021
Section 6.4, step 4, added text
that crews should sample at
least to 30m if limited by
equipment.
1.2
February 18, 2021
Section 13.4, step 2, added
text that crews need to record
grab number in the App.
1.2
March 1, 2021
Insertion of Section 2.8.5 to
reflect COVID-19 safety
considerations.
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1 Table of Contents
1 TABLE OF CONTENTS Ill
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 5
2.3.1 Site Sample-ability 5
2.3.2 Replacing Sites 6
2.4 Description of NCCA Indicators 6
2.4.1 In Situ Water Column Measurements 6
2.4.2 Water Chemistry (CHEM) and Associated Measurements 7
2.4.3 Algal Toxins (Cylindrospermopsin and Microcystins [MICX] and Microcystins [MICZ]) 7
2.4.4 Underwater Video (UVID) 7
2.4.5 Sediment Assessment, (SEDG, SEDC, SEDX, SEDO, and D15N) 8
2.4.6 Benthic Macroinvertebrate Assemblage (BENT) 8
2.4.7 Enterococci Fecal Indicator (ENTE) 8
2.4.8 Fish Tissue (FTIS, FPLG, HTIS) 8
2.4.9 Ocean and Coastal Acidification Research Indicator 9
2.5 Supplemental AAaterial to the Field Operations AAanual 9
2.6 Recording Data and Other Information 10
2.6.1 Electronic Field Forms 10
2.6.2 Paper Field & Tracking Forms 10
2.7 Data Management 12
2.8 Safety and Health 12
2.8.1 General Considerations 12
2.8.2 Safety Equipment 13
2.8.3 Safety Guidelines for Field Operations 13
2.8.4 General Safety Guidelines for Field Operations 14
2.8.5 Covid-19 Safety Considerations 15
3 INTRODUCTION TO SAMPLING 16
3.1 Site Visit Duration 16
3.2 Field Crew Makeup 16
3.3 Sampl i ng Sequence 16
3.4 Sampling Considerations 16
3.4.1 Considerations for Fish Tissue Collection 16
3.4.2 Listed Species Considerations 17
3.4.3 Considerations for Enterococci Collection 17
3.4.4 Other Considerations 18
A PRE-DEPARTURE ACTIVITIES 23
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4.1 Daily Itineraries 24
4.2 Instrument Checks and Calibration 24
4.2.1 Initial Assembly and Setup Procedures for LI-COR frame, sensor and Datalogger 25
4.3 Equipment and Supply Preparation 27
5 INITIAL SITE PROCEDURES 29
5.1 Site Verification 29
5.1.1 Equipment & Supplies 29
5.1.2 Site Verification Procedures 29
5.1.3 Site Relocation 30
5.1.4 Site Characteristics 30
5.2 Site Photograph 31
5.3 Sample Collection 31
5.4 Secondary Sediment or Fish Collection Zones 32
5.4.1 Sediment Samples 33
5.4.2 Fish Samples 34
6 WATER QUALITY MEASUREMENTS 36
6.1 Summary of Method for In Situ Measurements of Water Column Transparency, Dissolved Oxygen, pH,
Salinity, Conductivity, Temperature, and Light Attenuation 36
6.1.1 Equipment and Supplies 36
6.2 Sampling Procedure - Water Column Transparency (Secchi Depth) 36
6.3 Sampling Procedure - Multi-Parameter Sonde 37
6.3.1 Calibration 37
6.3.2 Dissolved Oxygen Meter 39
6.3.3 pH Meter 39
6.3.4 Salinity/Conductivity Meter 40
6.3.5 Temperature Meter 40
6.4 Sampling Procedure - Dissolved Oxygen, pH, Temperature and Salinity/ Conductivity 40
6.5 Photosynthetically Active Radiation (PAR) Meter 41
6.5.1 Sampling Procedure—Light Attenuation (LI-1400 Datalogger) 42
7 TOTAL ALKALINITY [ALKT] 44
7.1 Summary of Method 44
7.2 Equipment and Supplies 44
7.3 Sampling Procedure 44
7.3.1 Sample Collection 45
8 WATER CHEMISTRY [CHEM], CHLOROPHYLL-^ [WCHL], AND NUTRIENTS [NUTS] SAMPLE
COLLECTION AND PRESERVATION 48
8.1 Summary of Method 48
8.2 Equipment and Supplies 48
8.3 Sampling Procedure 48
9 ALGAL TOXINS (CYLINDROSPERMOPSIN [MICX] AND MICROCYSTES [MICZ]) 50
9.1 Summary of Method 50
9.2 Equipment and Supplies 50
9.3 Sampling Procedure 51
9.3.1 Sample Collection 51
9.3.2 Sample Storage 51
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10 FECAL INDICATOR (ENTEROCOCCI, [ENTE]) 52
10.1 Summary of Method 52
10.2 Equipment and Supplies 52
10.3 Sampling Procedure 52
11 PHYTOPLANKTON [PHYT] (GREAT LAKES ONLY) 54
11.1 Summary of Method 54
11.2 Equipment and Supplies 54
11.3 Sampling Procedure 54
12 UNDERWATER VIDEO [UVID] (GREAT LAKES ONLY) 56
12.1 Summary of Method 56
12.2 Equipment and Supplies 57
12.3 Underwater Video Carriage Set-up 57
12.3.1 Setting up Video Carriage System 57
12.3.2 Operating Camera and Lights 61
12.4 Underwater Video Collection 62
12.4.1 Summary of Method 62
12.4.2 Deploying GoPro video carriage 64
12.4.3 Transferring and Backing up Video Files 66
13 SEDIMENT COLLECTION 69
13.1 Summary of Method 69
13.2 Equipment and Supplies 70
13.3 Sampling Procedure 70
13.4 Processing Procedure - Benthic AAacroinvertebrate [BENT] Composition And Abundance 72
13.5 Processing Procedure - Sediment Composition, Chemistry, Toxicity AND Nitrogen Isotopes 74
14 FISH TISSUE COLLECTION 78
14.1 Ecological Contamination Fish Tissue Collection [FTIS] 79
14.1.1 Summary of Method 79
14.1.2 Equipment and Supplies 81
14.1.3 Sample Storage and Shipping Preparation 83
14.2 Fish Tissue Plug [FPLG] 88
14.2.1 Summary of Method 88
14.2.2 Equipment and Supplies 89
14.2.3 Sampling Procedure 89
14.2.4 Sample Storage 91
14.3 Human Health Fish Tissue Collection [HTIS] (Great Lakes Nearshoreand Lake Michigan Enhancement
SITES ONLY) 94
14.3.1 Summary of Method 94
14.3.2 Fish Tissue Distribution Scheme 95
14.3.3 Equipment and Supplies 96
14.3.4 Sampling Procedure 97
14.3.5 Sample Storage and Shipping Preparation 98
15 FINAL SITE ACTIVITIES 101
15.1 General Site Assessment 102
15.1.1 Shoreline Activities and Disturbances 102
15.1.2 Site Characteristics 102
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15.1.3 General Assessment 102
15.2 Processing The Fecal Indicator 103
15.2.1 Summary of Method 103
15.2.2 Equipment and Supplies 103
15.2.3 Processing Procedure - Fecal Indicator Filter Blank 103
15.2.4 Processing Procedure - Fecal Indicator Sample 104
15.3 Processing The Chlorophyll-4 & Dissolved Nutrients Indicators 106
15.3.1 Summary of Method 106
15.3.2 Equipment and Supplies 107
15.3.3 Processing Procedure 107
15.4 Post-Measurement Calibration Check of Multi-Parameter Sonde 109
15.5 Field Data & Tracking Form Review 110
15.6 Sample Packaging and Label Review 110
15.7 Sample Shipment & Tracking Form Submittal 111
15.7.1 Time-Sensitive Samples 111
15.7.2 Other Samples 112
15.8 Equipment Cleanup & Check 112
15.8.1 Boat & Trailer Cleanup 113
15.8.2 Post Sampling Equipment Care 113
15.8.3 Additional Decontamination Information 113
16 POST-SAMPLING ACTIVITIES 115
16.1 Sample Shipping 115
16.2 Data Submittal 115
16.3 Data and Tracking Reminders 116
16.4 Site Evaluation Spreadsheet Submittal 116
17 FIELD QUALITY CONTROL 117
17.1 Standardized Training 117
17.2 Standardized Field Data Collection App 117
17.3 Repeat Sampling 117
17.4 Field Evaluation And Assistance Visits 118
17.4.1 Specifications for QC Assurance 119
17.4.2 Reporting 119
18 LITERATURE CITED 121
APPENDIX A: EQUIPMENT AND SUPPLIES LISTS 123
Base Kit 123
Site Kits 125
Marine Site Kit 125
Great Lakes Site Kit 126
Human Health Fish Tissue Sampling Kit 127
Crew Supplied Equipment 127
APPENDIX B: SAMPLE LABELS a PACKING SLIPS 130
Sample Labels (Marine) 130
Sample Labels (Great Lakes) 132
APPENDIX C: SHIPPING AND TRACKING GUIDELINES 135
Tracking Forms in the App 135
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T1 - Daily Water Chemistry Samples 136
77 - Chilled Batched Samples (Marine = Daily, Great Lakes = Weekly) 136
T3 - Frozen Batched Samples 137
T4 - Non-Chilled: Batched Samples 137
T5 - Eco Fish Tissue 138
T6 - Human Health Whole Fish Tissue Composite Sample - NGL20, ISA20, and NPA20 sites Only 138
T7 - Underwater Video UVID Form [Great Lakes Only] 139
Shipping Guidelines 139
APPENDIX D: MICROPLASTICS COLLECTION AND PROCESSING 148
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List of Tables
Table 1.1 Contacts xii
Table 1.2 Regional Coordinators xm
Table 2.1 Guidelines for recording field measurements & tracking inforaaation 11
Table 2.2 General health & safety considerations 12
Table 4.1 Stock solutions, uses & methods for preparation 28
Table 5.1 Equipment & supplies: site verification 29
Table 6.1 Equipment & supplies: transparency, DO, pH, salinity/conductivity, temperature, & light
ATTENUATION 36
Table 6.2 Example depth measurement intervals 41
Table 7.1 Equipment & supplies: total alkalinity sample collection 44
Table 8.1 Equipment & supplies: water chemistry & chlorophyll-a sample collection 48
Table 9.1 Equipment & supplies: algal toxins (cylindrospermopsin and microcystins) 50
Table 10.1 Equipment & supplies: fecal indicator (Enterococci) sampling 52
Table 11.1 Equipment & supplies: phytoplankton 54
Table 12.1 Equipment & supplies: underwater video 57
Table 12.2 GoPro Hero 7 camera and light settings and directions 62
Table 13.1 Equipment & supplies: sediment collection 70
Table 14.1 Equipment & supplies: eco fish tissue collection 81
Table 14.2 Northeast region primary and secondary marine target species - whole body fish tissue collection
(Ecofish) 84
Table 14.3 Southeast region primary and secondary aaarine target species - whole body fish tissue collection
(Ecofish) 84
Table 14.4 Gulf region priaaary and secondary aaarine target species - whole body fish tissue collection
(Ecofish) 85
Table 14.5 Western region priaaary and secondary aaarine target species - whole body fish tissue collection
(Ecofish) 86
Table 14.6 Great Lakes priaaary and secondary target species - whole body fish tissue collection (Ecofish) ... 87
Table 14.7 Equipment & supplies: fish tissue plugs 89
Table 14.8 Priaaary and secondary aaarine target species for fish plug collection 92
Table 14.9 Priaaary and secondary Great Lakes target species for fish plug collection 93
Table 14.10 Equipment & supplies: huaaan health fish tissue collection 96
Table 14.11 Priaaary and secondary Great Lakes target species for huaaan health fish tissue collection 99
Table 15.1 Equipment & supplies: Enterococci processing 103
Table 15.2 Equipment & supplies: chlorophyll-a & dissolved nutrients processing 107
Table 15.3 General cleaning of sampling gear after each site 112
Table 17.1 General inforaaation noted during field evaluation 119
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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 intra-coastal estuarine system 4
Figure 2.3 Hypothetical Great Lakes Nearshore target population 5
Figure 3.1 Marine Field Sampling Scenario - Active Fishing Methods 19
Figure 3.2 Marine Field Sampling Scenario - Passive Fishing Methods or Ability to Filter Samples Immediately .20
Figure 3.3 Great Lakes Field Sampling Scenario - Active Fishing Methods 21
Figure 3.4 Great Lakes Field Sampling Scenario - Passive Fishing Methods or Ability to Filter Samples
Immediately 22
Figure 4.1 Overview of base site activities 23
Figure 4.2 Attachment of the underwater sensor to the mounting rings (adapted from LI-COR, 2006) 26
Figure 4.3 Lowering frame assembly with sensor, weight, and lowering line (adapted from LI-COR, 2006) ....26
Figure 5.1 Primary, secondary and tertiary sample collection zones 34
Figure 5.2 Primary and secondary fish collection zones 35
Figure 7.1 Target fill range for total alkalinity sample bottles 46
Figure 7.2 Total alkalinity filter detail 47
Figure 12.1 Underwater video assembly 56
Figure 12.2. GoPro camera A and light mounted on carriage 59
Figure 12.3. Approximate aiming angle of camera and lights for the oblique (B) and down-looking (A) cameras.
60
Figure 12.4. GoPro camera B and light mounted on carriage 60
Figure 12.5. Manufacturer's instructions for lights 61
Figure 12.6 Example of verification form in the NCCA App showing header information prior to underwater
CAMERA DEPLOYMENT 63
Figure 12.7 Example of sample collection form in the NCCA App showing header and benthic grab information
PRIOR TO UNDERWATER CAMERA DEPLOYMENT 64
Figure 12.8 Example of sample collection form in the NCCA App, UVID inforaaation 68
Figure 13.1 Illustration of acceptable & unacceptable grabs for benthic community analysis 72
Figure 14.1 Fish Tissue Distribution Scheme to be used at all Great Lake Sites with the prefix NGL20, ISA20, or
NPA20 96
Figure 15.1 Final site activities summary 101
Figure 15.2 Filtering set-up for Enterococci filtering 106
Figure 15.3 Filtering set-up for chlorophyll-a and nutrients filtering 109
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Acronyms/Abbreviations
CPR Cardiopulmonary resuscitation
Dl Deionized
DO Dissolved oxygen
EPA Environmental Protection Agency
ESA Endangered Species Act
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
HDPE High density polyethylene
HQ Headquarters
IM Information Management
MED Mid-Continent Ecology Division, U.S. EPA Office of Research and Development
NAD 83 North American Datum of 1983
NARS National Aquatic Resource Surveys
NCA National Coastal Assessment
NCCA National Coastal Condition Assessment
NEP National Estuary Program
NIST National Institute of Standards and Technology
NM Nautical miles
NOAA National Oceanic and Atmospheric Administration
ORD Office of Research and Development, U.S. EPA
OSHA Occupational Safety and Health Administration
PAR Photosynthetically active radiation
PBS Phosphate Buffer Solution
PDF Portable Document Format
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PET polyethylene terephthalate
PETG polyethylene terephthalate copolyester, glycol modified
PFD Personal flotation device
PSI Pounds per square inch
QAC Quality Assurance Coordinator
QAPP Quality Assurance Project Plan
QA/QC Quality assurance/quality control
QCS Quality Check Solution
SAV Submerged aquatic vegetation
SOP Standard Operating Procedure
SRM Standard Reference Material
TOC Total organic carbon
USDA United States Department of Agriculture
USGS United States Geological Survey
VHS Viral Hemorrhagic Septicemia
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Contact List
Table 1.1 Contacts
Role
Name
Phone/Email
Address
NCCA Team
Hugh Sullivan
202-564-1763
US EPA Office of Water
Lead
sullivan.huch@epa.p-ov
1200 Pennsylvania Avenue NE
(45031)
Washington DC 20460
NCCA QA
Danielle
202-566-2876
US EPA Office of Water
Coordinator
Grunzke
prunzke.danielle@epa.piov
1200 Pennsylvania Ave NE
(4503T)
Washington DC 20460
NARS QA
Kendra Forde
202-566-0417
US EPA Office of Water
Coordinator
forde.kendra@epa.pov
1200 Pennsylvania Ave NE
(45031)
Washington DC 20460
EPA Logistics
Brian Hasty
202-564-2236
US EPA Office of Water
Coordinator
hastv.brian@epa.pov
1200 Pennsylvania Ave NE
(45031)
Washington DC 20460
Contractor Field
Chris Turner
715-829-3737
Great Lakes Environmental
Logistics
cturner@plec.com
Center, Inc.
Coordinator
739 Hastings Street
Iraverse City, MI 49686
NARS IM
Michelle Gover
541-754-4793
GDI1
Coordinator
pover.michelle@epa.pov
200 SW 35th Street
Corvallis OR 97333
Great Lakes
Le anne Stahl
202-566-0404
US EPA Office of Water
Human Health
stahl.leanne@epa.pov
1200 Pennsylvania Avenue NE
Fish Tissue
John Healey
202-566-0176
(43051)
Manager
(Alternate)
he ale v. j ohn @ep a. po v
Washington DC 20460
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Table 1.2 Regional Coordinators
Role Name Phone/Email Address
EPA Region 1
Hilary Snook
617-918-8670
snook.hilarv@epa.p-ov
USEPA Region 1 — New
England Regional Laboratory
11 Technology Drive
North Chelmsford, MA
01863-2431
EPA Region 2
Emily Neriiig
732-321-6764
nerinp-.emilv@epa.p-ov
USEPA Facilities
Raritan Depot
2890 Woodbridge Avenue
Edison, NJ 08837-3679
EPA Region 3
Bill Richardson
215-814-5675
richardson.william@epa.pov
USEPA Region 3
1650 Arch Street
Philadelphia, PA 19103-2029
EPA Region 4
Chris McArthur
404-562-9391
mca rthur. chris tophe r@ep a. po v
USEPA Region 4
15165 — Sam Nunn Atlanta
Federal Center.
Atlanta, GA 30303-8960
EPA Region 5
Mari Nord
Ed Hammer
312-886-3017. nord.mari@epa.pov
312-886-3019. hammer.edward@epa.pov
USEPA Region 5
77 West Jackson Boulevard
Chicago," IL 60604-3507
EPA Region 6
Rob Cook
214-665-7141
cook.robert@epa.pov
USEPA Region 6
1445 Ross Avenue
Suite 1200
Dallas, TX 75202-2733
EPA Region 9
Matt Bolt
415-972-3578
bolt.matthew@epa.pov
USEPA Region 9
75 Hawthorne Street
San Francisco, CA 94105
EPA Region
10
Lillian Herger
206-553-1074
herper.lillian@epa.pov
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 estuan'ne 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 2005; 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 2020 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 2005 and 2020.
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With input from the states and other partners, EPA used an unequal probability, stratified
design to select 1000 probabilistic sampling events, of which roughly 50% are resample
sites (sites that were sampled in 2010 or 2015 and will be sampled again in 2020).
Resample sites from 2010/2015 are identified as Base 10 sites; while newly drawn sites
are identified as Base 20 sites. Approximately 7% of the 2010/2015 resample sites are also
designated "revisit sites," which indicates that they will be sampled twice in 2020 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 2020, 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
probabilistic 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.
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, Figure 2.2 and Figure 2.3 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 Great Lakes target population excludes embayments with connection to open
water that are less than 200 meters in width. 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 NCA developed by EPA Office of Research and
Development (ORD) Gulf Ecology Division (GED). The prior GED sample frame was
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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 the first NCCA in 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 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 2020 sample frame. For the Californian Province excluding San
Francisco Bay, the GED sample frame was changed to match a 2004 sample frame used for
the 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 was necessary to
provide the information required to estimate change between the periods of 2010 and
2015. The sample frame for the Great Lakes sites was obtained from EPA ORD Mid-
Continent Ecology Division (MED).
Please refer to the NCCA 2020: Site Evaluation Guidelines for more detailed information
on the target population and exclusion criteria.
Figure 2.1 Example of an estuarine system comprised of an embayment plus a
complex of bays and tidal rivers and creeks
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Figure 2.2 Example of an intra-coastal estuarine system
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Outside
Figure 2.3 Hypothetical Great Lakes Nearshore target population.
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
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
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the appropriate "Non-Sampleable-Permanent" category on the Verification Form in the
NCCA App. 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/organization. A
site can be deemed unsampleable for any number of reasons, including being too shallow
to properly operate sampling equipment or in the middle of a navigational channel where
it is unsafe.
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 20 sites must be replaced with Base 20
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/organization. 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 updated 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),
then the replacement site takes on the RVT2 assignment. That is the replacement site
must be visited twice in 2020.
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 mark that indicator as not collected and document the reason why
the indicator could not be collected in the comment area of the NCCA App. See Section
13 and Section 14 for information regarding the collection of sediment and fish samples,
the two indicators which crews most likely may experience difficulty collecting.
2.4 Description of NCCA Indicators
Indicators for the 2020 survey will basically remain the same as those used in 2015 and
other past coastal surveys, with a few modifications. Additionally, 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
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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 SecchiDisk 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 (WCHL)
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 Toxins (Cylindrospermopsin and Microcystins [MICX] and Microcystins
[MICZ])
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
sample is taken to analyze for both cylindrospermopsin and microcystins, 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 array 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. If the benthic macroinvertebrate sample
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is collected at a location other than the Y-Location, a second video focused on the
substrate at the benthos collection location will be taken.
2.4.5 Sediment Assessment, (SEDG, SEDC, SEDX, SEDO, and D15N)
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. The nitrogen
stable isotope ratio [D15N] in sediment will be measured to evaluate its utility as a
measure of anthropogenic development in watersheds of estuaries and will also be
collected at marine sites only.
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 and
is also referred to as the ecofish sample, 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. At revisit
sites, ecofish samples will only be targeted during visit 1. If a successful collection is not
possible at visit 1, crews should attempt to collect ecofish at visit 2.
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
they were collected. At revisit sites, fish plug samples will only be targeted during visit 1.
