Forest Service National 

Protocols for Sampling Air 

Pollution-Sensitive Waters

Prepared for the USDA Forest Service’s 

Air Resource Management Program

United States

Department of

Agriculture

Forest Service

Rocky Mountain Research 

Station

General Technical Report

RMRS-GTR-278WWW

April 2012
Authors

T.J. Sullivan, E&S Environmental Chemistry, Inc., Corvallis, OR. 

A.T. Herlihy, Oregon State University, Corvallis, OR. 

G.B. Lawrence, U.S. Geological Survey, Troy, NY.

J.R. Webb, University of Virginia, Charlottesville, VA.

Acknowledgments

These protocols were prepared by E&S Environmental Chemistry, Inc., under contract to the 

U.S. Forest Service, Air Resources Management Program, under the direction of A. Mebane. 

Protocol development benefited greatly from extensive review comments received on earlier 

drafts from Forest Service staff. In particular, helpful guidance and suggestions were 

provided by A. Mebane, C. Huber, J. Gurrieri, B. Gauthier, S. Grant, B. Roper, M. Hudy, C. 

Carlson, and W. Shaw. The final document was prepared by J. Charles. Field testing of the 

draft sampling protocols was kindly provided by C. Huber, T. Porwoll, and T. Wickman. 

Cover Photo

Todd McDonnell, E&S Environmental Chemistry, Inc., Corvallis, OR.

Citation

Sullivan, T.J., ed. 2012. USDA Forest Service national protocols for sampling air pollutionsensitive waters. Gen. Tech. Rep. RMRS-GTR-278WWW. Fort Collins, CO: U.S. Department 

of Agriculture, Forest Service, Rocky Mountain Research Station. 334 p. 

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endorsement purposes.

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communication of program information (Braille, large print, audiotape, etc.) should contact USDA’s 

TARGET Center at 202-720-2600 (voice and TDD). 
Forest Service National Protocols

for Sampling Air Pollution-Sensitive Waters

Prepared for the USDA Forest Service’s 

Air Resource Management Program by: 

T.J. Sullivan, E&S Environmental Chemistry

A.T. Herlihy, Oregon State University

G.B. Lawrence, U.S. Geological Survey

J.R. Webb, University of Virginia 

iii

Table of Contents

Section 1. Water Chemistry Field Sampling Protocols ........................................ 1

1.1 BACKGROUND................................................................................................................................................. 1

1.1.1 Resources Sensitive to Atmospheric Deposition .................................................................................... 2

1.1.2 Study Purpose and Objectives ............................................................................................................... 7

1.2 STUDY DESIGN..............................................................................................................................................11

1.2.1 Lake or Stream Characterization.........................................................................................................12

1.2.2 Synoptic Survey....................................................................................................................................14

1.2.3 Long-Term Monitoring ........................................................................................................................14

1.2.4 Characterization of Stream Chemistry During Hydrologic Events.....................................................15

1.2.5 Other Uses of Resulting Data ..............................................................................................................15

1.3 WHERE, WHAT, AND WHEN TO SAMPLE.....................................................................................................16

1.3.1 Where to Sample ..................................................................................................................................18

1.3.2 What to Measure..................................................................................................................................24

1.3.3 When to Sample ...................................................................................................................................31

1.4 FIELD METHODS...........................................................................................................................................34

1.4.1 Pre-Trip Preparations .........................................................................................................................35

1.4.2 Sample Collection................................................................................................................................40

1.4.3 On-Site Measurements.........................................................................................................................48

1.4.4 Post Collection Sample Processing, Documentation, and Cleanup ....................................................54

1.4.5 Sample Packaging and Transportation................................................................................................54

1.4.6 Sequence of Field Activities.................................................................................................................57

1.4.7 Safety in Field Activities......................................................................................................................57

Section 2. Laboratory Protocols............................................................................59

2.1 INTRODUCTION.............................................................................................................................................59