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If a successful collection is not possible at visit 1, crews should attempt to collect fish
plugs at visit 2.
In the Great Lakes only, additional fish composite samples will be collected at all of the
225 probabilistic nearshore Great Lakes sites (prefix = NGL20), all 38 Great Lakes island
sites (prefix = ISA20), and all 12 Great Lakes park sites (prefix = NPA20) for a combined
total of 275 sites. Fillet tissue from these samples will be homogenized in the lab 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 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, but HTIS will only be collected at one of the two
visits to a revisit site. Note that human health fish tissue samples will NOT be collected at
Great Lakes enhancement sites other than those listed above.
2.4.9 Ocean and Coastal Acidification Research Indicator
2.4.9.1 Total Alkalinity (ALKT)
At marine sites only, crews will collect a water sample in two bottles for the
measurement of total alkalinity. Total alkalinity (TA) is a characteristic of seawater that,
in combination with other measurements, can be used to calculate total pH (i.e., coastal
acidification) and the availability of carbonate ions used by marine organisms to produce
structural materials such as corals and shells. TA is also used to calculate the fate of
carbon that enters coastal waters in various forms and is useful as a direct indicator of
seawater buffering capacity. TA is defined differently from the alkalinity measurements
typically used in freshwater monitoring. In addition, the above seawater calculations are
sensitive to tiny errors in TA determination, so monitoring programs aim for extreme care
in the collection, handling, and analysis of TA samples.
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.
The primary means of data collection during the 2020 NCCA will be through a specifically
designed application for use on iOS devices (e.g., the NCCA App).Within the NCCA App,
there are a number of information (i) buttons that contain tables, figures, pictures, and
other information summarizing field activities and protocols from the Field Operations
Manual. Field crews are also required to keep the equipment manuals (probes, etc.)
available in the field for reference and for possible protocol clarification.
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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 2020: Quality Assurance Project Plan (EPA-841-R-14-003).
2.6 Recording Data and Other Information
Field data and sample information must be recorded completely, 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. Incorrect
information can result in substantially increased time to process information from the
electronic field forms to the National Aquatic Resource Surveys Information
Management (NARS IM) system. Guidelines for recording field measurements are
presented in Table 2.1.
All samples need to be identified and tracked, and associated information for each sample
must be recorded. To assist with sample identification and tracking, packing slips and
sample labels with sample ID numbers are preprinted and provided by EPA.
2.6.1 Electronic Field Forms
Field crews will utilize the NCCA App to complete data collection. The NCCA App is
available in the iTunes store and will come preloaded on iPads that will be distributed to
all non-contract field crews. These iPads will be designated for crew use during the 2020
season and will be returned to EPA at the end of the field season.
The NCCA App is the required format for data submission as it reduces 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 data entry fields. In addition, the NCCA App generates all sample IDs
based on the initial entry of the CHEM sample ID and includes fish pick lists for consistent
naming of fish species. 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 only to be used if the App fails and will be provided prior to the
field season. Paper packing slips (provided with label packets in site kits) must be
included in every cooler to maintain chain of custody.
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Table 2.1 Guidelines for recording field measurements & tracking information
Activity
Guidelines
Field Measurements
Data Recording
• If recording using the NCCA App, populate all values in the App
• If recording using paper forms (in the event of an App failure):
• 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.
Data Comments
• Comment fields are found throughout the App and associated with every sample and all key
data points.
• Use the provided areas to make comments about any data or sample that will explain any
deviation for normal protocol or will otherwise assist data reviewers in better understanding
the data.
• Be as clear as possible in your comments to convey all necessary information.
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 in Sample Collection form
in the App
Sample Collection and Tracking
Sample
Comments
• Comment fields are found throughout the App and associated with every sample and all key
data points.
• Use the provided areas to make comments about any data or sample that will explain any
deviation for normal protocol or will otherwise assist data reviewers in better understanding
the data.
• Be as clear as possible in your comments to convey all necessary information.
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 App data for accuracy, completeness, and legibility.
• The Field Crew Leader must review all data on the App. Submission of data to NARS IM
confirms review.
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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.
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
Training
• First aid and cardiopulmonary resuscitation (CPR)
• 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
• Flandling 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 VF1F 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
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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
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:
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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
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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2.8.5 Covid-19 Safety Considerations
Safety is the number one concern for all personnel. In implementing the NCCA, crews
should follow their agencies' guidance on maintaining social distance, use of personal
protective equipment, travel restrictions, sanitizing equipment, vehicles and boats, and,
if necessary, hotel rooms. NCCA training and assistance visits will be implemented in a
manner that takes into account Covid-19 safety requirements and restrictions.
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3 Introduction to Sampling
This Field Operations Manual describes procedures for collecting samples for the NCCA
2020. 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, whole fish composite samples to analyze fillets
for human health risks, and will record underwater video of the bottom substrate.
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 Listed Species Considerations
Field crews have the potential to encounter federally listed species and critical habitats
that are protected under the Endangered Species Act (ESA) while conducting field
sampling. Field crew leads are expected to have an understanding of the federally listed
species and their critical habitats and state species of concern that have the potential to
occur at or near a given sampling site, including habitats that will be used to access the
sampling site. Crew leads are responsible for making their crew members aware of
potential occurrences of listed species and their critical habitat. Efforts should be made
to minimize risks to listed species and their critical habitats and avoid the take3 of listed
species while implementing the NCCA field protocols. For example, crews are expected
to:
• abide by all boating speed regulations, including "No Wake" and "Minimum
Wake" zones;
• remain a respectful distance from marine mammals and sea turtles'5;
• designate a marine animal spotter for when the boat is in motion;
• understand the circumstances when it would be necessary to shut down a
vessel due to the presence of a listed species;
• allow a listed species to naturally move away from the sampling area (do not
herd or harass);
• immediately release listed taxa if they are unintentionally collected while
implementing the sediment, benthic macroinvertebrate, or fish tissue sampling
protocols (do not preserve); and
• implement additional limitations that may be established in the scientific
sampling permits.
These best practices are not an exhaustive list of requirements for field crews.
Regulations and guidelines that have been developed for marine life viewing provide
useful risk minimization practices when boating in area that may support listed manatee,
whales, turtles, sea lions, and sharks. Field crews are expected to be aware of the
recommendations and guidelines that apply in a given state and for a given species.
Additional information on boating best practices is available on the NOAA Fisheries Marine
Life Viewing page and provided by the Florida Fish and Wildlife Conservation Commission.
3.4.3 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
a "Take" means to harass, harm, pursue, hunt, shoot, wound, kill, trap, capture, or collect, or to attempt to
engage in any such conduct.
b For whales, remain at least 100 yards away unless other restrictions apply (e.g., 200 yards from killer
whales in Washington State inland waters). For seals, sea lions or turtles in the water, or on shore,
remain at least 50 yards away. To learn more, visit the Marine Mammal Viewing Guidelines and
Distances page, as well as 50 CFR 216.3 and 50 CFR 224.103
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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.
3.4.4 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-Srte ^1
[5.1.2]
Sanralearile
V
Not Ssmpleable
Site Characteristics
[5.1.4]
Relocate
[5.1.3]
VERIFICATION
Secchi Depth Ł6.21
Hydrpgraphk Profile
with Sonde [6.4]
ar*j PAR Meter [6.55
COLLECTION
(MARINE WITH ACTIVE
FISHING GEAR)
If the crew is able to filter the samples
on the vessel, the Enterococci collection
should take place immediately following
the hydrographic profile.
PROCESSING
JL
r
NUT5(N,I>) >
/
CHLA
<
ENTE J
Aliquot [15.3] *
Filters [15.3]
Filters [15.2] *
Figure 3.1 Marine Field Sampling Scenario - Active Fishing Methods
r
k
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Confirm X-Site ¦ ¦ ¦¦¦ ¦ ¦ i
[5.1.2]
f^ampleable Hot Sampleable
Site Characteristics
[5.1.4]
Fishing
(set passive *earI
V
Secchi Depth [5.2]
Hydrographk: Profile
with Sonde [6.4]
and PAR Meter [6.5]
Relocate
[5.1.3]
VERIFICATION
COLLECTION
(MARINE WITH PASSIVE
FISHING GEAR or
ABILITY TO FILTER ENTE
IMMEDIATELY)
CHEM, WCHUB.3]
BENT [13.4]
SEDO, 5EDK, 5EDC, SEDG,
D15N [13.5]
Retrieve
Fishing Gear
PROCESSING
*
NLTTS (N,P> j
/
CHLA
<
ENTE a
ABquot [15.3] '
\
Filters [15.3]
Filters [15.2] *
v
L
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Confirm X-Kte
[5.1.2]
SarnpleaWe I Not Sampleable
^ 7k
Site Characteristics
J5.1.4]
Y
Relocate
[5.1.3]
Secchi Depth [6.2]
Hydrograph*: Profile
with Sonde: [6.4]
and PAR Meter 16.51
" Second UVlD taken if BENT collected
at location other than Y-Lacaticr
VERIFICATION
COLLECTION
(GREAT LAKES WITH
ACTIVE FISHING GEAR)
If the crew is able to filter the samples
on the vessel, the Enterococci collection
should take place immediately following
the hydrographic profile.
Active
Fishing
PROCESSING
JL
r
Figure 3.3 Great Lakes Field Sampling Scenario - Active Fishing Methods
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Confirm X-Site ¦ ¦ ¦¦¦ ¦ ¦'
[5-1-2]
Sairipleable Hot 5am pleable
Site Characteristics Relocate
[5.1.4] [5.1.3]
Fishing
(set passive gear'l
Y
Secthi Depth [6.2]
Hydrographic Profile
with Sonde [6.4]
and PAR Meter [6.5]
CHEM, WCtiL [83
VERIFICATION
COLLECTION
(GREAT LAKES WITH
PASSIVE FISHING GEAR
or ABILITY TO FILTER
ENTE IMMEDIATELY)
MIOCr MICZ [9.3]
PHYT [11.31
BENT [13.4]
UV1D [12.41*
• Second UViD taken if BENT collected
at location other than Y-Location
SEDO, SEDX, 5EDC. SECN6,
[133]
Re-rieve
Fishing Gear
PROCESSING
i
v
k
Figure 3.4 Great Lakes Field Sampling Scenario - Passive Fishing Methods or Ability to Filter Samples Immediately
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4 Pre-Departure Activities
Page 23
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
(Figure 4.1). 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; (3)
that equipment and supplies needed to complete the sampling effort are available and in
good working order; and (4) any and all federally listed species that have the potential to
occur at the sites. After sampling, crews must ensure that: (1) samples are labeled,
packed, and shipped appropriately; (2) the sampling event is communicated to EPA via the
NCCA App submissions; and (3) equipment and supplies are cleaned and replenished as
necessary.
»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 App data submission
•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 iPad or replace batteries
•Refuel vehicle and boat
•Obtain ice and other consumable supplies as
needed
•Package and ship samples & data forms
Figure 4.1 Overview of base site activities
Pre-departure activities are included here, while post-sampling activities are also
discussed in Section 15: Final Site Activities and Section 16: Post-Sampling Activities.
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Pre-departure 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. If applicable and per field crew's standard operating
procedures, Site Packets should include information on federally listed species that may
occur at the site, how to avoid them, and actions to be taken if they are encountered.
Additional pre-sampling 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-parameter 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 on a
weekly basis 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.
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• 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.
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 two years for
calibration. Field crews must use the procedures for the initial setup of the LI-
COR 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 batteries of the
underwater cameras and lights are charged and all components are correctly
attached to the frame.
4.2.1 Initial Assembly and Setup Procedures for LI-COR frame, sensor and
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 the 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 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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Screw (one of three)
Weight (moderate)
Figure 4.2 Attachment of the underwater sensor to Figure 4.3 Lowering frame assembly with sensor,
the mounting rings (adapted from LI-COR, 2006) 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 lightweight 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 -125.0fjmols-1m-2/fjAmp (Each sensor has a unique multiplier value
supplied from the factory)
1. Connect the Quantum LI-190 ambient light sensor to the BNC connector on
top of the LI -1400 labeled 11.
2. Turn on the LI-1400 meter.
3. Press the [Setup] key.
4. Use the left ([<-]) or right ([->]) arrow keys to navigate to "SETUP
CHANNELS".
5. Press the [Enter] key to begin the sensor setup.
6. Use the left ([<-]) or right ([->]) arrow keys to navigate to "M=Light", press
Enter".
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").
8. Press the down ([j]) 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.
9. Press the down ([j]) arrow key; enter "AMB" for the unit label.
10. Press the down ([j]) 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.
11. Press the down ([j]) arrow key; select "Log Routin=none"
12. The remaining options do not need to be set as they apply only when using
a Log Routine.
13. Repeat this entire procedure for channel 12 to setup the underwater sensor
("l2=Light") using "UW" as the label for the channel.
4.3 Equipment and Supply Preparation
Field crews must check the inventory of forms, supplies, and equipment prior to
departure using Appendix A; 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 and/or boat 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 site kits through the supply request form at least two weeks
prior to sampling. Site kits will include sample labels, packing slips, and necessary
shipping labels for sampling one site and are specific to either a marine or Great Lakes
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site. Great Lakes crews sampling at designated human health whole fish tissue sites will
also need to request a whole fish tissue sampling kit along with the site kit. Crews will
automatically receive extra labels and paper form packets as a backup to electronic data
collection prior to sampling, and can request additional as needed. 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 2020 cannot be provided at the beginning of
the field season. Consequently, site kits will be provided to crews 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 & methods for preparation
Solution
Use
Preparation
Bleach (1-10%)
Clean nets, gear, and inside of boat
Add 10 - 100 mL bleach to 1 L distilled water.
Quality Check
Solution for multi-
parameter sonde
Weekly check of meter calibration
In place of weekly internal meter checks
No preparation needed (if purchased as ready-to-use
solution)
Buffered Formalin
Preserve benthic samples
Add 8 tablespoons Borax to 2 gallons 100% Formalin (37%
formaldehyde) solution.
FOR USE AT ALL SITES: Add V+ teaspoon Rose Bengal
crystals to above solution.
Lugol's Solution
Preserve phytoplankton samples
(Great Lakes sites only)
None (included in GL base kits); Lugol's 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
verifying site
sampling permit and landowner access (if required)
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
measurements
Verification form in App
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 Evaluation Spreadsheet. This process
can be completed 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
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sampling will begin) is referred to as the Y-location. If the X-site is not within
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 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 Verification Form in the
App.
6. Record the coordinates of the Y-location on the Verification Form in the App
form in decimal degrees in the NAD 83 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 Verification Form
in the App. 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 Verification Form in the App. No
further sampling activities are conducted at this site.
7. Replace the original site with the next oversample site on the estuary/state
list.
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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• At 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. 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 sketch/map to the NARS SharePoint site when you submit your
data forms.
7. Record the names of the Field Crew Leader, fish taxonomist, and all crew
members. The same name may be recorded twice if the Field Crew Leader is
also the fish taxonomist.
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 if 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.
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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 13.3).
Note: Acceptable means:
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, SN15 isotopes in benthic organic matter).
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) Toxicity [SEDX]
b) Organics/Metals [SEDO]
c) Total Organic Carbon [SEDC]
d) Silt/Clay (Grain Size) [SEDG]
e) Nitrogen Isotopes [D15N] at marine sites only
Indicate if any of the sediment samples were not successfully collected by
marking the "no sample collected" box(es) in the App for each pertinent
sample and supplying a reason for not collecting in the adjacent comment
field.
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] at targeted Great Lakes
sites. 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 and/or fish sample. If
benthos, sediment, and/or fish are collected from a secondary or tertiary location, in situ
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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 form
in the App, 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 37
m 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 or
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 form in
the App, indicate the sediment collection location by filling in the "between
37-100 m 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 or 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 form in the App, indicate the sediment collection zone by
filling in the "between 100-500 m 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
explanations must be completed within the App, as well as pertinent "no
sample collected" boxes.
Note: The Field Crew Leader may choose to make additional sediment grab
attempts.
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m
Figure 5.1 Primary, secondary and tertiary sample collection zones
5.4.2 Fish Samples
The primary fish collection zone at all sites is a radius 500 m from the X-site. 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 when considering sampling locations for fish
samples:
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 primary
500 m radius without success in order to move to the secondary fish collection
zone.
3. When relocating crews should concentrate on 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. Record 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.
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
in repeated attempts to locate suitable i
|
benthos/sediment sampling locations i I
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)
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
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500 m 1000 m 1500 m
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
If no suitable fish are collected in primary zone, and
• Crew observes signs of fish or
favorable habitat/structure
Human Health Fish Tissue (HTIS) ONLY'.
1000 - 1500 meters from X-site
If 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 Primaiy 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 Hydrographic Profile form in the App.
Table 6.1 Equipment & supplies: transparency, DO, pH, salinity/conductivity, temperature, ft light
attenuation
For taking measurements and
calibrating the water quality meter
For recording measurements
multi-parameter water quality meter with DO, pH, salinity/conductivity,
and temperature probes,
extra batteries
de-ionized water (lab certified preferred, but not required)
calibration cups and standards
QCS (used if internal meter checks are not possible)
barometer to use for calibration
thermometer
Secchi disk (20 cm diameter, weighted) & 100' line with clip (marked in
0.5 m intervals)
PAR meter (with LI-190 Quantum Sensor and LI-192 Underwater
Quantum Sensor & cables, independent datalogger)
NCCA App Hydrographic Profile form
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 0.1 m 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 Hydrographic Profile Form in the App,
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 0.5 m markings on the rope to the nearest 0.1 m.
• If the disk hits the bottom before disappearing, water column transparency
depth is greater than the water depth. Fill in the "Yes" circle in the App
next to "Clear to Bottom?" and record the station depth as both the
disappearance and reappearance depth in the "Reading 1" row in the App.
No further measurements or recording are necessary in this case.
3. Slowly raise the Secchi disk until it just becomes visible and record the depth
in the "REAPPEARS" column. Interpolate between the 0.5 m markings on the
rope to the nearest 0.1 m.
4. Repeat steps 1-3 two more times, recording both disappearance and
reappearance depths each time.
5. Use the comment space provided on the Hydrographic Profile Form in the App
to comment on any measurements that the crew feels needs further
explanation 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/QA form in the App.
• 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 requirement
applies for calibrations done both pre-sampling and post-sampling.
• The calibration buffers must be accurate to 0.02 pH units or better.
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• The calibration buffers should be replaced with fresh solutions every three
to four days or sooner if the crew suspects it has become contaminated.
• 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 Calibration/QA
form in the App.
• 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 three to four 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 standard 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).
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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 three to four
days or sooner if the crew suspects they have become contaminated.
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 (N 1ST)-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 Calibration/QA form in the App. 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
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"Quality Control Check" section of the Calibration/QA form in the App. 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
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 temperature ranges below are
from the NCCA 2010 dataset. On the Calibration/QA Form in the NCCA App, record two of
the results (a high and a low temperature from the pertinent range) from the annual
temperature check in the fields provided.
• Northeast: 6.8 °C < T < 32.3 °C
• Southeast: 21.2 °C < T < 33.42 °C
• Gulf Coast: 22.4 °C < T < 36 °C
• Great Lakes: 3.54 °C < T < 30.9 °C
• West Coast: 9 °C < T < 24.1 °C
See below methods for measuring DO, pH, salinity (marine sites) or conductivity
(freshwater sites), and temperature.
6.4 Sampling Procedure - Dissolved Oxygen, pH, Temperature and
Salinity/ Conductivity
1. Measure the total water depth at the Y-location to the nearest 0.1 m and
record on the Hydrographic Profile form in the App. If the sonde is attached to
a data recorder, crews may submit the hydrographic profile data via an
electronic file. If a crew chooses to use this option, ensure that all the data are
saved correctly and check the "Submitted data via eFile" box on the form.
2. Lower the sonde into the water and record DO, pH, salinity/conductivity, and
temperature measurements at the following depths: 0.1 m below the surface,
0.5 m below the surface, every 1 m from depths of 1.0 to 10.0 m, and if the
site is greater than 10 m, every 5 m thereafter. Take the last set of
measurements at 0.5 m from the bottom, making sure to not let the sonde
touch the bottom. Record these results in the Hydrographic Profile form in the
App.
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• NOTE: if the station depth is less than 1 meter, take the measurements at
0.1 meters and mid-depth. Record the pertinent measurements at these
two depths on both the upcast and the downcast.
3. Take a second reading at 0.5 m and then repeat the full sets of measurements
at each of the same depth intervals as the probe is retrieved (upcast) in the
Hydrographic Profile form in the App. Make sure to slide the 'Upcast?' toggle
on the left side of the pertinent data rows in the App to indicate the
measurements that were taken during the upcast (sliding the toggle to the
right turns it green and indicates an upcast measurement). Two examples are
provided below in Table 6.2 that illustrate the depths at which measurements
will be taken.
4. Flag any measurements that the crew feels needs further comment or when a
measurement cannot be made in the Hydrographic Profile form in the App.
Note, measurements may be limited by the length of sonde cable and crews
should record measurements as deep as the cable allows. However, at a
minimum, crews should record to a depth of 30 m in these situations and
comment about the cable limitation.
5. After all in situ measurements have been completed for the sampling day,
perform a "Post-Measurement Calibration Check" of the pH and conductivity
probes. Record these values on the Calibration/QA form in the App.
Table 6.2 Example depth measurement intervals
EXAMPLE 1: EXAMPLE 2: EXAMPLE 3:
Water Depth = 0.8 meters Water Depth = 7.2 meters Water Depth = 23.9 meters
0.1 m 0.1 m 0.1 m
0.4 m 0.5 m 0.5 m
1.0 m 1.0 m
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
8.0 m
9.0 m
10.0 m
15.0 m
20.0 m
23.4 m
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
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two years for calibration. Crews measure PAR at the same depths as other water column
indicators but should not be completed at the same time to prevent the multiparameter
sonde from interfering with the PAR sensor. 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) to the BNC connector
labelled 11 and an underwater sensor (LI-192 Underwater Quantum Sensor)
to the BNC connector labelled 12 as described in Section 4.2.1.2. Enter the
calibration factors (supplied by the manufacturer) for each probe if not
already entered.
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.
7. Scrolling down will move the cursor to the second row of data.
8. Using the left or right keys, navigate until channel I2I is displayed; this
shows the instantaneous reading for that channel allow viewing of both
channels of instantaneous data at once.
9. 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 0.1 m
(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 include using two people to view the two
readings, taking a photograph of the screen, etc.
10. 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 10 m
d) 0.5 m from the bottom
• NOTE: if the station depth is less than 1 meter, take the measurements
at 0.1 meters and mid-depth. Record the pertinent measurements at
these two depths on both the upcast and the downcast.
11. Allow the readings to stabilize at each depth before pressing "Enter" or
recording the values on the data form.
12. Repeat the procedure at the same depths, but in reverse order on the
upcast.
13. Review the saved data by pressing Esc and using the right or left key to
select "LOG DATA" and pressing Enter.
14. Select "View=ALL." Press Enter.
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15. 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.
16. Record the values from both sensors (/jE/m2/s), at the appropriate water
depths of the underwater sensor, in the App. Record the deck sensor
reading in the ambient (AMB) column, and the underwater sensor reading in
the underwater (UW) column.
17. If the sensor hits bottom, allow two to three minutes for the disturbance to
settle before taking the reading.
18. If the light measurements become negative before reaching the bottom
measurement, terminate the profile at that depth and begin to take the
upcast measurements.
19. 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 (see Section
4.2.1).
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 Total alkalinity [ALKT]
Total alkalinity (TA) is a characteristic of seawater that, in combination with other
measurements, can be used to calculate total pH (i.e., coastal acidification) and the
availability of carbonate ions used by marine organisms to produce structural materials
such as corals and shells. TA is also used to calculate the fate of carbon that enters
coastal waters in various forms and is useful as a direct indicator of seawater buffering
capacity. TA is defined differently from the alkalinity measurements typically used in
freshwater monitoring. In addition, the above seawater calculations are sensitive to tiny
errors in TA determination, so monitoring programs aim for extreme care in the
collection, handling, and analysis of TA samples.
7.1 Summary of Method
At marine sites only, two water samples will be taken from the Y-location using the
EPA-provided hand-held peristaltic pump at 0.5 m below the water surface or mid-depth
if station depth is less than 1.0 m. Store sample in cool, dark location (cooler) until
ready to ship.
7.2 Equipment and Supplies
Table 7.1 Equipment & supplies: total alkalinity sample collection
For collecting samples
nitrile gloves
Hand-operated peristaltic pump with flexible gas-impermeable tubing installed
Threaded tubing adapters (3)
Stainless steel 3/8 inch pipe to weight end of intake tube
In-line disposable groundwater filter (0.45 |xm)
HDPE bottle (125 mL, white, rectangular) (2)
Bucket, 5 gallon
Electrical tape, plastic
cooler with ice
For recording
NCCA App
measurements
Total alkalinity sample labels (2)
fine-tipped indelible markers (for labels)
clear tape strips
7.3 Sampling Procedure
See below for step by step procedure for collecting total alkalinity. Collect at the Y-location
and at marine sites only.
Note: Alkalinity samples should be collected between the hydrographic profile measurements
and when the water chemistry samples are collected.
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7.3.1 Sample Collection
1. Fill out both total alkalinity sample labels with the Site ID, visit number, date,
and salinity. The salinity value (0.5 m downcast) will auto-populate from the
Hydrographic Profile form in the App. If the TA sample is not collected at the
same time or location as the hydrographic profile and a new salinity value is
determined, edit the salinity value in the Sample Collection form accordingly.
2. Put on nitrile gloves
3. Ensure that the sampling apparatus is set up with the intake tubing, pump,
weight and threaded tubing adapters inserted into both ends of the intake
tubing (Figure 7.2). The adapter at the inlet end of the tubing holds the
weight in place while the adapter at the outlet side of the pump will receive
the disposable filter.
4. Pre-assemble the filter assembly by attaching the short piece of tubing to the
outlet side of a new filter (note the flow direction arrow on the side of the
filter). This assembly will be attached to the outlet side of the pump after
priming and flushing the system.
5. Fully submerge the inlet portion of the tube into a full bucket of site water and
begin pumping to prime the system. Holding the pump and outlet tubing near
the level of the bucket will help achieve prime more easily.
6. Once good flow is achieved through the tubing, fold and pinch the tube
between the pump and the threaded outlet adapter and quickly remove the
weighted inlet end from the bucket and lower the weighted end to desired
sample collection depth (0.5 m below the surface or at mid-depth if station
depth is less than 1.0 m). Flush the pre-filter tubing by hand cranking the pump
for 30 seconds.
7. While pointing the inlet adapter upward, fold and pinch the tube between the
pump and the threaded outlet adapter and attach the filter (with exit tubing)
to the outlet adapter (do not allow water to enter the filter unless its exit
opening is pointed upward). Continuing to point the filter outlet upward, crank
the pump to fill the filter from the bottom up and expel the air. All pump
cranking should be done slowly and carefully at a speed of approximately one
to two revolutions per second until water is flowing and there are no observed
bubbles in either filter or tubing. During this process, adjust the angle of the
filter as needed to allow air bubbles to exit through the outlet tubing.
8. After all air is expelled, crank the pump for an additional 20 seconds to rinse
filter and outlet tubing. It is no longer necessary to hold the filter upright once
all air has been expelled.
9. Rinse sample container lids with a few ml of sample three times. The process
of overflowing the sample bottle in Step 11 below will provide adequate rinsing
of the container itself.
10. Put outlet tube in sample container so that it is all the way on the bottom. Fill
sample container from the bottom in a controlled manner by turning the
peristaltic pump slowly (approximately one to two revolutions per second).
11. Allow sample container to overflow with bubble-free water by at least three
times the time needed to fill the container to the top (example: if it requires
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10 seconds to fill the bottle, the overflow process should continue for 20
additional seconds for a total of 30 seconds).
12. While continuing to pump slowly, pinch the outlet tube before beginning to
withdraw from the bottom so that, after withdrawal, the water surface is in
the lower half of the threaded portion of the bottle neck (Figure 7.1). If the
removal of the tube leaves the water surface too low in the bottle, start the
overflow process again and try withdrawing the tube a small amount before
pinching and resuming withdrawal.
13. If any air bubbles appear in tubing while collecting the sample, restart the
overflow timing in Step 11.
14. Cap the bottle.
15. Repeat the above steps for the second sample bottle.
16. Tape both lids with electrical tape and place them in a cooler (on ice) and shut
the lid. Do NOT freeze samples.
17. Ensure the Sample ID is recorded on the Sample Collection form in the App.
18. Once the samples are placed on ice, check the 'Chilled?' box on the form in the
App.
19. Both the time that the sample was collected and the salinity value are very
important to the analysis of the TA sample:
a) In the Sample Collection form in the App, enter the time that the sample
was collected as accurately as possible.
20. Discard the filter after use, a new filter will be used at each site. Both pieces
of tubing as well as the threaded adapters should be saved and reused.
21. At the end of the sampling day, rinse the tubing, weight, and pump housing
with Dl water to avoid cross-contamination and corrosion.
TOTAL ALKALINITY BOTTLES
Once filled, the water surface should be in the lower
half of the threaded portion of each bottle neck
Figure 7.1 Target fill range for total alkalinity sample bottles
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r ^
i> Ol
I ••
Peristaltic pump, tubing and weight
Filter and fill tubing
pre-assembled
1. Expel Air
Peristaltic
Pump
^ filter
terminal tube
adapter
2. Rinse/Flush
Figure 7.2 Total alkalinity filter detail
Tubing from pump is flushed. Filter is connected to adapter, inverted and filled with sample to expel air.
Additional water is run through to rinse tubing, filter, and bottle. Bottle filling procedure is then begun.
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8 Water Chemistry [CHEM], Chlorophyll-^ [WCHL],
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.
8.1 Summary of Method
The water chemistry samples will be analyzed for chlorophyll-a [WCHL], 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 HDPE bottle. Water for the chlorophyll-a sample
will be collected and transferred to a separate reusable 2 L amber HDPE 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.
8.2 Equipment and Supplies
Table 8.1 Equipment ft supplies: water chemistry ft chlorophyll-a sample collection
For collecting samples
water sampling device or water pumping system
nitrile gloves
HDPE bottle (250 mL, amber) [CHEM]
HDPE bottle (2 L, amber) [WCHL]
Electrical tape, plastic
cooler with wet ice
For recording
measurements
NCCA App
water chemistry sample label
fine-tipped indelible markers (for labels)
clear tape strips
8.3 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 HDPE 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 HDPE bottle (for water chemistry) and the 2 L amber
HDPE 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 form in the NCCA App.
a) Enter the water chemistry sample ID on the Tracking Form in the NCCA
App.
b) Once the sample is placed on ice, check the 'Chilled?' box in the App.
c) Note anything that could influence sample chemistry (heavy rain,
potential contaminants, etc.) in the Comments section.
d) 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 in the App.
10. Proceed to Section 15.3 for instructions on processing chlorophyll-a and
nutrients water sample to obtain a chlorophyll-a filter and the nutrients
filtrate.
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9 Algal Toxins (Cylindrospermopsin [MICX] and
Microcystins [MICZ])
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 DO 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.
9.1 Summary of Method
Two water samples for algal toxin analysis are taken from the Y-location: one for both
cylindrospermopsin and microcystin [MICX] and one for microcystin only [MICZ]. 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.
9.2 Equipment and Supplies
Table 9.1 Equipment & supplies: algal toxins (cylindrospermopsin and microcystins)
For collecting samples
nitrile gloves
water chemistry sample collection device
1 HDPE bottle (500 mL, white, round, wide mouth) [MICZ)
1 PETG bottle (500 mL, clear, square) [MICX]
Electrical tape, plastic
cooler with ice
For recording
measurements
NCCA App
cylindrospermopsin sample label
microcystin sample label
fine-tipped indelible markers (for labels)
clear tape strips
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9.3 Sampling Procedure
See below for step-by-step procedures for collecting both algal toxin 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.).
9.3.1 Sample Collection
1. Complete the MICZ 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 sample bottles and cover
with clear plastic tape.
a) MICX = clear square 500 mL PETG bottle
b) MICZ = white round 500 mL HDPE bottle
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.
9.3.2 Sample Storage
1. Place the 500 mL bottles in a cooler (on ice) and shut the lid.
2. Ensure the Sample IDs are recorded on the Sample Collection form in the App
along with any pertinent sample information.
3. As soon as you return to your base site (hotel, lab, office, etc.), freeze sample
bottles and keep frozen until shipping.
4. Once the samples are placed in the freezer, check the 'Frozen?' box on the
form.
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10 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.
10.1 Summary of Method
Crews collect and preserve the fecal indicator sample at the Y-location using the method
described in the Sampling Procedure (Section 10.3) below. In addition, crews observe the
area around the X-site and record (on the Site Assessment form in the App) signs of
disturbance that may contribute to the presence of fecal contamination to the waterbody.
10.2 Equipment and Supplies
Table 10.1 Equipment ft supplies: fecal indicator (Enterococci) sampling
For collecting samples
nitrile gloves
PETG bottle (250 mL, clear, square, pre-sterilized)
sodium thiosulfate tablet
wet ice
i i cooler
For recording measurements N(] \ App
10.3 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 0.3 meters below the water surface (or mid-depth if station
depth is less than 0.6 meters).
• 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.
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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 gently shake the bottle 25
times.
5. In the Sample Collection Form in the NCCA App, note the time and
depth (typically 0.3 meters) of the Enterococci collection.
6. Immediately after collection, place the sample on wet ice.
7. Store the sample in a cooler on wet ice to chill (not freeze) for at least
15 minutes prior to beginning the filtration process. Do not hold samples
longer than six hours before filtration and freezing.
8. The filtration procedure is contained in Section 15.2.
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11 Phytoplankton [PHYT] (GreatLakes only)
11.1 Summary of Method
At 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 HDPE 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.
11.2 Equipment and Supplies
Table 11.1 Equipment ft supplies: phytoplankton
For collecting and
preserving samples
water sampling device or water pumping system
nitrile gloves
HDPE bottle (1 L, white, narrow mouth)
wet ice
cooler
Lugol's solution
Pipet (10 mL)
Pipet Bulb
Electrical tape, plastic
For recording
measurements
NCCA App
phytoplankton sample label
fine-tipped indelible markers (for labels)
clear tape strips
11.3 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.
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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.
7. The sample must be preserved by adding 10 mL of Lugol's solution to the bottle
within two hours of collection.
8. After preservation, replace the lid and seal tightly with electrical tape.
9. Record the collection data on the Sample Collection form in the App. Include
the depth of collection, time of collection, and time of preservation.
10. Ensure the sample ID is recorded on the Sample Collection form in the App.
11. After the sample is preserved, check the 'Preserved?' box on the form in the
App.
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12 Underwater Video [UVID] (Great Lakes only)
12.1 Summary of Method
At Great Lakes sites only, crews will use underwater video cameras to capture at least 1
minute of benthic video at the Y-Location. Video will be used to document the benthic
habitat composition and record the presence of invasive species like zebra and quagga
mussels and round gobies, or other organisms. The underwater video carriage consists of a
steel frame onto which two cameras and two lights are attached. One camera looks down
(Camera A) and the other has an oblique view (Camera B). Figure 12.1 shows the fully set
up underwater camera assembly.
Figure 12.1 Underwater video assembly.
Includes cameras, lights, frame with scale markings on footings, and pre-attached leader.
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12.2 Equipment and Supplies
Table 12.1 Equipment ft supplies: underwater video
Component
GoPro Hero 7 Black camera
(2 — Units A and B)
Description
Camera with waterproof housing, battery, and
charging cord. Safety tether attached.
Function
Record video
Lens covers (2)
Protective lens cover, removed for insertion
into the dive housing. Replace when camera is
taken out of housing.
Lens protection
Suptig LED underwater video lights
(2Befng^
Light with internal battery and charging cord.
Light
Video carriage
Stainless, welded, 18" tall; 21" wide at base.
Leader with plastic tubing attached. Footings
have black and yellow scale bar tape with each
cube measuring 24 mm.
Frame for cameras
for deployment from
boat
Camera Clamps
Light Clamps
2 pipe clamps with safety tethers
2 GoPro clamps
Lowering line
100 ft rope, with carabiner and float attached.
Attach camera to the
video carriage
Attach lights to the
video carriage
Lowering video
carriage
Media (Cards A and B)
Crews provided with at least 2 Micro SD cards
for capturing, storing, and sending video to
USEPA
Memory
USB Power Supply
Laptop/computer
USB power station
Provided by crew
Recharging cameras
and lights
Transfer files and
store data
Supplies
Allen wrench
Screwdriver
Metal coin
Zip Ties
Lens cleaner cloth
Carrying case
Index cards and sharpie
Description/F unction
Used to tighten set screws on clamps
Used to tighten clamps
Used to unscrew waterproof chargingŁort on lights.
Used as back-up to secure clamps to carriage
Used to clean camera lens
Protective case for storing and transporting cameras, lights and supplies
Used to indicate site number in video at each site.
12.3 Underwater Video Carriage Set-up
Underwater camera settings will be adjusted prior to shipment to field crews. Information
here will allow field crews to verify camera setup and assemble the underwater camera
system. Camera settings should not need to be changed. If settings have been altered
between shipment and deployment, see Table 12.2 to verify and restore settings. The
video carriage system should be assembled for the field day and does not need to be
disassembled between sites.
12.3.1 Setting up Video Carriage System
Set up cameras on small boat video carriage as described in the steps below and as shown
in Figure 12.2 through Figure 12.4. The process of renaming files and backing the videos
up to a computer will not require removal of the Micro SD cards from the cameras. If
removal of a card is necessary, be sure not to interchange the cards so that each card only
has either oblique or down-looking videos on it.
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1. Make sure the camera settings are as shown in Table 12.2. Important note: unlike
previous versions of the GoPro camera, the protective outer lens on the Hero 7 has
been removed in order to fit in the dive housing. When handling the camera
outside the housing for charging or downloading, use care to avoid scratching the
lens, as it does not have a cover. If the camera needs to be stored outside the dive
housing, put the protective lens cover back on the camera.
2. To remove camera from waterproof dive housing, push the button on top of black
latch to the right, then lift front of latch up and release from the back cover. To
place camera in waterproof dive housing, close the back cover and pull the black
top latch over back cover. Push latch down to click into place. Ensure no debris is
stuck in rubber O-ring such that the case would not be completely sealed.
3. Attach Camera A with Micro SD card A in the dive housing in the down-looking
position as show in Figure 12.2 and Figure 12.3 using the clamp. Attach safety
tether as shown. Use a screw driver or other tool to tighten the clamp. Add zip ties
around clamps and carriage as back-up.
4. Attach the A light in the down-looking position opposite the camera as shown in
Figure 12.2 and Figure 12.3 using the clamp. Tighten the clamps until the camera
is held firmly. Add zip ties around clamps and carriage as back-up.
5. Attach Camera B with Micro SD card B in the oblique position as show in Figure
12.3 and Figure 12.4 using the clamp. To capture the foreground, the camera
should be facing slighty downward rather than straight out from the frame. Attach
safety tether as shown. Add zip ties around clamps and carriage as back-up.
6. Attach the B light in the oblique position next to camera B as shown in Figure 12.3
and Figure 12.4 using the clamp. Tighten the clamps until the camera is held
firmly. Add zip ties around clamps and carriage as back-up.
7. Turn on the cameras and lights and check the aim. Down-looking camera should be
pointing straight down with both of the taped video carriage arms visible in the
frame for size referencing. Oblique camera should be tilted down approximately
20° from vertical to place the benthic horizon at the approximate midpoint of the
image.
8. If a safe location is available for storage, cameras/lights can remain on the video
carriage at the end of the day. If there is risk of theft or jostling, remove cameras
and lights for storage. GoPro and dive housing can be removed by unscrewing
GoPro clamp (with screwdriver if needed). When reassembling, make sure GoPro
clamp is tight enough to prevent movement of the camera/light, but not
overtightened, because the plastic parts can break with overtightening.
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Figure 12.2. CoPro camera A and light mounted on carriage.
Note the orientation of clamps (screws for tightenting facing in), tether on camera, and cable ties on clamps.
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VowN- LOO^Ifiltr
Figure 12.3. Approximate aiming angle of camera and lights for the oblique (B) and down-looking (A) cameras.
Note the orientation of the tripod adapter (the fitting between the red locking thumbscrew and the camera or
light) which is facing backward relative to the camera lens.
Figure 12.4. GoPro camera B and light mounted on carriage.
Note the orientation of clamps (screws for tightening facing in), aiming of camera, tether on camera, and cable
tie on light. To capture the foreground, the camera should be facing slighty downward rather than straight out
from the frame.
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INSTRUCTIONS
Thank you for choosing this product
please read the instructions before use
USB
Charging batteries
Unscrew the waterproof plug connect the 5V USB cable
to charge the built in battery, the LED will flicker when
charging, the charging completed LED will light, the
charging duration is about 3 5 hours
• LED1 bright that there are 1 2S% power
• LED2 bright that there are 26-50% power
• LED3 bright that there are 51-75% power
• LŁ04 bright that there are 76-100% power
»Built in battery capacity 2600MA/H. the input voltage
5V / 1000MA
Power on / off
Press and hold the button for 1 second to turn the LED
on or off.
LED mode switch
LED light mode switch, there are four modes, each
press the key can be different mode switch
• LED1 flashes to indicate SOS mode
• LED2 flashing I file about 2B00Lui 7 87 inches
• LED3 flashing II file about 3800Lux 7 87 inches
• LED* flashing III file about SSOOLux 7 87 inches
» Before diving, please confirm that the USB waterproof
screw is tightened
• Pay attention to USB thread waterproof rubber nng
cleaning, tiny sand may cause leakage products
leakage
• When the LED lights up, it will generate heat please
do not block it wtth cotton cloth and other objects,
keep it well ventilated when using
Figure 12.5. Manufacturer's instructions for lights.
12.3.2 Operating Camera and Lights
GoPro Camera settings should be pre-set when sent to field crews. Settings should match
settings shown in Table 12.2. Directions for changing settings are also given in Table 12.2
To turn on the camera and/or to change setting, press the mode button on the right side
of the camera. It is much easier to change settings with the camera out of the housing
using touch screen, but be careful not to touch the lens, which does not have a lens cover
in order to fit in waterproof housing.
• To enter settings mode, touch screen once and follow the directions in Table 12.2.
• To view saved video files, swipe bottom to top on back screen and select video to
view.
Cameras should always be set to QuickCapture. When set to QuickCapture, pushing the
record button on top of camera when the camera is off will turn the camera on and start
recording. Pushing the same button again will stop recording and turn the camera off. This
setting helps saves battery life.
The brightest light setting (mode 4) should be used for sites deeper than 3 m. A lower
setting can be used for shallow sites to help preserve battery. Turn the light on by
pressing and holding ON button. Press button again to toggle through light settings. Table
12.2 also describes settings for lights. Figure 12.5 includes manufacturer's instructions
for lights.
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Table 12.2 GoPro Hero 7 camera and light settings and directions.
Option
Mode
Resolution
Frames per second
Field of View
Low light
Stabilization
Protune
Short Clips
Capture Mode
Light Level
GoPro Hero 7 Black
Setting
Video
1440
60
Wide
Auto
OfF~
Off"""
X
Video
Off
Voice Command/Voice
Control (audio icon)
Camera Beeps (music icon) | On
. ____
QuickCapture (bunny icon)
Auto Screen Lock (lock icon)
Wireless Connections
Beep volume
Auto power off
'led
Video Compression
Default Mode
Voice Control
Landscape Lock
Screensaver
Brightness
OPS
Language
Video Format
IIDMI Output
Audio input
Off
Off
High
Never
All on
H.264 + HEVC
Video
Off
Off [important!]