2.2 LABORATORY PREPARATION PRIOR TO SAMPLE ANALYSIS ......................................................................59

2.2.1 Bottle Cleaning ....................................................................................................................................59

2.2.2 Sample Processing, Preservation, and Storage ...................................................................................63

2.3 CHEMICAL ANALYSIS...................................................................................................................................67

Section 3. Quality Assurance/Quality Control Protocols...................................71

3.1 INTRODUCTION.............................................................................................................................................71

3.2 ATTRIBUTES OF DATA QUALITY..................................................................................................................72

3.2.1 Method Detection and Reporting Limits..............................................................................................72

3.2.2 Precision and Accuracy .......................................................................................................................76

3.2.3 Completeness.......................................................................................................................................77

3.2.4 Comparability ......................................................................................................................................78

3.2.5 Representativeness...............................................................................................................................78

3.2.6 Recommended Laboratory Data Quality Objectives ...........................................................................78

3.3 QA/QC SAMPLE TYPES................................................................................................................................79

3.3.1 Filter Blanks, Analytical Blanks, and Field Blanks.............................................................................80

3.3.2 Replicate Environmental Samples .......................................................................................................81

3.3.3 Spiked Project Samples........................................................................................................................82

3.3.4 External Quality Assurance Samples...................................................................................................82
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

iv

3.4 FIELD QUALITY ASSURANCE .......................................................................................................................84

3.4.1 Sample Containers...............................................................................................................................84

3.4.2 Field Measurements.............................................................................................................................84

3.5 REPORTING QUALITY ASSURANCE DATA ...................................................................................................85

3.6 LABORATORY AUDITS AND CERTIFICATION ...............................................................................................87

3.7 DATA ENTRY.................................................................................................................................................88

3.8 SUMMARY .....................................................................................................................................................88

Section 4. Data Analysis Protocols........................................................................91

4.1 BACKGROUND AND OBJECTIVES..................................................................................................................91

4.2 EVALUATION OF DATA QUALITY.................................................................................................................93

4.2.1 Charge Balance ...................................................................................................................................93

4.2.2 Calculated Versus Measured Conductivity..........................................................................................97

4.2.3 Calculated Versus Measured ANC ......................................................................................................98

4.2.4 Other Validation Procedures.............................................................................................................100

4.2.5 Final Data Quality Determination.....................................................................................................101

4.3 APPLY PROCEDURES TO PREPARE RAW DATA FOR GRAPHICAL AND STATISTICAL ANALYSIS .............102

4.3.1 Censored Data ...................................................................................................................................102 

4.3.2 Outliers and Missing Values..............................................................................................................102

4.3.3 Multiple Observations........................................................................................................................105 

4.3.4 Treatment of Zeros and Negative Values...........................................................................................106 

4.3.5 Treatment of Seasonality ...................................................................................................................106 

4.4 CONDUCT EXPLORATORY ANALYSES........................................................................................................107

4.4.1 Analysis of Water Quality Status .......................................................................................................107

4.4.2 Graphing and Quality Analysis .........................................................................................................108 

4.4.3 Recommended Data Analyses............................................................................................................109

4.5 CONDUCT STATISTICAL ANALYSES ...........................................................................................................128

4.5.1 Statistical Tests for Difference...........................................................................................................128

4.5.2 Trend Detection ................................................................................................................................. 130

4.5.3 Statistical Power................................................................................................................................133 

4.6 REPORT DATA IN STANDARDIZED FORMATS ............................................................................................134

4.7 SUMMARY AND CONCLUSIONS...................................................................................................................134

Section 5. Field Sampling Protocols for Aquatic Biota ....................................135

5.1 BACKGROUND ........................................................................................................................................135

5.1.1 Lake Zooplankton ..............................................................................................................................135

5.1.2 Stream Macroinvertebrates ...............................................................................................................136

5.2 STUDY DESIGN .......................................................................................................................................137