1 min
80% or as needed
On
English
NTSC
Media
Directions
Swipe screen left/right to switch among time lapse, video, and
photo modes.
Touch Video Settings button on bottom of screen.
Touch box on left of screen and click x.
Touch button in lower left of screen. Looks like video camera
if set to video, looks like circular arrow if set to looping.
Swipe down, icons are blue if on OR Swipe down, touch
preferences, touch Voice Control.
Swipe down, icons are blue if on
Swipe down, touch preferences, touch connections
Swipe down, touch preferences, touch General
Swipe down, touch preferences, touch Voice Control
Swipe down, touch preferences, touch Touch Display
Swipe down, touch preferences, touch Regional
Swipe down, touch preferences, touch Input/Output
N/A |
Suptig 84 LED light
I in <3m water to |
save battery; use
IV in deep water.
Set to II at power on. Push on button to cycle through levels.
12.4 Underwater Video Collection
At all Great Lakes sites, a minimum of 1 minute of video of the benthic habitat will be
collected using an underwater video camera system. Video will be used to document the
bottom composition and record the presence of invasive species like zebra and quagga
mussels' round gobies, or other organisms.
12.4.1 Summary of Method
Follow directions in Section 12.4.2 to deploy camera and collect underwater video.
Before deploying, crews should always verify that all cameras and lights are fully
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waterproof and have adequate battery/memory life, are on, and pointed correctly. After
cameras and lights are turned on, the iPad displaying the NCCA App Verification Form or a
notecard with the site and date is held in front of EACH camera lens (see Figure 12.6).
When using the App screen, ensure date has been entered and is visible to the camera
along with the blue header information. Make sure information is close enough to be read
but not so close that information is cut off the screen. Video carriage is then lowered
slowly to the bottom and slack is let out to prevent the carriage from dragging.
NGL20_IL-10001, Visit 1
Version 2.6
gP This form has been thoroughly reviewed and is ready for submission
Site name
Lake Michigan
Date collected
06/12/2020
Crew
GL4
Did you sample this site? (•) YES Q NO
Choose method used:
O Marine
(•) Great Lakes
Station Depth (m):
21.8
Arrival Time: 09:53
Depart Time:
Figure 12.6 Example of verification form in the NCCA App showing header information prior to underwater
camera deployment.
The target length of videos is at least one minute. In some cases (e.g., in swift currents or
shipping lanes) it may be difficult to collect a full minute of video. In those cases, crews
should collect whatever they can. Short videos are preferred over no videos, but if there
is risk to crew or equipment, do not deploy the underwater video carriage. If the video
carriage is dragging along bottom it should be retrieved immediately. In areas of swift
current or suspected underwater obstructions (wood, marine debris), do not deploy the
video carriage.
After video collection, the carriage is then retrieved, and cameras and lights are turned
off. Video quality should be verified by reviewing on the GoPro camera after collecting at
each site. Video section(s) of the Sample Collection form in the App should be filled out in
the field, except file names.
Video is collected at the Y-Location. The camera can be deployed at approximately the
same time as the in-situ measurements and water collection activities from the opposite
side of the boat. Avoid heavy disturbance of the bottom with anchors or sediment
samplers during the video. One person can operate and lower the cameras.
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If a benthos grab is successful at the Y-Location, this is the only video that is collected. If
the benthos grab is unsuccessful at the Y-Location, then the sediment collection protocol
(Section 13.3) should be followed to move within 37 m, then 100 m, then 500 m of the X-
site to obtain a successful benthos sample. When a successful benthos sample is obtained,
a 2nd video should be collected at the same location as the benthic grab was collected. At
the start of this video, scroll the benthic collection portion of the Sample Collection form
on the App to just below the header and hold it in front of EACH lens (see Figure 12.7).
Ensure that the distance from the X-site where the video is being collected is displayed
(or print that information on the previously used notecard).
Menu
NGL20JL-10001. Visit 1 MM
Version 2.6
Benthic (BENT)
(1L HDPE bottle)
No Sample
Collected ~
Benthic Collection Location:
O Within 37m from X-site (•) Between 37-100m from X-site O Between 100-500m from X-site
Grab area (m2):
0.05
Select grab type:
STANDARD.PO...
Number of grabs: (•) 1 *" Note: 2 grabs are required Sieve size: O 0.5 mm
for samples less than 0.03m2 ***
O 2 O 1.0 mm
Depth (cm)
(Should be >7 cm)
8.5
No. of jars:
Preserved? Q
Figure 12.7 Example of sample collection form in the NCCA App showing header and benthic grab information
prior to underwater camera deployment.
Fill out the information on the Sample Collection form in the App for the 2nd video.
Additional space is included on the field form if multiple videos are collected for any
other reason as well. At the end of each field day, video files will need to be transferred
from GoPro cameras to a computer and backed up, at which point the rest of the field
form (video filenames) can be completed. Directions for doing this are given in Section
12.4.3. Crews will also need to charge lights and camera batteries using charging cords
and USB power supply.
12.4.2 Deploying GoPro video carriage
The detailed steps for deploying GoPro Video carriage and collecting underwater video are
shown below. The video carriage should be treated with the care and clean technique of a
PAR sensor, Hydrolab sonde, or other delicate and expensive instrument. Store video
carriage in a dedicated space where cameras will not be jostled between sites.
1. Before every deployment, check the following:
• Both camera cases are fully closed
• Both cameras have sufficient battery life
• Both cameras have adequate memory space
• Both cameras are both aimed correctly
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• Both lights are on
• Both lights and cameras are aimed correctly
• Both charging ports on lights are closed securely
• All clamps are tight
2. Attach the lowering line to the video carriage above the permanent leader with
the carabiner provided. The purpose of the leader is to prevent the line from
dropping inside the carriage and fouling the cameras or lights.
3. Attach the other end of the lowering line to the boat and make sure there is
always a float on the end of the lowering line in case it is dropped accidentally or
must be jettisoned. If deploying in swift currents, do not tie off to the boat.
Instead, have a second crew member hold the float end during deployment.
4. Determine the approximate depth and clear 2x length of line to reach the bottom.
5. Prepare the Site Verification screen on the App that displays the site ID and date
(or notecard with site ID and date). For second videos and if the video is not
collected at the Y-Location, display the benthic section of the Sample Collection
form to show the site ID and if the video location is within 37 m or within 500 m of
the X-site (or print on notecard).
6. Turn on both lights:
a. Press on button once to turn on.
b. Three indicator lights show the brightness setting.
c. Set brightness to mode 4 for deployment, unless the site is very shallow.
Toggle through brightness settings by pressing on button again. All four
LEDs above the on button will be illuminated when set to brightest setting.
If site is shallow, a lower setting can be used to save battery.
d. Verify light is on by waving hand in front of it.
7. Start collecting video:
a. Turn on the GoPro cameras by pressing the red circle button (black square
when inside waterproof housing) on top of the camera (Quick capture mode
set to ON). You will hear three quick beeps indicating the camera is
recording.
b. Hold the iPad screen or card with site information and date in front of EACH
camera.
c. Double check the battery life and settings.
8. Lower the camera slowly on the windward side of the boat until the camera hits
the bottom. When bottom is reached, let out enough slack to prevent the boat
from dragging the carriage.
9. If there is risk of dragging, pull carriage up immediately and re-deploy if safe. In
areas of swift current or suspected underwater obstructions (wood, marine debris),
do not deploy the video carriage.
10. Continue recording until you have captured at least 1 min of footage at the
bottom.
11. Retrieve carriage:
a. Pull up line. Pull up from bottom quickly to avoid dragging along the
bottom. Avoid banging the rig against the side of the boat.
b. Stop video recordings by pressing the red circle button.
c. Turn off lights by pressing and holding on/off button until light goes off.
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d. Verify the light is turned off by waving hand in front of light.
12. Review video footage to ensure capture and quality:
a. Dry camera and remove camera from waterproof case, ensuring camera
does not get wet.
b. Press on/mode button on right side of camera.
c. Swipe bottom to top to view files stored on camera.
d. The most recently collected video will play automatically. To see older
videos, select the gallery icon in the upper left corner of the screen. The
most recent video will be shown in the upper left. Touch the desired video
to review, and touch play to watch.
e. Turn off camera by pressing and holding side on/mode button to turn off
camera.
f. Return camera to waterproof case, snap shut, and re-collect if necessary.
13. The Sample Collection form in the App should be filled out in the field, except for
the filenames for A (downlooking) and B (oblique) videos, which will be completed
at the end of the day when files are transferred to a computer. Data should
include:
a. GPS coordinates of video location
b. Distance from X-site (e.g., within 37 meters, 37-100 meters, or 100-500
meters)
c. Depth of video location
14. At the end of each field day, charge lights and camera batteries using charging
cords and USB power supply.
12.4.3 Transferring and Backing up Video Files
Procedures for transferring video files via a USB cable to a computer and/or external hard
drive are described below. Best practice is to always store videos in at least two locations
in case of failure or damage to one.
1. Plug USB cable into Mini USB port on side of camera and laptop/computer. Take
care not to scratch lens when camera is outside waterproof housing.
2. Turn on camera by pressing On/Mode button. Camera should show up as a drive on
laptop. Note if camera "locks up" or stops responding, remove and replace the
battery to reboot the camera.
3. Review each file for the site information at the beginning and change file name of
the videos stored on the Micro SD card using the following convention:
SITEID_V1_DATE_X_01, where:
• SITEID - the NCCA SitelD
• V1 -- visit number, either V1 or V2
• DATE -- the date collected in format YYYYMMDD
• X -- either A for the downlooking camera or B for the oblique camera
• 01 -- the number video collected (always 01 unless the camera is deployed
more than once at a site)
¦ Examples:
• N G L20 J L -10001 _V1 _20200612_A_01
• NGL20 IL-10001 V1 20200612 B 01
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• N G L20 J L -10001 _V1 _20200612_A_02
• NGL20JL-10001 _V1 _20200612_B_02
4. Copy all files to the computer.
5. Do not delete files from Micro SD card, as this is how files will be transferred to
EPA. The file names on the computer and on SD card should be identical and both
follow the naming convention above.
6. If your Micro SD cards are getting close to full, request additional cards using the
Supply Request Form.
7. Complete field forms (Figure 12.8) by entering the filenames associated with the
videos collected for each site.
8. If no underwater video was taken at the site, fill in the "no sample collected" box
and provide reason in the comment field.
9. As the Micro SD cards fill, request an additional pair of cards and send both full
Micro SD cards with videos to GLTED using a T7 FedEx label from your base kit. Be
sure to request the replacement cards with enough lead time to receive and
replace the cards before the cards are completely full to ensure videos can be
taken at every site.
10. In the NCCA App, access the Tracking Form for each site from which videos were
collected and are being shipped. Mark each UVID sample (full set of videos form
one site) as shipped and submit the tracking form.
11. At the end of the field season, each crew will receive instructions on how to return
all underwater camera equipment including all remaining Micro SD cards with
videos saved to GLTED.
12. Do not delete back-up videos on your computer for one year in case Micro SD cards
are damaged or lost in transit.
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12:49 PM Wed Mar 11
•=¦ 56% ¦
Menu
NGL20JL-10001, Visit 1
Version 2.8
SAVE
NCCA/NGLA 2020 SAMPLE COLLECTION
Underwater: Digital Video Recording (UVID)
Y-Location Video:
No Sample Collected ( ]
Latitude:
xx.xxxxxx
Longitude:
-XXX.XXXXXX
Depth (m):
Video taken:
O Within 37m from X-site O Between 37-100m from X-site O Between 100-500m from X-site
A Camera
Filename:
SITEID_V1_DATE_A_01
B Camera
Filename:
SITEIDV1_DATE_B_01
Additional Video: Use if more than one video is collected at the Y-Location or if the benthos sample is collected
at a location other than the Y-Location.
Latitude:
XX.XXXXXX
Longitude:
-XXX.XXXXXX
Depth (m):
Video taken:
O Within 37m from X-site O Between 37-100m from X-site Q Between 100-500m from X-site
A Camera
Filename:
SI TEI D_V 1 _D AT E_A_0 2
B Camera
Filename:
SITEID V1 DATE B 02
©
Additional Video: Use if more than one video is collected at the Y-Location or if the benthos sample is collected
at a location other than the Y-Location.
Latitude:
XX.XXXXXX
Longitude:
-XXX.XXXXXX
Depth (m):
Video taken:
O Within 37m from X-site O Between 37-100m from X-site O Between 100-500m from X-site
A Camera
Filename:
SITEID V1 DATE A 03
B Camera
Filename:
SITEID V1 DATE B 03
o
Figure 12.8 Example of sample collection form in the NCCA App, UVID information.
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13 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,
acute whole sediment toxicity, and nitrogen isotopes in sediment. 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 separate sample containers.
Crews collect 2.5 L of sediment (1.5 liters at Great Lakes sites) to submit for chemistry
(contaminants), toxicity, and grain size analyses. At marine sites only, an additional
sample is collected for nitrogen isotopes.
13.1 Summary of Method
A 1/25 (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 sediment contaminant, toxicity and the marine-only
D15N sample collection. 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 form in the App. 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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13.2 Equipment and Supplies
Table 13.1 Equipment ft supplies: sediment collection
For collecting samples
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)
Phosphate-free detergent such as Liquinox
Formalin (100% buffered) with stain
Graduated cylinder for measuring formalin
Rose Bengal Stain (for staining formalin solution)
Borax
ruler (cm)
wash bottle (for ambient water)
stainless steel mixing pot or bowl with lid
Spoon, stainless steel (15") or Teflon spoons/scoops/spatula
HDPE bottle(s) (1 L, wide-mouth) [BENT]
glass jar (2, 60 mL, amber) [SEDC, D15N]
glass jar (120 mL, amber) [SEDO]
plastic 6 mil bags (2, 1 quart) [SEDG]
Bucket w/screw top lid (0.6 gallon) for marine SEDX
Bucket w/snap top lid (1 quart) for Great Lakes SEDX
scrub brush
cooler with wet ice
For recording
NCCA App
measurements
pencils (for inner labels)
fine-tipped indelible markers (for outer labels)
clear tape strips
13.3 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. At the next
station the sampler, spoons and mixing bowl or bucket must be washed with phosphate-
free detergent such as Liquinox 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.
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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.
3. Lower the grab sampler through the water column such that travel through the
last five meters is no faster than about 1 m/sec. This procedure 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 cm
for the benthic macroinvertebrate grab (see Figure 13.1).
• 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 cm 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 form in the App 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 form in the App.
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 least 7 cm deep with even surface
Unacceptable grab Unacceptable grab
Sloping surface Insufficient volume
Figure 13.11llustration of acceptable Et unacceptable grabs for benthic community analysis.
An acceptable grab is at least 7 cm in depth, but not oozing out of the top of the grab and has a relatively
level surface.
13.4 Processing Procedure - Benthic Macroinvertebrate [BENT]
Composition And Abundance
Grab samples obtained to assess the benthic macroinvertebrate community are processed
as outlined below.
1. Record the deepest recorded salinity at sampling location and write on the
BENT bottle label.
2. Measure the depth of the sediment at the middle of the sampler to the
nearest Vi centimeter and record the value on the Sample Collection form
in the App. The depth should be >7 cm if possible (see previous section).
• Record the grab number in the app and any descriptive information
about the grab, such as the presence or absence of a surface floe, color
and smell of surface sediments, and visible fauna.
Unacceptable grab
Wash-out
Unacceptable grab
Overfilled
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3. 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.
• 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.
• In the rare event that a federally listed benthic species is observed in
the grab sample (e.g., listed freshwater mussels, black or white
abalone, etc.), gently remove the individual from the sample and
quickly return it to an area where is it unlikely to be sampled again to
minimize stress. Make notes in the comments area in the NCCA App
regarding any organisms removed from the sample, including best field
identification and "ESA".
4. 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 HDPE bottle (which should be placed in
the sieve or a bucket in case some of the sample spills over).
5. Complete the BENT sample label with Site ID, date collected, visit number,
and jar number.
6. Attach the completed label to the 1 L wide mouth sample bottle and cover
with clear plastic tape.
7. 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.
8. 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.
9. Record sample collection location and the total number of jars on the
Sample Collection form in the App.
10. Carefully inspect the sieve to ensure that all organisms are removed. Use
fine forceps (if necessary) to transfer fauna from the sieve to the sample
bottle. Once again, do not preserve organisms that have the potential to be
a federally listed species.
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11. 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.
• If rose bengal staining of sample is not evident, you may need to add
more preservative.
• 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 procedure
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 six percent.
• Once preserved, check the 'Preserved?' box on the Sample Collection
form in the App.
12. 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.
13. 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-
contamination of samples. Be sure to rinse the brush between each sieve
cleaning.
13.5 Processing Procedure - Sediment Composition, Chemistry,
Toxicity AND Nitrogen Isotopes
In addition to grab samples collected for benthic community analysis, additional grabs are
collected for chemical analyses (organics/metals and TOC), grain size determination,
acute toxicity tests, and (at marine sites) nitrogen isotopes. The top two centimeters of
these grabs are removed, homogenized, and split into these five 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 13.3 and Figure 13.1
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
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presence in the sediment sample collection zone comment field on the
Sample Collection form in the App.
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
phosphate-free detergent such as Liquinox and rinsed with ambient site
water 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 phosphate-free
detergent such as Liquinox and rinsed with ambient site water) 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 2.5L at marine
sites and 1.5 liters 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 form in the App. 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 describe the proportions of sediment collected from each
zone in the comments section for each sample.
6. Stir the sediment sample with a Teflon paddle or stainless steel spoon until
it's thoroughly homogenized and takes on a uniform color and consistency.
This will take between 2 and 10 minutes. Divide the composite into the
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 1/2 inch
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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.
• Record any comments on the Sample Collection form in the App.
• Freeze the sample as soon as possible and keep frozen until
shipped.
• Fill in the "frozen" bubble in the App 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 (for marine samples) or 1 quart bucket (for Great Lakes
samples) and cover with clear plastic tape.
• Using the stainless steel spoon, fill the bucket with the amount
of sediment specified below:
o For marine sites the preferred volume is 1800 mL (which
will fill the 0.6 gallon bucket approximately 2/3 full) but
if that is not possible the minimum volume required is
900 mL.
o For Great Lakes sites the preferred volume is 900 mL
(which will fill the bucket approximately % inches from
the rim of the 1 quart bucket) but if that is not possible
the minimum required is 400 mL.
• Replace the lid and tighten so that the locking mechanism
engages and holds the lid tightly closed.
• Record any comments on the Sample Collection form in the App.
• 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 in the App 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.
• 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 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.
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• Record any comments on the Sample Collection form in the App.
• Freeze the sample as soon as possible and keep frozen until
shipped.
• Fill in the "frozen" bubble in the App 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 any comments on the Sample Collection form in the App.
• 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 in the App to confirm that the sample
has been chilled.
e) NITROGEN ISOTOPE (D15N) marine sites only:
• Complete the D15N sample label with Site ID, date collected,
and visit number.
• 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 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.
• Record any comments on the Sample Collection form in the App.
• Freeze the sample as soon as possible and keep frozen until
shipped.
• Fill in the "frozen" bubble in the App to confirm that the sample
has been frozen.
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Page 78
Crews collect fish at all NCCA sites. At revisit sites, ecological contamination of fish tissue
or ecofish (FTIS) and fish tissue plugs (FPLG) collection should be attempted during visit 1.
If a crew is unsuccessful collecting FTIS or FPLG during visit 1, then attempt to collect
during visit 2. At Great Lakes revisit sites, crews that are unsuccessful at collecting the
human health fish tissue (HTIS) 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 14.1 for detailed information
regarding ecofish sample collection.
In addition to whole fish samples collected at all sites for ecological risk purposes, crews
will also collect fish tissue plugs at all non-enhancement 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 14.2 for
a detailed discussion offish tissue plug collection.
Finally, crews at all 225 probabilistic nearshore Great Lakes sites (sites whose prefix
begins with NGL20), all 38 Great Lakes island sites (sites whose prefix begins with ISA20),
and all 12 Great Lakes park sites (sites whose prefix begins with NPA20) will collect a fish
composite sample to analyze contaminants in fillet tissue for human health applications
(HTIS). Refer to Section 14.3 for detailed information regarding samples collected for
human health fish tissue contaminant analysis. Note that human health fish tissue samples
will NOT be collected at Lake Erie (LEA20) or Green Bay (GBA20) intensification sites.
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. For those instances, apply the
fish distribution scheme described in Section 14.3.2.
At all sites and for all sample types, crews are never to collect species that are federally
listed as threatened or endangered under the Endangered Species Act for tissue samples.
If a federally listed species (e.g., fish, mammal, sea turtle, etc.) is encountered while
fishing (netted, hooked, etc.), crew members are expected to immediately release the
individual following identification in an area where it is unlikely to be captured again and
cease sampling for five minutes to allow the individual to safely leave the area. Record
the encounter with the listed species by selecting the ESA button in the NCCA App and
record the species, number of individuals, and condition of the individuals. Prior to
restarting fish collection, field crews should evaluate whether alternative fishing methods
that are less likely to encounter listed species are available.
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14.1 Ecological Contamination Fish Tissue Collection [FTIS]
14.1.1 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 form in the App. Also record
sample ID, species retained, and specimen lengths on the Eco Fish Collection form in the
App. Crews will also indicate the date of collection and the coordinates of the location
where fish were ultimately caught.
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:
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 to collect fish at several sampling locations
within the original 500 m radius without success.
3. When relocating crews should concentrate on signs of 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. 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.
5. If fish are collected in more than one zone fill in the bubble of the zone where
the majority of the fish were collected and describe the proportions of fish
collected from each zone in the comments section for each sample in the App.
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 14.2
• Southeast - Table 14.3
• Gulf of Mexico - Table 14.4
• West -Table 14.5
• Great Lakes - Table 14.6
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If a full composite sample is not collected after three 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. Alternative fish
species should be limited to bony fish. Cartilaginous fish and Moray eels (Family
Muraenidae) should not be submitted for this indicator or for the fish plug sample.
Regardless of the species that is ultimately collected, all fish in the composite MUST be of
the same species and meet size requirements.
Crews are expected to know and be able to identify the federally listed species and state
species of concern that have the potential to occur at a given sampling site. If a listed
species is visually observed prior to initiating the sampling, allow the species to naturally
depart the area without herding or harassing. If a listed species is encountered (stunned,
netted, hooked, etc.), crews are expected to immediately release the fish following
identification in an area where it is unlikely to be captured again and cease sampling for
five minutes to allow the fish to safely leave the area.
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
in the App. 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 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 comment fields
provided in the fish sample collection form in the App. At Great Lakes sites where crews
will collect both ecological fish tissue and human health fish tissue samples, but they
collect 10 or fewer fish, they must follow the fish distribution scheme described in
Section 14.3.2.
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14.1.2 Equipment and Supplies
Table 14.1 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)
For storing and
preserving fish
composite sample
zip-top bag(s) (plastic, 2 gallon)
Plastic bag (large, composite)
zip-top bag(s) (sandwich size) — for labels
cooler
plastic cable tie
dry ice or wet ice (for temporary transport)
side cutter (cleaned with phosphate-free detergent such as Liquinox between sites)
For documenting the
fish composite
sample
NCCA App
fish tissue sample labels
fine-tipped indelible markers (for labels)
Tyvek label tag with grommet
clear tape strips
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 14.2 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.
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.
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• 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 form in the App.
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 form in the App. Record
sample ID, species name, and specimen lengths on the Eco Fish Collection
form in the App. Make sure the sample ID recorded on the collection form
match those on the sample labels.
7. While wearing clean nitn'le gloves, remove each fish retained for analysis
from the clean holding container(s). If needed, dispatch larger fish using
the most humane method available.
8. Place all fish from the composite in a two-gallon zip-top 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 phosphate-free detergent such as Liquinox and rinsed
with ambient site water 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 Eco Fish Collection form in the App. Be sure to record scientific name
and minimum and maximum lengths on the labels.
• Place the interior label inside a small (sandwich-size) zip-top 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.
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12. Prepare a sample label for the outer bag, ensuring that the label
information matches the information recorded on the Eco Fish Collection
form in the App. 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.
14.1.3 Sample Storage and Shipping Preparation
1. After the sample is packaged, immediately place it on dry ice for shipment.
• Check the "frozen" box on the Eco Fish Collection form in the App 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 two 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 they 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 14.2 Northeast region primary and secondary marine target species - whole body fish tissue collection
(Ecofish)
NORTHEAST REGION PRIMARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*
Ictaluridae
Ameiurus catus
White catfish
Primary
Ictalurus punctatus
Channel catfish
Primary
Moronidae
Morone americana
White perch
Primary
Paralichthyidae
Paralichthys dentatus
Summerflounder
Primary
Pleuronectidae
Pseudopleuronectes americanus
Winter flounder
Primary
Sciaenidae
Cynoscion regalis
Gray weakfish
Primary
Sciaenops ocellatus
Red drum
Primary
Sparidae
Stenotomus chrysops
Scup
Primary
NORTHEAST REGION SECONDARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*
Achiridae
Trinectes maculatus
Hogchoaker
Anguill idae
Anguilla rostrata
American eel
Secondary
Atherinopsidae
Menidia menidia
Atlantic silverside
Batrachoididae
Opsanus tau
Oyster toadfish
Ephippidae
Chaetodip terus faber
Atlantic spadefish
Moronidae
Morone saxatilis
Rock fish (or striped bass)
Secondary
Mugulidae
Mugil cephalus
Black mullet
Pomatomidae
Pomatomus saltatrix
Bluefish
Secondary
Sciaenidae
Bairdiella chrysoura
Silver perch
Menticirrhus saxatilis
Northern kingfish
Serranidae
Centropristis striata
Black sea bass
Triglidae
Prionotus carolinus
Northern searobin
Prionotus evolans
Striped searobin
* Indicates whether species also occurs in the primary or secondary fish plus 'Jsf (see Table 14.8).
Table 14.3 Southeast region primary and secondary marine target species - whole body fish tissue collection
(Ecofish)
SOUTHEAST REGION PRIMARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*!
Ariidae
Ariopsisfelis
Hardhead sea catfish
Pr
imary
Bagre marinus
Gafftopsail sea catfish
Pr
imary
Paralichthyidae
Paralichthys albigutta
Gulf flounder
Pr
imary
Paralichthys dentatus
Summerflounder
Pr
imary
Paralichthys lethostigma
Southern flounder
Pr
imary
Sciaenidae
Cynoscion arenarius
Sand weakfish (or seatrout)
Pr
imary
Cynoscion nebulosus
Speckled trout
Pr
imary
Cynoscion regalis
Gray weakfish
Pr
imary
Leiostomus xanthurus
Spot croaker
Pr
imary
Sparidae
Lagodon rhomboides
Pinfish
SOUTHEAST REGION SECONDARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*!
Cichlidae
Tilapia mariae
Spotted tilapia
Haemulidae
Haemulon aurolineatum
Tomtate
Sciaenidae
Bairdiella chrysoura
Silver perch
Menticirrhus americanus
Southern kingfish
Serranidae
Centropristis striata
Black sea bass
* Indicates whether species also occurs in the primary or secondary fish plus 'Jsf (see Table 14.8).
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Table 14.4 Gulf region primary and secondary marine target species - whole body fish tissue collection (Ecofish)
GULF REGION PRIMARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*
Ariidae
Ariopsis felis
Hardhead sea catfish
Pr
mary
Bagre marinus
Gafftopsail sea catfish
Pr
mary
Paralichthys albigutta
Gulf flounder
Pr
mary
Paralichthyidae
Paralichthys dentatus
Summer flounder
Pr
mary
Paralichthys lethostigma
Southern flounder
Pr
mary
Cynoscion arenarius
Sand weakfish (or seatrout)
Pr
mary
Cynoscion nebulosus
Speckled trout
Pr
mary
Sciaenidae
Cynoscion regalis
Gray weakfish
Pr
mary
Leiostomus xanthurus
Spot croaker
Pr
mary
Micropogonias undulatus
Atlantic croaker
Pr
mary
Sciaenops ocellatus
Red drum
Pr
mary
Sparidae
Lagodon rhomboides
Pinfish
GULF REGION SECONDARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*
Carangidae
Caranx hippos
Crevalle jack
Chloroscombrus chrysurus
Atlantic bumper
Diodontidae
Chilomycterus schoepfii
Burrfish
Gerreidae
Eucinostomus gula
Silver jenny
Haemulidae
Orthopristis chrysoptera
Pigfish
Ictaluridae
Ictalurus furcatus
Blue catfish
Lepisosteidae
Lepisosteus oculatus
Spotted gar
Lutjanidae
Lutjanus griseus
Gray snapper
Sciaenidae
Pogonias cromis
Black drum
Serranidae
Diplectrum formosum
Sand perch
Triglidae
Prionotus scitulus
Leopard searobin
* Indicates whether species also occurs in the primary or secondary fish plug list (see Table 14.8).
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Table 14.5 Western region primary and secondary marine target species
(Ecofish)
whole body fish tissue collection
WESTERN REGION PRIMARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*
Atherinopsidae
Atherinops affinis
Topsmelt silverside
Cottidae
Leptocottus armatus
Pacific staghorn sculpin
Primary
Oligocottus rimensis
Saddleback sculpin
Cynoglossidae
Symphurus atricaudus
California tonguefish
Embiotocidae
Cymatogaster aggregata
Shiner perch
Primary
Embiotoca lateralis
Striped seaperch
Primary
Gasterosteidae
Gasterosteus aculeatus
Three-spined stickleback
Citharichthys sordidus
Pacific sanddab
Primary
Paralichthyidae
Citharichthys stigmaeus
Speckled sanddab
Primary
Paralichthys californicus
California flounder
Primary
Isopsetta isolepis
Butter sole
Pleuronectidae
Parophrys vetulus
English sole
Primary
Platichthys stellatus
Starry flounder
Primary
Psettichthys melanostictus
Pacific sand sole
Sciaenidae
Genyonemus lineatus
White croaker
Primary
Serranidae
Paralabrax nebulifer
Barred sand bass
Primary
Paralabrax maculatofasciatus
Spotted sand bass
Primary
WESTERN REGION SECONDARY ECOFISH TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
FISH PLUG LIST*
Batrachoididae
Porichthys notatus
Plainfin midshipman
Porichthys myriaster
Specklefin midshipman
Embiotocidae
Amphistichus argenteus
Barred surfperch
Secondary
Paralichthyidae
Xystreurys liolepis
Fantail sole
Hypsopsetta guttulata
Diamond turbot
Secondary
Pleuronectidae
Microstomus pacificus
Dover sole
Secondary
Lepidopsetta bilineata
Rock sole
Lyopsetta exilis
Slender sole
Sciaenidae
Umbrina roncador
Yellowfin croaker
' Indicates whether species also occurs in the primary or secondary fish plug list (see Table 14.8).
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Table 14.6 Great Lakes primary and secondary target species - whole body fish tissue collection (Ecofish)
GREAT LAKES PRIMARY ECOFISH TARGET SPECIES
FAMILY SCIENTIFIC NAME COMMON NAME FISH PLUG LIST*
Catostomidae
Moxostoma macrolepidotum
Shorthead redhorse
Primary
Centrarchidae
Ambloplites rupestris
Rock bass
Primary
Lepomis qibbosus
Pumpkinseed
Primary
Lepomis macrochirus
Bluegill
Primary
Micropterus dolomieu
Smallmouth bass
Primary
Pomoxis annularis
White crappie
Pomoxis nigromaculatus
Black crappie
Cottidae
Cottus bairdii
Mottled sculpin
Cottus co gnat us
Slimy sculpin
Cyprinidae
Couesius plumbeus
Lake chub
Cyprinus carpio
Common carp
Primary
Pimephales notatus
Bluntnose minnow
Esocidae
Esox lucius
Northern pike
Primary
Esox masguinongy
Muskellunge
Primary
Gadidae
Lota lota
Burbot
Primary
Gasterosteidae
Gasterosteus aculeatus
Three-spined stickleback
Gobiidae
Neogobius melanostomus
Round goby
Proterorhinus marmoratus
Tubenose goby
Ictaluridae
Ameiurus nebulosus
Brown bullhead
Primary
Ictalurus punctatus
Channel catfish
Primary
Noturus flay us
Stonecat
Moronidae
Morone americana
White perch
Primary
Morone chrysops
White bass
Primary
Osmeridae
Osmerus mordax
American/ rainbow smelt
Percidae
Gymnocephalus cernuus
Ruffe
Perca flavescens
Yellow perch
Primary
Percina caprodes
Logperch
Sander canadensis
Sauger
Sander vitreus
Walleye
Primary
Percopsidae
Percopsis omiscomaycus
Trout-perch
Salmonidae
Coregonus artedi
Cisco/ lake herring
Coregonus clupeaformis
Lake whitefish
Primary
Oncorhynchus gorbuscha
Pink salmon
Oncorhynchus kisutch
Coho salmon
Primary
Oncorhynchus mykiss
Rainbow trout
Primary
Oncorhynchus tshawytscha
Chinook salmon
Primary
Sa/ve/inus namaycush
Lake trout
Primary
Sciaenidae
Aplodinotus arunniens
Freshwater drum
Primarv
GREAT LAKES SECONDARY ECOFISH TARGET SPECIES
FAMILY SCIENTIFIC NAME COMMON NAME FISH PLUG LIST*
Catostomidae
Catostomus catostomus
Longnose sucker
Catostomus commersonii
White sucker
Secondary
Moxostoma anisurum
Silver redhorse
Centrarchidae
Micropterus salmoides
Largemouth bass
Clupeidae
Alosa pseudoharengus
Alewife
Dorosoma cepedianum
American gizzard shad
Cyprinidae
Cyprinella spiloptera
Spotfin shiner
Luxilus cornutus
Common shiner
Notropis stramineus
Sand shiner
Esocidae
Esox niger
Chain pickerel
Fundulidae
Fundulus diaphanus
Banded killifish
Fundulus maialis
Striped killifish
Ictaluridae
Ameiurus melas
Black bullhead
Salmonidae
Prosopium cylindraceum
Round whitefish
Salmo trutta
Brown trout
Secondary
Salvelin us fon tinalis
Brook trout
Salvelinusfontinalis x namaycush
Splake
* Indicates whether species also occurs in the primary or secondary fish plus 'Jsf (see Table 14.9).
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14.2 Fish Tissue Plug [FPLG]
14.2.1 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 at least 190 mm 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 (Section 14.1). 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) (if all required HTIS and FTIS specimens have been collected)
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 remove fish plugs from
specimens that are part of the human health fish composite 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 14.1; if a plug is collected from a live fish, antibiotic
salve is placed over the wound and the fish is released.
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14.2.2 Equipment and Supplies
Table 14.7 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 form in the App.
Table 14.7 Equipment ft 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 mL)
nitrile gloves
measuring board
aspirator bulb
scale (in grams)
scalpel (disposable, sterile)
For recording
NCCA App
measurements
fish tissue plug sample label
fine-tipped indelible markers (for labels)
clear tape strips
14.2.3 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
14.8 and Table 14.9. When ecofish specimens meet the size (190 mm) 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 by people in or around the study area, with specimens
that have a minimum length of 190 mm) to obtain a sample from the species that are
available. Alternative fish species should be limited to bony fish. Cartilaginous fish and
Moray eels (Family Muraenidae) should not be submitted for this indicator or for the
ecofish sample. The alternative genus and species must be written in to the NCCA App. 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 commonly consumed, 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.
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Please note: There are no invertebrate organisms on this list. Crab, shrimp, mollusks,
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 repellant 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,
• 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,
• at least 190 mm in length.
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). The minimum acceptable length for a fish used
for any fish plug sample is 190 mm.
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.
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11. Record sample ID, species, and specimen length and weight in the Fish
Tissue Plug Samples section of the Eco Fish Collection form in the App.
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 mm 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 mL 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 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 zip-top
bag. Place the sample in a cooler with dry ice
19. Dispose of gloves, scalpel, and biopsy punch.
14.2.4 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 one
week of collection. Please see Appendix C: Shipping and Tracking
Guidelines for next steps.
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Table 14.8 Primary and secondary marine target species for fish plug collection
PRIMARY MARINE FISH PLUG TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
Ariidae
Ariopsisfelis
Hardhead sea catfish
Bagre marinus
Gafftopsail sea catfish
Cottidae
Leptocottus armatus
Pacific staghorn sculpin
Embiotocidae
Cymatogaster aggregata
Shiner perch
Embiotoca lateralis
Striped seaperch
Ictaluridae
Ameiurus catus
White catfish
Ictalurus punctatus
Channel catfish
Moronidae
Morone americana
White perch
Citharichthys sordidus
Pacific sanddab
Citharichthys stigmaeus
Speckled sanddab
Paralichthys albigutta
Gulf flounder
Paralichthyidae
Paralichthys californicus
California flounder
Paralichthys dentatus
Summerflounder
Paralichthys lethostigma
Southern flounder
Parophrys vetulus
English sole
Platichthys stellatus
Starry flounder
Pleuronectidae
Pseudopleuronectes americanus
Winter flounder
Cynoscion arenarius
Sand weakfish (or seatrout)
Cynoscion nebulosus
Speckled trout
Cynoscion regalis
Gray weakfish
Sciaenidae
Genyonemus lineatus
White croaker
Leiostomus xanthurus
Spot croaker
Micropogonias undulatus
Atlantic croaker
Sciaenops ocellatus
Red drum
Serranidae
Paralabrax maculatofasciatus
Spotted sand bass
Paralabrax nebulifer
Barred sand bass
Sparidae
Stenotomus chrysops
Scup
SECONDARY MARINE FISH PLUG TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
Anguillidae
Anguilla rostrata
American eel
Amphistichus argenteus
Barred surfperch
Embiotocidae
Amphistichus rhodoterus
Redtail surfperch
Embiotoca jacksoni
Black perch
Hyperprosopon argenteum
Walleye surfperch
Moronidae
Morone saxatilis
Rock fish (or striped bass)
Paralichthyidae
Hippoglossina oblonga
Fourspot flounder
Hippoglossoides platessoides
American dab
Pleuronectidae
Hypsopsetta guttulata
Diamond turbot
Liman da ferru ginea
Yellowtail flounder
Microstomus pacificus
Dover sole
Pomatomidae
Pomatomus saltatrix
Blue fish
Sciaenidae
Menticirrhus undulatus
California whiting
Scorpaena guttata
California scorpionfish
Sebastes caurinus
Copper rockfish
Sebastes entomelas
Widow rockfish
Scorpaenidae
Sebastes fl avid us
Yellowtail rockfish
Sebastes melanops
Black rockfish
Sebastes mystinus
Blue rockfish
Sebastes paucispinis
Bocaccio
Serranidae
Paralabrax clathratus
Kelp bass
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Table 14.9 Primary and secondary Great Lakes target species for fish plug collection
PRIMARY GREAT LAKES FISH PLUG TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
Catostomidae
Moxostoma macrolepidotum
Shorthead redhorse
Ambloplites rupestris
Rock bass
Centrarchidae
Lepomis gibbosus
Pumpkinseed
Lepomis macrochirus
Bluegill
Micropterus dolomieu
Smallmouth bass
Cyprinidae
Cyprinus carpio
Common carp
Esocidae
Esox lucius
Northern pike
Esox masquinongy
Muskellunge
Ictaluridae
Ameiurus nebulosus
Brown bullhead
Ictalurus punctatus
Channel catfish
Gadidae
Lota lota
Burbot
Moronidae
Morone americana
White perch
Morone chrysops
White bass
Percidae
Percaflavescens
Yellow perch
Sander vitreus
Walleye
Coregonus clupeaformis
Lake whitefish
Oncorhynchus kisutch
Coho salmon
Salmonidae
Oncorhynchus mykiss
Rainbow trout
Oncorhynchus tshawytscha
Chinook salmon
Salvelinus namaycush
Lake trout
Sciaenidae
Aplodinotus grunniens
Freshwater drum
SECONDARY GREAT LAKES FISH PLUG TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
Catostomidae
Catostomus commersonii
White sucker
Ictaluridae
Ictalurus furcatus
Blue catfish
Salmonidae
Salmo trutta
Brown trout
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14.3 Human Health Fish Tissue Collection [HTIS] (GreatLakes
Nearshore and Lake Michigan Enhancement sites only)
14.3.1 Summary of Method
Field crews collect human health fish composite samples at all 225 of the Great Lakes
nearshore sites (i.e., sites whose prefix begins with NGL20), all 38 Great Lakes island sites
(sites whose prefix begins with ISA20), and all 12 Great Lakes National Park sites (sites
whose prefix begins with NPA20). This will result in human health fish tissue being
targeted at 45 sites per lake, plus the 38 island sites and 12 park sites in Lake Michigan. If
a Great Lake site has been designated as a human health fish tissue site and is dropped, a
replacement site is identified following procedures described in Section 2.3.2 and human
health fish tissue should be collected at the replacement site. At revisit sites in the Great
Lakes, crews that are unsuccessful at collecting the human health fish tissue sample
(HTIS) during visit 1 are expected to attempt the collection of that sample during visit 2.
Note that human health fish tissue samples will NOT be collected at Lake Erie (LEA20) and
Green Bay (GBA20) enhancement sites.
Labs analyze fillet tissue for mercury, polychlorinated biphenyls (PCBs), per- and
polyfluoroalkyl substances (PFAS), and fatty acids.
This section contains the sampling procedures and target species for human health fish
composite collection. Note that the human health fish species table (Table 14.11)
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 or Great Lakes Fish Tissue Trainer.
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., hook and line, gill net, or otter trawl). 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 form in the App.
For each attempted fish collection method, record equipment details, start and stop
times, and fishing location(s) on the Human Health Fish Collection form in the App.
Record sample ID, species retained, and specimen lengths on the Human Health Fish
Collection form in the App.
Identify and measure the specimens collected for each composite sample. Record the
scientific name (genus and species) and total length for each specimen on the Human
Health Fish Collection form in the App. Human health fish composites should consist of
five 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. The minimum
acceptable length for a fish in any composite sample is 190 mm. Field crews should make
every effort to consistently obtain five fish for the human health fish composite sample;
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however, a sample of fewer than five fish is acceptable. Conversely, for the exceptions
where field crews collect five fish that are small, they should collect up to five additional
fish (for an overall composite of up to 10 fish) to provide adequate tissue for analysis.
Fish submitted as part of the human health fish composite sample should remain intact
and be submitted as whole specimens. Crews should not take fish plugs from human
health fish tissue specimens.
14.3.2 Fish Tissue Distribution Scheme
Ideally, at Great Lakes sites where crews will collect both human health fish tissue
samples (HTIS) and ecological fish tissue samples (FTIS), they will successfully collect 10
or more fish of the same species that are each >190 mm in length. That would allow them
to retain 5 fish for the HTIS sample and 5 (or more) for the FTIS sample. However, if 10
fish are not available at a site, field teams will apply a fish distribution or "fish-splitting"
scheme. It is important to understand that the 5 HTIS fish must be the same species and
the 5 FTIS fish must be the same species, but the HTIS sample and the FTIS sample from a
site may be different species. (Note that the following fish distribution scheme would
only apply when the same species of fish is collected and available for both human health
and ecological samples).
If only a single fish is collected at a site, it should be retained as the Ecological (FTIS)
sample.
If sampling yields two fish of the same species, one will be the Human Health (HTIS)
sample and one will be the FTIS sample. If an odd number of fish of the same species are
collected at a site, the "extra" fish should be included in the HTIS sample. For example,
if sampling yields three fish of the same species, two of them will be saved as the HTIS
sample and one fish will be retained as the FTIS sample (See Figure 14.1). Obviously, in
cases where an even number of fish (of the same species) are collected from a site, the
number of specimens will be split evenly between the HTIS sample and the FTIS sample.