5.3 SITE SELECTION....................................................................................................................................138

5.3.1 Laying Out the Support Reach for Stream Macroinvertebrates.........................................................138

5.3.2 Lake Selection for Zooplankton Sampling .........................................................................................141

5.4 PRE-TRIP PREPARATION ....................................................................................................................143

5.4.1 Equipment and Supplies.....................................................................................................................143

5.4.2 Equipment Cleaning Protocols..........................................................................................................144
TABLE OF CONTENTS

v 

5.5 COLLECTION PROCEDURES...............................................................................................................145

5.5.1 Stream Benthic Macroinvertebrates..................................................................................................145

5.5.2 Lake Zooplankton .............................................................................................................................. 149

5.6 SAMPLE PROCESSING, PRESERVATION, AND HANDLING.........................................................151

5.6.1 Stream Benthic Macroinvertebrates..................................................................................................151

5.6.2 Lake Zooplankton ..............................................................................................................................153

5.7 DOCUMENTATION AND TRACKING..................................................................................................154

5.8 LABORATORY ANALYSIS ....................................................................................................................154

5.9 QUALITY ASSURANCE..........................................................................................................................155

5.9.1 Sample Replication ............................................................................................................................155

5.9.2 Taxonomy...........................................................................................................................................155

5.10 INTERPRETATION..................................................................................................................................156

5.10.1 Biological Metrics .............................................................................................................................156

5.10.2 Multimetric Indices and Predictive Modeling ...................................................................................157 

5.10.3 Taxonomic Resolution........................................................................................................................157

5.10.4 Index of Biotic Integrity .....................................................................................................................158

5.10.5 Biotic Effects Analysis .......................................................................................................................161 

Section 6. Transition Plan....................................................................................167

6.1 BACKGROUND ........................................................................................................................................167 

6.2 TRANSITION STEPS ...............................................................................................................................167 

6.3 ANTICIPATED PROTOCOL CHANGES ..............................................................................................170 

6.4 DECISION OF WHETHER OR NOT TO CHANGE PROTOCOLS....................................................171 

Section 7. Training Plan.......................................................................................173

7.1 BACKGROUND ........................................................................................................................................173 

7.2 PLANNING THE TRAINING SESSIONS...............................................................................................174 

7.3 CLASSROOM TRAINING.......................................................................................................................176 

7.3.1 Handouts............................................................................................................................................177 

7.3.2 Questions...........................................................................................................................................177 

7.3.3 Prepare for Field Training ................................................................................................................177 

7.4 FIELD TRAINING....................................................................................................................................177 

Section 8. References............................................................................................179
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

vi

Appendices..................................................................................................... 189

APPENDIX A. ACRONYMS AND ABBREVIATIONS

APPENDIX B. GLOSSARY

APPENDIX C. SAMPLING PROCEDURE FOR SURFACE WATER CHEMISTRY

APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

APPENDIX E. DATA ENTRY FORMS AND INSTRUCTIONS FOR FIELD SAMPLING ACTIVITIES

APPENDIX F. LABELING INSTRUCTIONS FOR FIELD SAMPLE CONTAINERS

APPENDIX G. EXAMPLE STANDARD OPERATING PROCEDURES FOR LAB ANALYSIS PROTOCOLS

APPENDIX H. TRAINING CHECKLISTS

APPENDIX I. JOB HAZARD ANALYSIS (JHA)
TABLE OF CONTENTS

vii

List of Tables

Table 1-1. Recommended sensitive receptors and indicators for air quality related values affected by 

atmospheric deposition of air pollutants. ................................................................................... 3

Table 1-2. General guidelines for evaluating the likelihood of aluminum toxicity in fresh water. ............ 6

Table 1-3. Common management issues for FS ARM program staff, with associated field study 

approaches. ................................................................................................................................ 8

Table 1-4. Elements to be considered in formulating sampling questions. .............................................. 10