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Fish Tissue Sample Fish Distribution Scheme
Reminder: Apply this scheme only
when all fish are the same species
Human Health
(HTIS)
Ecological
(FTIS)
If ¦ ¦ ¦ 1 fish collected Then...
2 fish collected
3 fish collected
4 fish collected
5 fish collected
6 fish collected
7 fish collected
8 fish collected
9 fish collected
10 fish collected
4T
v-"' <¦'
/r/r
/W"
Figure 14.1 Fish Tissue Distribution Scheme to be used at all Great Lake Sites with the prefix NCL20, ISA20, or
NPA20.
14.3.3 Equipment and Supplies
Table 14.10 lists the equipment and supplies necessary for field crews to collect human
health fish composite 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.
Table 14.10 Equipment Et supplies: human health fish tissue collection
For collecting fish
composite sample
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)
For storing and
preserving fish
composite sample
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)
For documenting the
fish composite
sample
NCCA App
human health fish tissue sample labels
fine-tipped indelible markers (for labels)
Tyvek label tag with grommet
clear tape strips
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14.3.4 Sampling Procedure
Note: Do not handle any food, drink, sunscreen, or insect repellant 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 composite sample, select five whole fish. Criteria
for inclusion in the human health fish composite sample:
a) All fish are of the same primary target species or secondary fish species
(See Table 14.11)
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. The minimum
acceptable fish length is 190 mm.
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 form in the App:
• Ensure the sample identification number is entered.
• Check the boxes verifying that all samples are of similar length and the
same species.
• Record species selected for analysis, individual specimen lengths (total
length in mm), and any relevant comments. Extra rows are provided in the
App in the event that additional specimens are collected to ensure
adequate tissue for analysis (refer to Frequently Asked Questions for
further clarification).
• Make sure the sample ID and specimen numbers recorded in the App match
those on the sample labels.
6. Wearing clean nitrile gloves, remove each fish selected for analysis from the
clean holding container(s). If needed, dispatch each fish using the most
humane method available.
7. Wrap each whole fish in extra heavy-duty aluminum foil, with the dull side in
contact with the fish (foil is solvent rinsed and baked and will be provided by
EPA).
8. Prepare a sample label for each sample specimen, ensuring that the label
information matches the information recorded on the Human Health Fish
Collection form in the App. Be sure to record the fish genus and species and
specimen length on each label.
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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 form in the App. Be sure to record fish genus and 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.
14.3.5 Sample Storage and Shipping Preparation
1. After the fish sample is packaged, keep the sample chilled using either of the
following options (option "a" preferred):
a) (preferred option) immediately place the fish sample in a cooler of dry ice
until it can be properly frozen (at <-20°C in a laboratory or other interim
facility) or shipped to Microbac Laboratories (Baltimore, MD);
• If fish samples are held on dry ice in the field, the field crew should
replenish the supply of dry ice at least daily until the samples can be
properly frozen or shipped.
• Keep all specimens designated for a particular fish composite sample in
the same cooler for transport.
b) (alternate option for temporary holding or transport) immediately place
the fish sample in a cooler with wet ice (for temporary holding only).
• Packaged fish samples may be placed on wet ice in coolers if they will
be immediately transported to a nearby laboratory or other interim
facility to be frozen before shipment (wet ice should be replenished
frequently before it melts).
• Keep all specimens designated for a particular fish composite sample in
the same cooler for transport.
2. Crews have two options for freezing and shipping fish composite samples,
depending on site logistics:
a) Ship the samples via priority overnight delivery service (i.e., Federal
Express), packed on dry ice, so that they arrive at Microbac Laboratories
(Baltimore, MD) within 24 hours from the time of sample collection. Do NOT
ship on Fridays, Saturdays, or the day before federal holidays. Fish samples
must be packed on sufficient dry ice (50 pounds minimum, with blocks of
dry ice layered to ensure direct contact between fish and dry ice) to keep
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them frozen for up to 48 hours. Do not use dry ice pellets for shipping
human health fish samples. Remember to record the tracking number on
the Tracking Form in the App before submitting it to NARS IM.
b) Freeze the fish samples within 24 hours of collection at <-20° C and store
the frozen samples until shipment within two weeks of sample collection._|f
fish samples cannot be stored in a freezer within 24 hours of collection, the
field crew should replenish the supply of dry ice in the cooler containing
the samples, at least daily, until the samples can be properly frozen or
shipped. Frozen fish samples will subsequently be packed on at least 50
pounds of layered blocks of dry ice and shipped to Microbac Laboratories
(Baltimore. MD) 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
App tracking forms.
Table 14.11 Primary and secondary Great Lakes target species for human health fish tissue collection
PRIMARY HUMAN HEALTH FISH TISSUE TARGET SPECIES
FAMILY
SCIENTIFIC NAME
COMMON NAME
Ambloplites rupestris
Rock bass
Micropterus dolomieu
Smallmouth bass
Centrarchidae
Micropterus salmoides
Largemouth bass
Pomoxis annularis
White crappie
Pomoxis nigromaculatus
Black crappie
Cyprinidae
Cyprinus carpio
Common carp
Esox lucius
Northern pike
Esocidae
Esox masquinongy
Muskellunge
Esox niger
Chain pickerel
Ictaluridae
Ictalurus punctatus
Channel catfish
Gadidae
Lota lota
Burbot
Moronidae
Morone americana
White perch
Morone chrysops
White bass
Percaflavescens
Yellow perch
Percidae
Sander canadensis
Sauger
Sander vitreus
Walleye
Coregonus clupeaformis
Lake whitefish
Oncorhynchus gorbuscha
Pink salmon
Oncorhynchus kisutch
Coho salmon
Salmonidae
Oncorhynchus tshawytscha
Chinook salmon
Oncorhynchus mykiss
Rainbow trout
Salmo salar
Atlantic salmon
Salmo trutta
Brown trout
Sa/ve/inus namaycush
Lake trout
Sciaenidae
Aplodinotus grunniens
Freshwater drum
SECONDARY HUMAN HEALTH FISH TISSUE TARGET SPECIES
FAMILY SCIENTIFIC NAME COMMON NAME
Carpiodes cyprinus
Quillback
Catostomus catostomus
Longnose sucker
Catostomus commersonii
White sucker
Catostomidae
Hypentelium nigracans
Northern hogsucker
Ictiobus cyprinellus
Bigmouth buffalo
Ictiobus niger
Black buffalo
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| Lepomis cyanellus
| Green Sunfish
I Lepomis gibbosus
I Pumpkinseed
Centrarchidae
I Lepomis gulosus
I Warmouth
| Lepomis macrochirus
i Bluegill
I Lepomis megalotis
| Longear Sunfish
| Ameiurus melas
i Black bullhead
Ictaluridae
| Ameiurus natalis
| Yellow bullhead
I Ameiurus nebulosus
| Brown bullhead
I Coregonus artedi
I Cisco/ lake herring
Salmonidae
I Coregonus hoyi
i Bloater
I Prosopium cylindraceum
I Round whitefish
I Salvelinusfontinalis
I Brook trout
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15 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 15.1.
Figure 15.1 Final site activities summary
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15.1 General Site Assessment
After sampling, complete the Site Assessment form in the App. Record all observations
from the site that were noted during the course of the visit. The Site Assessment form is
by no means comprehensive, and crews are encouraged to record any additional pertinent
observations in the General Assessment from in the App.
15.1.1 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 of the App
to verify that blank fields indicate absence of the specific type of activity or disturbance.
15.1.2 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. For example, 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 the dominant land use
around the X-site. 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.
15.1.3 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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15.2 Processing The Fecal Indicator
15.2.1 Summary of Method
At each site, crews collect and filter water samples for fecal indicator analyses. Upon
receipt of the filters, the lab uses qPCR analysis to quantify Enterococci bacteria trapped
on the filter.
15.2.2 Equipment and Supplies
Table 15.1 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 subsequent filtering of the
chlorophyll-a sample at the same site, Enterococci must be filtered before filtering
chlorophyll-a samples.
Table 15.1 Equipment ft supplies: Enterococci processing
For processing samples
nitrile gloves
sterile screw-cap graduated 50 mL centrifuge tube (for measuring sample)
Filtration flask (side arm, 500 mL)
rubber stopper (#8 white, with 10 mm 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
measurements
NCCA App
fine-tipped indelible markers (for labels)
fecal indicator sample labels (2 or 3 vial labels and 1 bag label)
clear tape strips
15.2.3 Processing Procedure - Fecal Indicator Filter Blank
At revisit sites (sites that will be visited twice in the index period for QA 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 15.2). Set out:
a. 50 mL sterile centrifuge tube,
b. One bottle of chilled phosphate buffer solution (PBS), and
c. Two 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 seven 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 four 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 Form in the App.
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 15.2.4 for processing the water sample collected for
Enterococci.
15.2.4 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
six hours of collection. See below for field processing procedures.
Prior to beginning the filtering process, chill the Enterococci sample and PBS on wet ice
for at least 15 minutes and chill the microcentrifuge tubes on dry ice.
1. Put on nitrile gloves.
2. Set up the sample filtration apparatus on a flat surface and attach the vacuum
pump (Figure 15.2). Set out:
a. 50 mL sterile centrifuge tube,
b. one bottle of chilled PBS, and
c. two sterile forceps.
3. Attach the filter funnel with pre-loaded sterile filter onto the filtering flask with
reusable rubber stopper and adapter.
4. Gently 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 seven 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
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25 mL volume, discard the filter and, using a new sterile filter funnel with pre-
loaded filter, repeat the filtration using a lesser volume.
8. 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 five times. Remove
the cap and pour the rinse into the filter funnel to rinse the filter.
9. Filter the rinsate and repeat with another 10 mL of chilled PBS.
10. 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 four
times).
11. 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 form in the App. 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 form in the App.
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 form in the App.
Affix the exterior label on the outside of the bubble envelope and cover with clear
plastic tape.
18. Place the bubble envelope in the zip-top bag and then on dry ice for preservation
during transport and shipping.
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Sterile 100 mL .
filter funnel with
pre-loaded filter
(single use)
Reus
filterir
funne
Rubber stopper (white)
and small funnel adapter
Vacuum pump
(hand or electric)
m
Figure 15.2 Filtering set-up for Enterococci filtering
15.3 Processing The Chlorophyll-/! & Dissolved Nutrients Indicators
15.3.1 Summary of Method
At each site, crews collect and filter water samples for chlorophyll-a arid 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.
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15.3.2 Equipment and Supplies
Table 15.2 Equipment ft supplies: chlorophyll-a ft dissolved nutrients processing
For filtering
chlorophyll-a
sample
Filters - Whatman 47 mm glass fiber GF/F 0.7 |j, - Box
Nutrients filtering chamber
Silicone grease
Filtration unit (blue base filter funnel, 250 mL unit)
rubber stopper (#8 blue, with 15 mm hole) and large filter funnel adapter
vacuum pump (electric or hand)
Dl water
nitrile gloves
forceps
graduated cylinder (250 mL)
For recording
measurements
NCCA App
chlorophyll-^ & dissolved nutrients sample labels
fine-tipped indelible markers (for labels)
clear tape strips
For sample
collection and
preservation
Centrifuge tube (sterile, screw-top, 50-mL) in leak-proof bag
aluminum foil square
FIDPE bottle (250 mL, white)
cooler with dry ice
electrical tape
15.3.3 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.
1. Complete the NUTS sample label with Site ID, date collected, and visit number.
2. Attach the completed label to the 250 mL clear HDPE 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 15.3).
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 with Dl water
between sampling events. Prior to filtration of the sample, rinse the filter funnel
adapter and graduated cylinders three times with Dl water. After assembling the
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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 seven inches of Hg of vacuum -3.4 psig or
a filtration duration of more than five minutes for a single sample volume, to avoid
cell damage or loss of contents during filtering.
a. If the lid of the filtration chamber does not seal adequately, apply a small
amount of silicone grease to the gasket on the underside of the lid.
b. Applying downward pressure to the lid during initial application of vacuum will
also help the lid seal.
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 and
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 HDPE 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.
14. Record the dissolved nutrients sample information on the Sample Collection form in
the App. 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 form in the App 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.
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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 !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 zip-top plastic bag.
21. Close the plastic bag and place it on dry ice.
i. 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.
filter funnel -
(250 mL with blue base)
Figure 15.3 Filtering set-up for chlorophyll-a and nutrients filtering
15.4 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 of the Calibration/QA form in the App. If
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significant drift is detected as defined by 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.
15.5 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 fully completed.
• Confirm that the Site ID, visit number, and date of visit are correct.
• Ensure that the water chemistry sample ID has been entered on the
Tracking Form in the App.
• Verify the accuracy and clarity of all recorded information.
• Ensure that any pertinent data explanations are entered into the respective
comments sections.
• Ensure that comments are clear, with no "shorthand" or abbreviations.
• Make sure that any targeted sample that was not collected has a comment
recorded as to why the sample was not collected.
• Ensure that shipping/airbill tracking numbers have been recorded in the
Tracking Form in the App prior to shipping samples.
If information is missing from the forms, the Field Crew Leader must complete the missing
sections. When utilizing the NCCA App, the Field Crew Leader must ensure that the data is
submitted. The receipt of a submission is a confirmation that the data has been reviewed
by the Field Crew Leader.
15.6 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 labels are preprinted with the sample IDs, review the labels
and forms in the App 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.
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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 in
the NCCA App and labels before tracking forms are transmitted to NARS IM and samples
are packaged and sent to the labs.
15.7 Sample Shipment & Tracking Form Submittal
Each shipping group has been assigned a "T" number to help crews identify the correct
section of the Tracking Form in the App to use when sending samples. This "T" number is
located above each of the sample groups in the Tracking Form in the App. Crews will also
find reference to the same "T" numbers on the individual samples labels, on the packing
slips that crews will include in the coolers, and on the top of the pre-printed FedEx return
labels provided in the site kits.
Crews submit tracking information via the NCCA App and include packing slips in the
coolers when they send samples to the labs. Refer to Appendix C: Shipping and Tracking
Guidelines for additional details on preparing samples for shipping.
15.7.1 Time-Sensitive Samples
The field crew must ship or deliver time-sensitive samples (i.e., water chemistry (CHEM),
chlorophyll-a (WCHL), and dissolved nutrients (NUTS)) to the appropriate analytical
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 or start shipments on Saturday or 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 or his/her designee will complete the T-1 section of the Tracking
Form in the App for the samples being shipped. Shipping details which include the
destination lab, date shipped, FedEx airbill number, sender, and sender's phone number
will be entered for the group of samples. Once details are saved in the App, a date will
appear in the shipped column of the Tracking Form.
The Field Crew Leader or his/her designee will submit the Tracking Form via the NCCA
App along with any and all data forms. This initial submission serves as both the
notification of a sampling event and the tracking of the designated samples. After
submission, a data summary will be automatically emailed back to the email address from
which the submission was received. The Field Crew Leader or his/her designee should
review this data summary for accuracy and make any corrections necessary and re-submit
the pertinent form(s).
The field crew will place the samples and the appropriate packing slip (in a waterproof
bag or plastic sleeve) in the cooler provided with the site kit. The field crew 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
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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. If there are samples listed on the packing that are not included in the cooler, line
out the sample IDs for the samples not included.
15.7.2 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 16: Post-Sampling Activities
for further guidance.
15.8 Equipment Cleanup & Check
After each sampling event, crews will need to clean all sampling gear using the guidance
provided in Table 15.3. These steps are for general cleaning and do not include any
additional steps necessary to decontaminate equipment for the known or suspected
presence of nuisance species.
Table 15.3 General cleaning of sampling gear after each site
1 A|iiipnicn( lype
Cleaning Method
Water collection and filtering equipment
Rinse 3 times with D1 water (110 detergent)
Sediment collection/processing equipment
Wash with a phosphate-free detergent such as Liquinox,
rinse with DI water
Sieve box/bucket
Use copious amounts of forceful water and a stiff brush to
clean the sieve. Be sure to rinse the brush between each
sieve cleaning.
Fish collection/processing gear
Clean with 1% bleach solution, rinse with tap water
and/or DI water
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 (VHS) is an
invasive and deadly fish virus that is threatening Great Lakes fish. VHS 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 15.8.3.
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)
• Sea Grant Program (http://www.sgnis.org)
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• USDA Animal and Plant Health Inspection Service (http://aphis.usda.gov)
15.8.1 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.
15.8.2 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.
15.8.3 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 five
minutes.
2. Rinse with tap water (do not use sea or lake water) and remove remaining
debris.
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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 four hours.
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
one minute.
3. Dry gear in direct sunlight (at least 84 °F) for at least four 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 and filter 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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16 Post-Sampling Activities
16.1 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 or his/her
designee will complete the pertinent section(s) (e.g., T-2, T-3, T-4, and/or T-5) of the
Tracking Form in the App for the samples being shipped. Shipping details which include
the destination lab, date shipped, FedEx airbill number, sender, and sender's phone
number will be entered for the group of samples. Once details are saved in the App, a
date will appear in the shipped column of the Tracking Form.
The Field Crew Leader or his/her designee will submit the Tracking Form via the NCCA
App. After submission, a data summary will be automatically emailed back to the email
address from which the submission was received. The Field Crew Leader or his/her
designee should review this data summary for accuracy and make any corrections
necessary and re-submit the pertinent form(s).
The Field Crew Leader will place the samples and the correct batch packing slip (in a
waterproof bag or plastic sleeve) in a requested batch shipment cooler. If there are
samples listed on the packing that are not included in the cooler, line out the sample
IDs for the samples not included. 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.
16.2 Data Submittal
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 menu button and choose the form(s)
that you wish to submit. Click the green submit button at the bottom of the form list. An
email will pop up on your device addressed to NARSFieldData@epa.gov. Copy yourself, any
other crew members or managers and click send. To ensure that he email was sent, check
the SENT mailbox on your email App and look for the recent email containing the data. If
the email is not in the SENT mailbox, it was not sent and you should try again after
verifying an internet connection.
At any point, if it is determined that data needs to be revised or updated, crews should
feel free to do so in the App and re-submit any edited data forms using the steps above.
Newly revised data will automatically take the place of previous data. It is not necessary
to re-submit data forms that were unchanged however.
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16.3 Data and Tracking Reminders
It is very important to submit the data and tracking forms immediately after every
sampling event. Prompt submissions 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 2020 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 FLC contact information can be
found on Table 1.1 of this manual.
16.4 Site Evaluation Spreadsheet Submittal
Throughout the field season or at the end of the field season, EPA HQ needs field crews to
submit their updated 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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17 Field Quality Control
Page 117
The NCCA program requires that all cooperators and field crews follow strict QA and QC
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.
17.1 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.
17.2 Standardized Field Data Collection App
All field crews collect and record data using an app. The app serves several purposes.
First, it ensures that crews measure and record the same parameters. Second, it promotes
efficient data entry and minimizes the opportunities for data transcription errors. Finally,
the app facilitates field data 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.
17.3 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 7% 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
eco fish and fish plug samples, which are targeted only on the first visit). At Great Lakes
revisit sites, crews that are unsuccessful at collecting the human health fish composite
sample (HTIS) 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 must always collect the filter
blanks before the sample is filtered. See Section 15.2.3 for the procedure for collecting
filter blanks.
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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.
17.4 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.
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17.4.1 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 17.1 summarizes the plan, the checklist, and corrective action
procedures.
Table 17.1 General information noted during field evaluation
Field
Evaluation
Plan
• 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.
Field
Evaluation
and
Assistance
Visit
Checklist
• 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.
Corrective
Action
Procedures
• 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.
17.4.2 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
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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
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:
• Brian Hasty, EPA Field Logistics Coordinator
The Field Logistics Coordinator will contact the NCCA QA Lead and the NCCA Project Lead
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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18 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 6t Health Administration (OSHA). 2006. Regulations (Standards - 29 CFR).
Substance technical guidelines for formalin - 1910.1048 App A. Occupational Safety 6t
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-
1.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. 2020. 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 2020: Site Evaluation Guidelines (EPA-841-R-14-
006)
National Coastal Condition Assessment 2020: Field Operations Manual (EPA-841-R-14-
007)
National Coastal Condition Assessment 2020: 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 HDPE 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.