Table 1-5. Examples of questions that could be used to guide inventory, characterization, 

and monitoring study design. .................................................................................................. 11

Table 1-6. General guidance regarding water quality study design. ......................................................... 12

Table 1-7. Parameters to consider for possible inclusion in surface water acidification 

studies. ..................................................................................................................................... 27

Table 1-8. Parameters to consider for possible inclusion in studies of atmospheric 

nutrient nitrogen enrichment of fresh waters. .......................................................................... 28

Table 1-9. Suggested timing of surface water samples for evaluation of sensitivity to 

and effects from atmospheric deposition. ................................................................................ 31

Table 1-10. Key issues to consider when initiating and implementing an inventory of lake 

or stream water chemistry to assess the effects of atmospheric deposition. ............................ 36

Table 1-11. Key issues to consider when initiating and implementing a long-term monitoring effort to 

document and quantify trends in lake or stream chemistry over time 

in response to inputs of atmospheric deposition. ..................................................................... 36

Table 1-12. Key issues to consider when conducting modeling using the MAGIC model 

(Cosby et al. 1985) to estimate critical load or to calculate changes in lake/stream 

chemistry in response to future emissions controls. ................................................................ 36

Table 1-13. Ancillary measurements that may help in interpretation of lake or stream water 

chemistry data.......................................................................................................................... 53

Table 2-1. Summary of a typical bottle rinsing protocol for low-nutrient, low-ionic strength samples. .. 61

Table 2-2. Example laboratory aliquot schedule for a particular project. ................................................. 64

Table 2-3. Recommended laboratory holding times. ................................................................................ 65

Table 2-4. Example low-volume sample schedule. .................................................................................. 67

Table 3-1. Recommended DQOs for detection limits, precision, accuracy, and completeness. .............. 76

Table 3-2. Recommended laboratory quality control samples. ................................................................ 80

Table 3-3. Field quality control samples................................................................................................... 84

Table 4-1. Some example approaches for FS ARM program data analysis, tied to the 

purpose and general approach of the field study. .................................................................... 92

Table 4-2. Factors for converting mg/L units or pH units to μeq/L units. ................................................ 94

Table 4-3. Data validation quality control procedures. ............................................................................. 96

Table 4-4. Critical Q-values for Dixon’s outlier Q-test, at the 0.95 confidence level. ........................... 105
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

viii

Table 4-5. Example variables with which to subset water quality data, according to measured 

water chemistry for analysis. ................................................................................................. 110

Table 4-6. Example variables with which to subset water quality data for analysis according to 

features other than measured water chemistry. ..................................................................... 110

Table 4-7. Values of t. ............................................................................................................................ 131

Table 5-1. Example approaches for FS ARM program biological sampling, tied to the purpose 

of the field study. ................................................................................................................... 137

Table 5-2. Equipment and supplies for benthic macroinvertebrates. ...................................................... 143 

Table 5-3. Equipment and supplies for collecting zooplankton samples. ............................................... 144 

Table 5-4. List of metrics in each category used in the EPA National Wadeable Stream 

Assessment. ........................................................................................................................... 160 
TABLE OF CONTENTS

ix

List of Figures

Figure 3-1. Schematic illustration of precision and accuracy. ................................................................ 77

Figure 3-2. Results from analysis of low-concentration QC samples for nitrate analysis. ..................... 86

Figure 3-3. Low-concentration QC samples for sulfate analysis. ........................................................... 87

Figure 4-1. Cation sum versus anion sum. .............................................................................................. 95

Figure 4-2. Calculated versus measured conductivity. ........................................................................... 98

Figure 4-3. Calculated carbonate alkalinity versus laboratory titrated ANC with no 

obvious outliers. ................................................................................................................... 99

Figure 4-4. Patterns in DOC concentration in streams within a particular region. ............................... 101

Figure 4-5. Minimum stream ANC sampled at each site during each year versus median spring 

ANC for all samples collected at that site during that spring season. ................................ 107