Kit Type Item Quantity Protocol(s)
Regular
Aluminum foil squares - pack of 25
2
Chlorophyll A
Regular
Antibiotic Salve
1
Fish plug
Regular
Aspirator bulb
1
Fish Plug
Regular
Centrifuge tube stand
1
Chlorophyll A
Regular
Centrifuge tubes (screw-top, 50-mL) (extras)
10
Enterococci, Chlorophyll A
Regular
Clear tape strips - packs of 25
6
General
Regular
Electrical tape, plastic - roll
4
General
Regular
FedEx airbills (non-chilled batch, frozen batch, data)
10
Shipping
Regular
FedEx Dangerous Goods label (Class 9, for dry ice
shipments)
10
Shipping
Regular
Filters - Whatman 47 mm glass fiber GF/F 0.7 ju, - Box
1
Chlorophyll A
Regular
Filtration flask (side arm, 500 mL)
1
Chlorophyll A, Dissolved
Nutrients
Regular
Filtration unit (Sterile blue base 250 mL with funnel, cap, and
filter holder) - spares
5
Chlorophyll A, Dissolved
Nutrients
Regular
Filtration unit (white base, sterile, 100 mL units, includes pre-
loaded filter for ENTE) - spares
5
Enterococci
Regular
Filtration unit adapter (large)
3
Chlorophyll A
Regular
Filtration unit adapter (small)
3
Enterococci
Regular
Fish weigh scale, case, and spare batteries-]"
1
Fish plug
Regular
Forceps (fine-tipped, watchmakers types)
1
Benthics
Regular
Forceps (sterile, disposable) - spares
Enterococci, Chlorophyll A
Regular
Funnel (wide-mouth)
1
Benthics
Regular
Gloves (nitrile) - box
1
General
Regular
Graduated cylinder (100 mL)
1
Benthics
Regular
Graduated cylinder (250 mL)
1
Chlorophyll A
Regular
F1DPE bottle (1 L, white, wide-mouth) (extras)
Benthics
Regular
F1DPE bottle (2 L, amber)"]"
1
Chlorophyll A
Regular
Lowering line (100') with clips (marked in 0.5 m intervals)"]"
1
Depth Secchi
GL Only
Lugol's Solution
1
Phytoplankton
Regular
Microcentrifuge tubes containing glass beads (extras or for
filter blanks)
5
Enterococci
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1 Kit Type
Item
Quantity
Protocol(s)
Regular
Nutrients filtering chamber]"
1
Dissolved Nutrients
Regular
Packing tape (extra rolls)
2
Shipping
Regular
Packing tape (on holder)
1
Shipping
Regular
PAR Meter with frame, sensors and LI-1400 dataloger
1
Water Profile
Marine Only
Peristaltic pump with flexible gas-impermeable tubing
installed
1
Total Alkalinity
GL Only
Pipet (10 mL)
Phytoplankton
GL Only
Pipet Bulb
1
Phytoplankton
Regular
Plastic cable ties - spares
10
Eco Fish Tissue
Regular
Rubber bands (spares)
20
Sediment collection
Regular
Rubber stopper (#8 blue, with 15 mm hole)
2
Chlorophyll A, Dissolved
Nutrients
Regular
Rubber stopper (#8 white, with 10 mm hole)
2
Enterococci
Regular
Rubbermaid Roughneck tote (3 gallon)
1
General
Regular
Secchi disk (20 cm diameter, weighted)"]"
1
Water Profile
Regular
Shipping supplies organizer
1
Shipping
Regular
Sieve bucket (500 |xm)"]-
1
Benthics
Regular
Silicone grease
1
General
Regular
Sodium thiosulfate tablets (in vial)
1
Enterococci
Regular
Spoon, stainless steel (15")
1
Sediment collection
Regular
Tyvek tag with grommet - spares
10
Eco Fish Tissue
GL Only
Underwater video camera kit (includes frame, 2 cameras, 2
lights, lowering line with float, tools, Micro SD cards, etc.)
1
Underwater Video
Regular
Vacuum hand pump and clear plastic tubing]"
1
Enterococci, Chlorophyll A,
Dissolved Nutrients
Regular
Wash bottle (1 L Nalgene), One for ambient water, One for
DI
2
Sediment collection,
Chlorophyll A, Dissolved
Nutrients
Marine Only
Weight, stainless steel pipe, for TA intake tube
1
Total Alkalinity
Regular
Zip-top bags (2 gallon for ecoflsh) - extras
10
Eco Fish Tissue
Regular
Zip-top bags (sandwich size) - for ecoflsh labels - extras
10
Eco Fish Tissue
+ Item supplied if needed
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Site Kits
A site kit will be provided to the field crews for each sampling site. Site kits are specific to
marine sites and Great Lakes sites. Please submit an electronic request form well in advance
of field sampling. Kits must be requested at least two weeks before sampling is to take place.
Each site kit will include a label packet (specific to marine or Great Lakes sites) and 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.
Marine Site Kit
Item Quantity per Protocol(s)
Site Kit
Bucket w/screw top lid (0.6 gallon) for marine SEDX
1
Sediment Toxicity (Marine)
Centrifuge tube (sterile, screw-top, 50-mL) in leak-proof bag
1
Enterococci, Chlorophyll A
Cooler Liner (batch size, GLEQ
1
Shipping
Cooler liner (medium size, WRS)
1
Shipping
FedEx air bills (pre-addressed) plus handle tags, zip ties, etc.
1
Shipping
Filtration unit (blue base, 250 mL with funnel, cap, and filter holder)
1
Chi-A, Dissolved Nutrients
Filtration unit (white base, 100 mL, with pre-loaded ENTE filter)
Enterococci
Fish Tissue Plug Kit (includes vial, scalpel, punch, forceps, gloves,
bubble bag, and zip-top bag)
1
Fish Tissue Plugs
Forceps (sterile, disposable)
Enterococci, Chlorophyll A
Glass jar (120 mL, amber)
1
Sediment Organics/Metals
Glass jar (60 mL, amber)
1
Sediment TOC
Glass jar (60 mL, amber)
1
D15N
F1DPE bottle (1 L, white, wide-mouth)
1
Benthics
F1DPE bottle (125 mL, white, rectangular)
Total Alkalinity
F1DPE bottle (250 mL, amber)
1
Water Chemistry
F1DPE bottle (250 mL, white, round)
1
Dissolved Nutrients
F1DPE bottle (500 mL, white, round, wide mouth)
1
Algal Toxin
In-line disposable groundwater filter (0.45 |xm)
1
Total Alkalinity
Label and Packing slip packet
1
General
Microcentrifuge tubes with glass beads (in bubble and zip-top bag)
Enterococci
PETG bottle (250 mL, clear, square, pre-sterilized)
1
Enterococci
PETG bottle (500 mL, clear, square)
1
Microcystin
Plastic 6 mil bags (1 qt)
Sediment grain size
Plastic bag (large, composite)
1
Eco Fish Tissue
Plastic cable tie
1
Eco Fish Tissue
Sterile phosphate buffered solution (PBS)
1
Enterococci
Tyvek Tags w/Grommet
1
Eco Fish Tissue
Zip-top bags (plastic, 2 gallon)
2
Eco Fish Tissue
Zip-top bags (sandwich size) — for labels
2
Eco Fish Tissue
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Great Lakes Site Kit
Item
Quantity per
Site Kit
Protocol(s)
Bucket w/snap top lid (1 quart) for Great Lakes SEDX
1
Sediment Toxicity (GL)
Centrifuge tube (sterile, screw-top, 50-mL) in leak-proof bag
1
Enterococci, Chlorophyll A
Cooler Liner (batch size, GLEQ
1
Shipping
Cooler liner (medium size, WRS)
1
Shipping
FedEx air bills (pre-addressed) plus handle tags, zip ties, etc.
1
Shipping
Filtration unit (blue base, 250 mL with funnel, cap, and filter holder)
1
Chi-A, Dissolved Nutrients
Filtration unit (white base, 100 mL, with pre-loaded ENTE filter)
Enterococci
Fish Tissue Plug Kit (includes vial, scalpel, punch, forceps, gloves,
bubble bag, and zip-top bag)
1
Fish Tissue Plugs
Forceps (sterile, disposable)
Enterococci, Chlorophyll A
Glass jar (120 mL, amber)
1
Sediment Organics/Metals
Glass jar (60 mL, amber)
1
Sediment TOC
Glass jar (60 mL, amber)
1
D15N
F1DPE bottle (1 L, white, narrow mouth)
1
Phytoplankton
F1DPE bottle (1 L, white, wide-mouth)
1
Benthics
F1DPE bottle (250 mL, amber)
1
Water Chemistry
F1DPE bottle (250 mL, white, round)
1
Dissolved Nutrients
F1DPE bottle (500 mL, white, round, wide mouth)
1
Algal Toxin
Label and Packing slip packet
1
General
Microcentrifuge tubes with glass beads (in bubble and zip-top bag)
Enterococci
PETG bottle (250 mL, clear, square, pre-sterilized)
1
Enterococci
PETG bottle (500 mL, clear, square)
1
Microcystin
Plastic 6 mil bags (1 qt)
Sediment grain size
Plastic bag (large, composite)
1
Eco Fish Tissue
Plastic cable tie
1
Eco Fish Tissue
Sterile phosphate buffered solution (PBS)
1
Enterococci
Tyvek Tags w/Grommet
1
Eco Fish Tissue
Zip-top bags (plastic, 2 gallon)
2
Eco Fish Tissue
Zip-top bags (sandwich size) — for labels
2
Eco Fish Tissue
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Human Health Fish Tissue Sampling Kit
A human health fish tissue (HTIS) kit will be provided to the field crews for selected
sampling sites (separately from site kits). This kit will include materials for sampling HTIS at
one Great Lakes nearshore site. Please submit an electronic request form well in advance of
field sampling. Kits must be requested at least two 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
Aluminum foil
5
Packaging
H
a
w
Cooler (blue)
1
Storage & Shipping
Dry ice (Class 9) shipping label
1
Shipping
p
cn
FedEx airbill (pre-addressed)
1
Shipping
HH
H
Nitrile gloves
5 pairs
Packaging
X
(A
Plastic bags (large, composite)
1
Packaging
ft
X
X
Plastic cable ties
12
Packaging
Polyethylene tubing (heavy-duty, food grade)
1 roll
Packaging
Tyvek tags with grommets
1
Packaging
Crew Supplied Equipment
Item
Quantity
Protocol
Active/passive fish sampling device (e.g., trawl, seine, hook & line, etc.)
Fish Collection
Phosphate-free detergent such as Liquinox
Sediment Collection
Barometer (for calibration)
Water Profile
Batteries (AA)
GPS, Water Profile
Bleach (1-10% solution)
Decontamination
Borax
Sediment Collection
Buckets (large)
Sediment Collection
H
/
Calibration cups & standards
Profile
W
§
Cell phone, 2-way radios, walkie talkies
General
ft
HH
Clipboard(s)
1-2
General
P
a
De-ionized water (lab certified preferred, not required)
Water Profile
w
Digital camera (with extra memory card & batteries)
General
e!
Dip net
1
Fish Collection
w
/
Dry ice
~50 lbs/site
Shipping
w
a
Fine-tipped, indelible markers
General
Formalin (100% buffered) with stain
Sediment Collection
Fuses (10 amp)
Underwater Video
GPS unit (with manual & reference card, extra battery pack);
General
Knife
General
Livewell/buckets with aerator
Fish Collection
Maps & access instructions
General
Measuring board (mm scale)
1
Fish Collection
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Item Quantity Protocol
Multi-parameter probe water quality meter (with pH, DO, temperature,
and conductivity/salinity probes — e.g., Hydrolab, YSI, etc.)
1
Water Profile
NCCA 2020 Fact Sheets (available on NARS SharePoint)
10
General, Outreach
PAR meter (with LI-190 Quantum Sensor and LI-192 Underwater
1
Water Profile
Quantum Sensor & cables, independent datalogger)
Pencils (#2)
5
General
Plastic tub or bucket
1
Sediment Collection
QCS — quality check solution
If needed
Water Profile
Rose Bengal stain
1 bottle
Sediment Collection
Ruler (in cm)
1
General
Sampling permits/permission letters
General
Side cutter
1
Ecofish, Human
health fish
collection
Scissors
1
General
Scrub brush
1
Sediment Collection
Sieve box/frame (if necessary)
1
Sediment Collection
Spare parts
Various
Multi-probe
Stainless steel or Teflon spoons (large & small), spatulas, & scoops
Mixing and
dispensing sediment
Stainless steel mixing pot or bowl with lid
1
Sediment Collection
Stop watch
1
Underwater Video
Thermometer
1
Water Profile
Water sampling device (e.g., Niskin) or pump system
1
Chlorophyll A
Dissolved Nutrients
Phytoplankton
Water Chemistry
Microcystin
Weights & pads for grabs
Sediment Collection
Wet ice
-50
lbs/site,
additional
for shipping
Shipping
Young-modified Van Veen grab sampler (0.04 m2) OR standard OR
1
Sediment Collection
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)
H
/
W
§
Boat horn
Bow/Stern lights
Emergency tool kit
Ph
HH
D
Extra boat plug
O
a
§
Fire extinguisher
First aid kit
Float (to attach to anchor)
Gas Can
Fland bilge pump
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Item Quantity Protocol
Motor
PFDs (1/person)
Pingers
Sonar unit
Spare prop
Spare prop shear pin
Type IV PFD (throwable life saving device)
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Appendix B: Sample Labels & Packing Slips
Sample Labels (Marine)
**MARINE**
WATER CHEMISTRY (CHEM)
Site ID:
WATER COLUMN CHLOROPHYLL (WCHL)
Site ID:
Date:
/ /2020 Visit#: Ol 02
Date: / /2020 Visit#: Ol 02
T1
999000
Volume Filtered: ml.
T1 999001
Date:
NUTRIENTS (NUTS)
Site ID:
/ /2020 Visit#: Ol 02
ALGAL TOXIN (MICX)
(PETG Bottle (clear, square bottle)
Site ID:
T1
999002
Date: / /2020 Visit#: Ol 02
Salinity: (%>)
T3 999003
ALGAL TOXIN (MICZ)
(HDPE Bottle (round Nalgene bottle)
Site ID:
BENTHIC INFAUNA (BENT)
Site ID:
Date: / /2020 Visit#: Ol 02
Date:
/ /2020 Visit#: Ol 02
Salinity: (%o) Jar 1 of
T3
Salinity: (%o)
999004
T4 999005
SEDIMENT TOC(SEDC)
Site ID:
SEDIMENT GRAIN SIZE (SEDG)
Site ID:
Date:
/ /2020 Visit#: Ol 02
Date: / /2020 Visit #: Ol 02
T3
999006
999007
T2
SEDIMENT ORGAN ICS/METAL (SEDO)
Site ID:
SEDIMENT TOXICITY (SEDX)
Site ID:
Date:
/ /2020 Visit#: Ol 02
Date: / /2020 Visit #: Ol 02
T3
999008
T2 999009
SEDIMENT NITROGEN (D15N)
Site ID:
TOTAL ALKALINITY (ALKT)
Site ID:
Date:
/ /2020 Visit#: Ol 02
Date: / /2020 Visit#: Ol 02
T3
999010
Salinity: (%o) Jar 1 of 2
T2 999011
TOTAL ALKALINITY (ALKT)
Site ID:
Sample Type:
Site ID:
Date:
/ /2020 Visit#: Ol 02
Date: / /2020 Visit#: Ol 02
Salinity: (%o) Jar 2 of 2
Sample ID:
T2
999011
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FISH TISSUE PLUG (FPLG)
Site ID:
Date: / /2020 Visit #: Ol 02
999012
ECO FISH TISSUE - INNER BAG OF .
Site ID:
Date: / /2020 Visit ft: Ol 02
Genus Species:
Length (mm) Min.: Max.:
999013
ECO FISH TISSUE - INNER BAG OF .
Site ID:
Date: / /2020 Visit #: Ol 02
Genus Species:
Length (mm) Min.: Max.:
999013
ECO FISH TISSUE - OUTER BAG
T5 Site ID:
Date: / /2020 Visit #: Ol 02
Genus Species:
Length (mm) Min.: Max.:
999013
ENTEROCOCCI (ENTE) - BAG
Site ID:
Date: / /2020 Visit #: 01 02
Vol. Filt: 1 mL 2 mL
999014
Filter: 1
Vol. Filt: mL
999014
Filter: 2
Vol. Filt: mL
999014
Filter: Blank
Vol. Filt: mL
999014
Benthic Infauna (BENT)
Site ID:
Date: / /2020 Visit#: 01 02
Collector(s):
Jar of
SAMPLE ID:
BENTHOS - EXTRA JAR
Site ID:
Date: / /2020 Visit #: Ol 02
Jar of
SAMPLE ID:
ECO Fish Tissue (FTIS)
Site ID:
Date: / /2020 Visit #: 01 02
Length (mm) Min.: Max.:
Bag of
SAMPLE ID:
BENTHOS-EXTRA JAR
Site ID:
Date: / /2020 Visit #: Ol 02
Jar of
SAMPLE ID:
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Sample Labels (Great Lakes)
Page 132
**GREAT LAKES**
WATER CHEMISTRY (CHEM)
Site ID:
*, WATER COLUMN CHLOROPHYLL (WCHL)
Ł Site ID:
Date:
/ 12020 Visit#: Ol 02
p Date: / /2020 Visit #: Ol 02
T1
999020
55 Volume Filtered: mL
« T1 999021
*
Date:
NUTRIENTS (NUTS)
Site ID:
/ /2020 Visit#: Ol 02
ALGAL TOXIN (MICX)
(PETG Bottle (clear, square bottle)
Site ID:
T1
999022
Date: / /2020 Visit #: Ol 02
T3 999023
ALGAL TOXIN (MICZ)
(HDPE Bottle (round Nalgene bottle)
Site ID:
BENTHIC INFAUNA (BENT)
Site ID:
Date: / /2020 Visit#: Ol 02
Date:
/ /2020 Visit#: Ol 02
Jar 1 of
T3
999024
T4 999025
SEDIMENTTOC (SEDC)
Site ID:
SEDIMENT GRAIN SIZE (SEDG)
Site ID:
Date:
/ /2020 Visit#: Ol 02
Date: / /2020 Visit #: Ol 02
T3
999026
999027
12
SEDIMENT ORGAN ICS/METAL (SEDO)
Site ID:
SEDIMENT TOXICITY (SEDX)
Site ID:
Date:
T3
/ /2020 Visit#: Ol 02
999028
Date: / /2020 Visit#: Ol 02
T2 999029
PHYTOPLANKTON (PHYT)
Site ID:
Site ID:
Date' / /2020 Visit#'01 02
Date:
T2
/ /2020 Visit#: Ol 02
999035
Sample ID:
Site ID:
Site ID:
Date:
/ 12020 Visit#: 01 02
Date: / /2020 Visit#: 01 02
Sample ID:
Sample ID:
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FISH TISSUE PLUG (FPLG)
Site ID:
Date: / /2020 Visit #: Ol Q2
999012
ECO FISH TISSUE - INNER BAG OF .
Site ID:
Date: / /2020 Visit #: Ol Q2
Genus Species:
Length (mm) Min.: Max.:
999013
ECO FISH TISSUE - INNER BAG OF .
Site ID:
Date: / /2020 Visit #: Ol 02
Genus Species:
Length (mm) Min.: Max.:
999013
ECO FISH TISSUE - OUTER BAG
75 Site ID:
Date: / /2020 Visit #: Ol 02
Genus Species:
Length (mm) Min.: Max.:
999013
ENTEROCOCCI (ENTE) - BAG
Site ID:
Date: /
/2020 Visit#: 01 02
Vol. Filt: 1
mL 2 mL
999014
Filter : 1
Vol. Filt: mL
999014
Filter: 2
Vol. Filt: mL
999014
Filter: Blank
Vol. Filt: mL
999014
Benthic Infauna (BENT)
Site ID:
Date: / /2020 Visit#: 01 02
Collector(s):
Jar of
SAMPLE ID:
BENTHOS - EXTRA JAR
Site ID:
Date: / /2020 Visit #: Ol 02
Jar of
SAMPLE ID:
BENTHOS - EXTRA JAR
Site ID:
Date: / /2020 Visit #: Ol 02
Jar of
SAMPLE ID:
ECO Fish Tissue (FTIS)
Site ID:
Date: / /2020 Visit #: 01 02
Length (mm) Min.: Max.:
Bag of
SAMPLE ID:
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HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date; 7/2020 Visit #: Ol 02
Genus Species:
Length: mm
999016.01
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: / /2020 Visit #: Ql <32
Genus Species:
Length: mm
999016.02
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: 7 72020 Visit#: Ol 02
Genus Species:
Length: mm
999016.03
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: 7 72020 Visit #: Ql 02
Genus Species:
Length: mm
999016.04
HH FISH TISSUE WHOLE (HTIS)
Site ID:
Date: 7 72020 Visit #: Ql 02
Genus Species:
Length: mm
999016.05
HH FISH TISSUE WHOLE (HTIS) - BAG
Site ID:
Date: 7 72020 Visit #: Ol 02
Genus Species:
Length range: mm to mm
rc
999016
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Appendix C: Shipping and Tracking Guidelines
Tracking Forms in the App
Each shipping group has been assigned a "T" number to help crews identify the correct
section of the Tracking Form in the App to use when sending samples. This "T" number is
located above each of the sample groups in the Tracking Form in the App. Crews will also find
reference to the same "T" numbers on the individual samples labels, on the packing slips that
crews will include in the coolers, and on the top of the pre-printed FedEx return labels
provided in the site kits. Crews submit tracking information via the NCCA App and include
packing slips in the coolers when they send samples to the labs. If any sample listed on the
packing slip is not being shipped, line out the sample ID to indicate that the sample is not in
the cooler.
Procedure for filling out and submitting tracking via the App
1. After ensuring all of the samples to be shipped are properly preserved and prepared
for shipment, access the Tracking Form in the App.
2. Ensure the correct water chemistry sample ID has been entered at the top of the form.
Doing so will populate the sample IDs of all other collected samples. Samples that
were not collected will display a blank sample ID field and the not collected bubble
will be transferred from the individual sample collection forms. The not collected
bubbles are not editable in the Tracking Form; to change the collection status of a
sample, access the pertinent sample collection forms (e.g., Sample Collection, Eco
Fish Collection, and/or Human Health Fish Collection).
3. In the pertinent section of the Tracking Form, check the box under the 'To Ship'
column for each sample being sent in the shipment.
4. Click the 'Enter Shipping Details' button and fill out the resulting popup window with
the destination lab, date shipped, airbill number, sender and sender's phone number.
5. Click the 'save shipping info' button to save the details and the Tracking Form.
6. Once the shipping details have been saved in the App, a date will appear in the
shipped column of the Tracking Form. If the shipping details for a sample need to be
edited, click the date in the shipped column to access the saved shipping details.
Editing the details in this manner changes ONLY one sample at a time. The only way to
enter shipping details for an entire group of samples is during the initial details entry.
a. If the status of the sample needs to change from shipped to not shipped, click
the date in the shipped column to access the saved shipping details and delete
all the shipping info. Click the "save shipping info" button after deleting all the
shipping information. The sample will no longer be marked as shipped and the
"to ship" checkbox will reappear.
7. After all pertinent shipping details have been saved, click the SUBMIT menu button
and select the button next to 'Tracking' and any other the forms that you wish to
submit. Click the green submit button at the bottom of the form list. An email will pop
up on your device addressed to NARSFieldData@epa.gov. Copy yourself, any other
crew members or managers and click send. To ensure that he email was sent, check
the SENT mailbox on your email app and look for the recent email containing the data.