Figure 4-6. Box plots comparing hypothetical lake ANC values measured in samples 

collected during the spring versus the fall season. ............................................................. 108

Figure 4-7. Changes in the concentration of major water chemistry constituents during a 

hypothetical hydrological episode in one stream. .............................................................. 112

Figure 4-8. Time series of major ions and discharge in Treasure Lake in the Sierra Nevada during 

snowmelt in 1993. .............................................................................................................. 113

Figure 4-9. Daily discharge (A) and nitrate concentration (B) in Icy Brook and Andrews Creek 

within the Loch Vale watershed, Rocky Mountain National Park, in 

April-September 1992. ....................................................................................................... 114

Figure 4-10. Ratio of NO3

-

:(SO4

2- + NO3-

) concentration versus ANC in stream water samples 

collected during hydrological episodes in four streams included in the Adirondack 

region of EPA’s Episodic Response Program. ................................................................... 115

Figure 4-11. Map of summer NO3

-

 concentrations in drainage lakes sampled by the Adirondack 

Lakes Survey Corporation in the Adirondack region of New York. ................................. 116

Figure 4-12. Example space-for-time substitution analysis for four variables across a gradient of 

expected response. ............................................................................................................. 117

Figure 4-13. Results of a classification system devised to indicate expected low-ANC streams 

(in this example, based on geology and elevation) compared with actual low-ANC 

streams, which are represented as dots on the map. ........................................................... 118

Figure 4-14. A histogram of the frequency of occurrence within a region (or forest) of surface 

water SO4

2- concentration can reveal the typical distribution of the range of values 

that may be attributable to broad patterns of regional atmospheric inputs versus 

the more sporadic occurrence of high values that more likely derive from geological 

sources of S. ....................................................................................................................... 119

Figure 4-15. Critical load simulated by the MAGIC model to protect streams in Shenandoah 

National Park against acidification to ANC below 0 (top panel) and 20 μeq/L 

(bottom panel) by the year 2040 is plotted as a function of 1990 ANC. ........................... 120

Figure 4-16. Example trends analyses of ANC in lake and stream waters, based on data from 

EPA’s Long Term Monitoring program. ........................................................................... 122
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

x 

Figure 4-17. Plot of ANC versus ion ratio of SO4

2- concentration divided by the sum of base 

cation concentrations (SBC) for low ANC (≤ 200 μeq/L) surface waters in 

(a) Adirondacks, (b) Catskill Mountains, and (c) Cascade Mountains. ............................. 124

Figure 4-18. Relationship between lake size and lake ANC in the Adirondack Mountains. .................. 125

Figure 4-19. Nitrogen outputs in soil water or stream water versus N deposition inputs throughout 

Europe. ............................................................................................................................... 126

Figure 4-20. Observed relationship between NO3- leaching loss in runoff and mean air temperature 

at an experimental watershed site at Gårdsjön, Sweden. ................................................... 127 

Figure 4-21. Example patterns of NO3- concentration in surface water at four sites at various stages 

of watershed N-saturation. ................................................................................................. 128 

Figure 4-22. Schematic representation of data normality. ...................................................................... 129 

Figure 4-23. Plot, with regression line, of SO42- concentration in Deep Run, Shenandoah National 

Park, over the period of monitoring record. ....................................................................... 130 

Figure 5-1. Equipment, supplies, and sequence for collecting benthic macroinvertebrate samples. .... 141

Figure 5-2. Modified D-frame kick net. ............................................................................................... 146

Figure 5-3. Wisconsin net and collection bucket diagram. Some microzooplankton nets 

have a reducing collar attached. ......................................................................................... 150

Figure 5-4. Location of the nine ecoregions used in Wadeable Streams Assessment IBI 

development. ...................................................................................................................... 159

Figure 5-5. Schematic depicting mayfly richness over time in response to changes in stream 

chemistry. ........................................................................................................................... 162

Figure 5-6. Average number of families of aquatic insects for each of 14 streams in Shenandoah 

National Park versus the mean (left) or minimum (right) ANC of each stream. ............... 164

Figure 5-7. Zooplankton taxonomic richness versus ANC for a combined Adirondack dataset, 

based on 111 lake visits to 97 lakes in the EMAP, ELS, and STAR zooplankton 

surveys. .............................................................................................................................. 166
1

SECTION 1. 