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If the email is not in the SENT mailbox, it was not sent and you should try again after
verifying an internet connection.
8. At any point, if it is determined that data needs to be revised or updated, crews
should feel free to do so in the App and re-submit any edited data or tracking forms
using the steps above. Newly revised data will automatically take the place of previous
data. It is not necessary to re-submit data or tracking forms that were unchanged
however.
9. After submission, a data summary will be automatically emailed back to the email
address from which the submission was received. The Field Crew Leader or his/her
designee should review this data summary for accuracy and make any corrections
necessary and re-submit the pertinent form(s).
Shipping Groups:
T1 - Daily Water Chemistry Samples
• Complete the T1 section of the tracking form for the samples that are shipped
immediately after each sampling event
• water chemistry (CHEM)
• chlorophyll A (WCHL)
• dissolved nutrients (NUTS).
• Send the tracking form and all data forms from the site to the IM Team via the
NCCA App. This serves as the "status report" for that sampling event.
• Ship all of the samples to the lab in the same cooler with the packing slip that
was provided with the label packet. If any sample listed on the packing slip is
not being shipped, line out the sample ID to indicate that the sample is not in
the cooler. If samples from multiple sites are shipped together, then multiple
packing slips must be used.
• 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 as much fresh wet ice as will fit in the cooler
liner.
• Water chemistry samples should be shipped within 24 hours of collection.
T2 - Chilled Batched Samples (Marine = Daily, Great Lakes = Weekly)
• Use this section of the App tracking form for shipping batches of chilled
samples:
• Sediment toxicity (SEDX)
• Sediment grain size (SEDG)
• Total alkalinity (ALKT at marine sites only
• Phytoplankton (PHYT) at Great Lakes sites only
• At marine sites, ship 1 day's worth of samples (up to 3 sites) together in a
single cooler.
• At Great Lakes sites, ship up to 7 site's worth of samples together in a single
cooler.
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• Ship all of the samples to the lab in the same cooler with the packing slip that
was provided with the label packet. If any sample listed on the packing slip is
not being shipped, line out the sample ID to indicate that the sample is not in
the cooler. If samples from multiple sites are shipped together, then multiple
packing slips must be used.
• Samples need to be shipped on as much fresh wet ice as will fit in the cooler
liner.
• At marine sites, chilled batched samples should be shipped the same day as
sampling or the next day.
• At Great Lakes sites, chilled batched samples should be shipped at least every
week
T3 - Frozen Batched Samples
• Use this form for shipping batches of frozen samples:
• Algal Toxins - Microcystes and Cylindrospermopsin (MICX)
• Algal Toxins - Microcystes (MICZ)
• Enterococci (ENTE)
• Fish tissue plugs (FPLG)
• Sediment TOC (SEDC)
• Sediment Organics/Metals (SEDO)
• Nitrogen isotope (D15N) marine sites only
• Ecofish samples (FTIS) (may be shipped separately as a standalone T5
shipment)
• 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.
• Ship all of the samples to the lab in the same cooler with the packing slip that
was provided with the label packet. If any sample listed on the packing slip is
not being shipped, line out the sample ID to indicate that the sample is not in
the cooler. If samples from multiple sites are shipped together, then multiple
packing slips must be used.
• Samples need to be shipped with approximately 20 pounds of dry ice in a cooler
with a two-piece dry ice liner.
• Frozen batched samples should be shipped at least every week.
T4 - Non-Chilled: Batched Samples
• Use this section of the App tracking form for shipping batches of non-chilled
samples:
• Benthic Macroinvertebrates (BENT)
• 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.
• Ship all of the samples to the lab in the same cooler with the packing slip that
was provided with the label packet. If samples from multiple sites are shipped
together, then multiple packing slips must be used.
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• Samples need to be shipped with absorbent material and no ice. Place all
samples and absorbent material inside the cooler liner.
• 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 PG 2 as our guideline. The limited quantity exclusion allows ground shipping
of PG 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 - Eco Fish Tissue
• Use this section of the App tracking form for shipping batches of frozen eco fish
(FTIS) samples.
• Eco Fish samples may be sent in the same cooler as the other frozen batched
samples (T3) 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.
• Ship all of the samples to the lab in the same cooler with the packing slip that
was provided with the label packet. If samples from multiple sites are shipped
together, then multiple packing slips must be used.
• Samples need to be shipped with approximately 20 pounds of dry ice in a cooler
with a two-piece dry ice liner.
• Eco Fish samples should be shipped at least every 2 weeks.
T6 - Human Health Whole Fish Tissue Composite Sample - NGL20, ISA20, and NPA20
sites Only
• Use this section of the App tracking form for shipping frozen human health fish
tissue samples (HTIS).
• Only one human health fish composite sample may be shipped in a single
cooler.
• Ship the sample to the lab in the same cooler with the packing slip that was
provided with the label packet.
• Samples need to be shipped with a minimum of 50 pounds of dry ice (blocks of
dry ice only).
• Human health fish composite samples should be shipped within 2 weeks of
collection.
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T7 - Underwater Video UVID Form [Great Lakes Only]
• Use this section of the App tracking form for shipping the EPA-provided Micro
SD Cards containing all underwater video recorded during the season.
• Before shipping, make backups 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 section(s) of the Tracking
Form in the App for the samples being shipped and will submit tracking via the App. The field
crew will place the samples and the packing slip (in a waterproof bag or plastic sleeve) in a
shipment cooler. If any sample listed on the packing slip is not being shipped, line out the
sample ID to indicate that the sample is not in the cooler. The field crew 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
• See chart for specific
preservation information
for each sample
Holding Time
• 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
Shipping
• 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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Wet Ice
• Ensure that the ice is fresh immediately prior to shipment;
• Line the cooler with a large plastic liner bag.
• Place samples and ice inside the cooler liner and seal the liner with the provided
cable tie.
• Secure the cooler lid with packing tape.
Dry Ice
• 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 Office® locations, FedEx Retail locations such as Walgreens/Wal-
Mart/OfficeMax, or at 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 packing 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.
No Ice
• Line the cooler with a large plastic liner bag.
• Surround the jars with crumpled newpaper or other absorbent material
• If the cooler is not full, add material to keep all bottles upright to prevent leakage.
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Water Chemistry
[CHEM]
•Ship within 24 hours
•Ship 250 mL brown HDPE bottle
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler liner
•Ship on wet ice
Chlorophyll-o
[WCHL]
•Ship with CHEM and NUTS samples
•Ship foil wrapped centrifuge tube
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in provided leak-proof zip-top bag
•Place in cooler liner
•Ship on wet ice
Dissolved Nutrients
[NUTS]
•Ship within 24 hours
•Ship 250 mL white HDPE bottle
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler liner
•Ship on wet ice
r
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Sediment Grain Size
[SEDG]
•Ship daily at marine sites and within 1 week at GL sites
•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 daily at marine sites and within 1 week at GL sites
•Ship in bucket (0.6 gal for estuarine, 1 quart for Great Lakes)
•Confirm label completed & taped
•Tighten the lid securely and ensure it will not loosen in
shipping
•Place in lined cooler with other chilled batched samples.
•Ship on wet ice
Total Alkalinity
[ALKT]
(Marine only)
•Ship within 24 hours
•Ship two 125 mL white rectangular HDPE bottles
•Confirm label completed & taped
•Seal with plastic electrical tape
•Place in cooler liner
•Ship on wet ice
Phytoplankton
[PHYT]
(GL only)
•Ship within 1 week
•Ship in HDPE bottle (1 L, white, narrow mouth)
•Confirm preserved with 10 ml 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
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Algal Toxins
[MICX] and [MICZ]
r
Enterococci
[ENTE]
•Ship at least every week
•Freeze after collection
•Ship in HDPE 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 lbs of dry ice
•Ship at least every week
•Ship in frozen, microcentrifuge tubes
•Confirm labels completed
•Place each tube in small bubble bag with label on outside
•Place bag in zip-top bag
•Place in cooler lined with dry ice insert along with other
frozen batched samples
•Pack cooler with 20 lbs of dry ice
Fish Plugs
[FPLG]
A
Sediment TOC
[SEDC]
•Ship at least every week
•Freeze after collection
•Ship in glass scintillation vial
•Confirm label completed & taped
•Place vial in small bubble bag and then 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 lbs of dry ice
•Ship at least every week
•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
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Sediment
Organics/Metals
[SEDO]
A
Nitrogen Isotopes
[D15N]
Benthic
Macroinvertebrates
[BENT]
•Ship at least every week
•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
•Ship at least every week
•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
•Ship every 2-3 weeks
•Preserve benthos samples immediately upon collection
•Ship in HDPE 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
•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 lbs of dry ice
Ecological Whole Fish
Tissue
[FTIS]
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Human Health Whole
Fish Tissue
[HTIS]
(NGL20, ISA20, and
NPA20 sites only)
•Ship at least every 2 weeks
•Freeze after collection, as soon as possible (-20 °C cooler)
•Ship in bags
•Confirm label completed & taped
• Pack cooler with 50 lbs of dry ice
Underwater Video
[UVID] (GLonly)
•Ship at end of season or as Micro SD cards get full
•Back up files to computer hard drive
•Be sure files are named appropriately
•Package EPA-provided Micro SD cards in plastic cases
•Ship in FedEx envelope
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[Tl]
Daily Water
Chemistry Samples
1 - HDPE bottle
(250 mL, amber)
CHEM
1 - Filter in centrifuge
tube (50 mL), in
zip-top bag
WCHL
1 - HDPE bottle
(250 mL, white)
NUTS
N
' [T2] i
Chilled Samples (Batched)
1 - Plastic bag (quart size, double
SEDG
- Sediment bucket
(0.6 gal) marine OR (1 qt) GL
SEDX
- HDPE (125 mL white)
ALKT {marine only)
- HDPE (1L, white narrow mouth)
PHYT (GL only)
T
J
Pack 1 day 's worth
of samples (up to 2
sites) in lined cooler
with wet ice
ICE
MARINE
Pack 1 day's
worth of samples
(up to 3 sites) in
lined cooler with
wet ice
I
[T3]
Frozen Samples (Batched)
1 - Glass scintillation
vial with 2 plugs
FPLG
1 - PETG bottle (500 mL, clear,
square)
MICX
1 - HDPE bottle (500 mL, white,
round, wide-mouth)
MICZ
2 - Filters in centrifuge tubes in
bag (plus 1 blank if revisit site)
ENTE
1 - Glass jar (60 mL)
SEDC
1 - Glass jar (120 mL)
SEDO
1 - Glass jar (60 mL)
D15N (marine
only)
GREATLAKES
Pack 1 week's
worth of samples
(up to 7 sites) in
lined cooler with
wet ice
SHIP WITHIN 24
HOURS
(MON-FRI)
Express
PRIORITY
OVERNIGHT
SHIP WITHIN 24
HOURS
(MON-FRI)
Express
PRIORITY
OVERNIGHT
SHIP WITHIN 1
WEEK
(MON-FRI)
Express
PRIORITY
OVERNIGHT
[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
[T4]
Non-Chilled
Samples
(Batched)
1 or more -
HDPE bottle
(1 L, white,
wide-mouth)
BENT
[T6]
Frozen HH Whole
Fish Samples
(Batched)
5-10 Fish in large
plastic composite
bags
HTIS
Great Lakes
Nearshore
(NGL20) sites,
Great Lakes Island
(ISA20) sites,
and
Great Lakes Parks
(NPA20) sites
Pack 2-4 sites' worth of samples (depending
on whether Eco Fish are included and size
of Eco Fish sample) in cooler M>ith dry ice
insert and 20 pounds dry ice
DRY
ICE
...
[T7]
UW Video
(Batched)
2 or 4 -
Videos
per site
ITVID
Great Lakes
sites only
Pack up to 12
sites' -worth of
samples in lined
cooler with
NO ICE
I
Pack frozen fish in
cooler with 50
pounds of
dry ice
I
Rename files,
backup to
computer.
Place in
plastic case
and padded
envelope
SHIP WITHIN 1 WEEK
(MON-FRI)
Express
PRIORITY OVERNIGHT
SHIP EVERY
2-3 WEEKS
ANYTIME
Ground
GROUND
SHIP MON -
I III 'RS (No
Saturday
Delivery)
St
Express
PRIORITY
OVERNIGHT
SHIP AT
END OF
SEASON
Express
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National Coastal Condition Assessment 2020
Field Operations Manual
Version 1.1 Page 147
SHIPPING
GROUP
SAMPLE
SAMPLE TARGET
VOLUME
CONTAINER
PRESERVATIVE
PACKAGING FOR
SHIPMENT
HOLDING TIME
DAILY WATER
CHEMISTRY
SAMPLES
(Tl)
Water Chemistry
(CHEM)
250 mL
HDPE bottle
(250 mL, amber)
Wet ice
Line cooler with heavy
plastic bag cooler liner
Ship in cooler with wet ice
24 hours
(ship same day as
sampling or next day)
to
WRS lab - Corvallis or
approved State lab
Chlorophyll-^
(WCHL)
Collection: 2L
HDPE bottle
(2 L, amber)
Wet ice prior to filtering
Processing: readily
visible stain on filter,
max 2000 mL
Filter in 50 mL
centrifuge tube
(foil wrapped)
Dry ice in field
Dissolved Nutrients
(NUTS)
250 mL of filtrate from
chl-^ filtering
HDPE bottle
(250 mL, white)
Wet ice in field
CHILLED
BATCHED
SAMPLES
(T2)
Sediment Grain Si2e
(SEDG)
100 mL
Plastic bag
(double bagged, quart
size^^
Wet ice in field
Line cooler with heavy
plastic bag cooler liner
Ship in cooler with wet ice
MARINE:
24 hours
(ship same day as
sampling or next day)
to
GLEC lab -
Traverse City
GREAT LAKES:
Sediment Toxicity
(SEDX)
MARINE:
target = 1800 mL
minimum = 900 mL
GREAT LAKES: target
= 900 mL, minimum
-100 mL
MARINE = screw top
bucket (0.6 gal)
GREAT LAKES = snap
lid bucket (1 qt)
Wet ice in field
Total Alkalinity
(ALKT) Marine only
125 mL in each of 2
bottles
HDPE bottles (2)
(125 mL, white)
Wet ice in field
Batch; ship at least every
week
to
GLEC lab -
Traverse City
Phytoplankton
(PHYT)
Great Lakes only
1 L
HDPE bottle
(1 L, white, narrow-
mouth)
Add 10 mL Lugol's
solution
Wet ice in field
Hold chilled in the dark
FROZEN
BATCHED
SAMPLES
(T3)
Algal Toxins
(MICX)
500 mL (leave
headspace)
PETG bottle
(500 mL. clear, square)
Wet ice in field,
Freeze as soon as possible
Ship in cooler with
2-piece dry ice insert
and 20 pounds of
DRY ICE
Note: Frozen Batched
(T3)and Ecofish samples
(T5) may be shipped
together or separately
Batch, ship within 1
week to GLEC lab —
Traverse City
Algal Toxins
(MICZ)
500 mL (leave
headspace)
HDPE bottle
(500 mL. white, wide-
mouth, round^^
Wet ice in field,
Freeze as soon as possible
Enterococci
(ENTE)
Collection:
250 mL
HDPE bottle
(250 mL, pre-sterilized,
clea^^^
Wet ice prior to filtering
(at least 15 minutes)
Processing:
Two 50 mL filtrations
2 filters in
microcentrifuge tubes
Dry ice in field; hold in
freezer; MUST be filtered
& frozen within 6 hours of
collection;
Hold in freezer or on dry
ice
Filter blanks
(revisit sites only):
One 20 mL filtration of
PBS
1 filter in microcentrifuge
tube
Fish Tissue Plugs
(FPLG)
2 plugs
glass scintillation vial
(20 mL)
Dry ice in field; Hold in
freezer or on dry ice
Sediment Total Organic
Carbon
(SEDC)
50 mL (leave headspace)
glass jar
(60 mL, amber)
Dry ice in field
Sediment
Organics/Metals
(SEDO)
100 mL (leave
headspace)
glass jar
(120 mL, amber)
Dry ice in field
Nitrogen Isotopes
(D15N) Marine only
50 mL (leave headspace)
glass jar
(60 mL, amber)
Dry ice in field
ECOFISH
SAMPLES
(T5)
Whole Fish Tissue
Sample
(FTIS)
5-20+ fish (300 g whole
body tissue)
2 gallon self-sealing bags
Large outer bag
Dry ice in field; Hold in
freezer
NON-
CHILLED
BATCHED
SAMPLES
(T4)
Benthic
Macroinvertebrates
(BENT)
All organisms in grab(s)
HDPE bottle(s)
(1 L, white, wide-mouth)
Stained formalin solution
Ship in cooler lined with
plastic bag cooler liner
Batch, ship at least every
2-3 weeks to GLEC lab
— Traverse City
HUMAN
HEALTH
WHOLE FISH*
(T6)
Human Health Whole
Fish Tissue Sample
(HTIS)*
Great Lakes only
5-10 whole fish (500 g of
fillet weight) minimum
fish length of 190 mm
Wrapped individually in
solvent rinsed foil
Sealed in poly tubing
Large outer plastic bag
Dry ice in field; Hold in
freezer
Ship in provided HTIS
cooler with 50 pounds of
DRY ICE
Batch, ship weekly
(except on Fridays,
Saturdays, or the day
before Federal holidays)
to HTIS lab
UW VIDEO
(T7)
Underwater Video
(UVID)
Great Lakes only
1 minute videos of
benthic habitat on both
A and B cameras
2 Micro SD Cards
(one per camera)
Rename files on Micro SD
Cards and back up files to
computer
Place Micro SD Card in
plastic case,
Ship in padded envelope
Ship when cards are
nearly full and/or at end
of season to EPA
Duluth lab
* Human Health Fish composite sample is collected at all 225 probabilistic nearshore Great Lakes sites (prefix = NGL20), all 38 Great Lakes island
SITES (PREFIX = ISA20), AND ALL 12 GREAT LAKES PARK SITES (PREFIX = NPA20).
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National Coastal Condition Assessment 2020
Field Operations Manual
Version 1.1
Page 148
Appendix D: Microplastics Collection and
Processing
At specifically targeted sites in the northeast, crews will collect a sample of sediment for the
analysis of microplastics. Microplastics have been found in every aquatic environmental
matrix on the globe from coastal waters to artic ice, and deep-sea trench sediments. These
reports demonstrate that the likelihood of exposure to microplastics by aquatic organisms is
ever-increasing. There are many routes of contamination for microplastics to enter the
aquatic environment (e.g., surface run-off, wastewater treatment plants, and poor trash
management). Once in aquatic systems, due to bio-fouling and aggregation, sediments are the
likely ultimate sink for most microplastics. Sediments also serve as the habitat for many
organisms at the base of aquatic food chains and therefore act as a vector for many
anthropogenic contaminants, including possibly microplastics, to enter food webs.
Quantification of microplastics in sediments is one of the first steps to understand the fate
and effects of microplastics in aquatic food webs including those that may ultimately affect
humans. By better understanding how much microplastics are present in marine coastal
sediments, we improve our ability to predict potential microplastics exposure to humans.
This methodology describes the procedure for sampling sediments for microplastic analysis.
The sampling procedure is very similar to conventional sediment sampling for other
anthropogenic contaminants (e.g., metals, PCBs, etc.). However, special care must be taken
to avoid accidental contamination of the sediment sample by the collectors' clothing or
sources of plastics associated with the sampling vessel (e.g., fraying polymer lines). To
address this source of contamination, while the sediment sample is being collected, an "air
blank" will be collected to allow quantification of any microplastics entering the sample via
airborne-sources.
Equipment ft supplies: mk:i aplastics collection and processing
For collecting samples
nitrile gloves
sediment sampling collection device
2 (500mL) glass jars
1 metal spatula or stainless steel teaspoon
Electrical tape, plastic
For recording
microplastics sample label
measurements
fine-tipped indelible markers (for labels)
clear tape strips
1. Rinse all collection equipment with sea water in between sites to remove sediment
debris, followed by Dl water to avoid cross contamination.
2. Collect sediment using appropriate collection device (the same device used for the
collection of sediment for chemical analysis will suffice).
3. Minimize exposure to synthetic fibers:
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National Coastal Condition Assessment 2020
Field Operations Manual
Version 1.1
Page 149
o wear cotton laboratory coats or cotton t shirts. All crew members should wear
the same color, but not white, blue or black. Note the color worn on the
sample label.
o Note the color of the nitrile gloves worn on the sample label,
o Minimize exposure to fraying ropes or other plastic items onboard. Do not use
synthetic wipes such as Kimwipes for equipment cleaning.
4. After retrieving the grab sampler, ensure the grab is acceptable (e.g., level sediment
layer, not overfilled, etc. - see Figure 13.1).
5. Open one of the glass jars before transferring sediment to the sample jar to collect an
air blank. Position this empty air blank jar open to the atmosphere, close to the
sediment sample whenever it's exposed to air.
6. Using a metal spatula or stainless steel teaspoon to transfer sediment directly from
the grab sampler to the 500 mL glass sample jar, collect the top 5 cm of the grab.
o The target collection volume is 400mL of sample, which will likely require
multiple grabs, depending on the size of your grab sampler and the depth of
sediment in each collection,
o Leave approximately 1 inch of headspace in the sample jar.
7. Close both the sample and air blank jars and secure the lids with electrical tape.
8. Fill out sample labels for the both the sample and air blank jars and cover with clear
tape.
9. If plastic debris is present on the boat (i.e., - shedding equipment or rope), take a
small sample of plastic and retain it in aluminum foil for potential analysis. This will
help reduce uncertainty in the air blank contamination.
o Label this sample with an extra label from the label packet with the same
sample ID as the air blank sample.
10. Ship the sample and air blank to the lab with samples of benthic macroinvertebrates,
or whenever is convenient. No chilling or freezing is necessary
o Be sure to keep each sample jar upright to help prevent leakage,
o Pad all glass jars with packing material to avoid breakage.
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