WATER CHEMISTRY 

FIELD SAMPLING PROTOCOLS T.J. Sullivan, J.R. Webb, A.T. Herlihy, and G.B. Lawrence

1.1 BACKGROUND

The first step in designing a surface water sampling program is identifying one or more problems 

or questions that require information on water quality. Common water quality problems include 

nutrient enrichment (from a variety of causes), effects of atmospheric deposition (acidification, 

eutrophication, toxicity), and effects of major disturbances such as fire or pest infestations. Once 

the problems or questions have been clearly defined, a sampling program can be designed that 

addresses where to sample, what to measure, and when and how to conduct the sampling. The 

selection of measurements should be 

tailored to specific study objectives and 

to the study design, which guides the 

specifics of field, laboratory, and data 

analysis protocols.

A variety of air pollutants have the 

potential to stress aquatic ecosystems 

through contributions from the 

atmosphere to the Earth’s surface. The 

main focus of this protocol is on 

atmospheric pollutants that contribute 

to surface water acidification and 

eutrophication (nutrient enrichment). 

Both atmospheric sulfur (S) and 

nitrogen (N) have the potential to cause 

acidification. Atmospheric N can also 

cause eutrophication of aquatic 

ecosystems in which the N supply is 

limiting for algal or plant growth. 

Sampling for atmospherically 

deposited toxic materials is also 

addressed, but with lesser coverage. 

Acidification

When fossil fuels are burned, as in the combustion 

of coal in power plants, sulfur and nitrogen are 

emitted into the atmosphere. The sulfur and 

nitrogen can contribute to the acidification of surface 

waters and soils in the ecosystem. 

Large areas of the Northeast, Appalachian 

Mountains, northern Florida, Upper Midwest, and 

mountainous portions of the Western United States 

have many lakes and streams with low acid 

neutralizing capacity and are therefore potentially 

sensitive to acidification from atmospheric 

deposition of sulfur and/or nitrogen. 

There are already many acidified surface waters in 

the Eastern United States, while in the Western 

United States many surface waters are susceptible 

to acidification, but levels of acidic deposition are 

relatively low and acidic surface waters are rare. 

Many of the areas having acid-sensitive surface 

waters, especially in the Northeastern United States 

and Appalachian Mountains also contain extensive 

areas with acid-sensitive soils.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

2

Many fresh waters in the United States are thought to be phosphorus (P)-limited (U.S. EPA 

2008). In such waters, the addition of P would be expected to increase plant and/or algal growth, 

whereas the addition of N would not. Nevertheless, there are also fresh waters considered to be 

N-limited or N and P co-limited. 

The U.S. Forest Service (FS) collects and analyzes data to identify ecosystems and resources 

sensitive to air pollution and to determine if these resources are being impacted. Such studies are 

often conducted at the individual forest level. This protocol provides FS staff with a consistent 

framework regarding decisions of where, when, and how to conduct water sampling for the 

purpose of evaluating and monitoring air pollution effects on aquatic ecosystems. It is based on 

protocols developed by the U.S. Environmental 

Protection Agency, U.S. Geological Survey, and 

Forest Service, including documents prepared by 

Herlihy (1997), Turk (2001), Webb et al. (2004), 

Eilers (2007), and Sullivan and Herlihy (2007). This 

framework allows the user to build a site-specific 

project plan based on the relevant management 

questions. The primary focus of this section is on 

monitoring effects from atmospheric pollutants that 

contribute to surface water acidification and 

eutrophication (nutrient enrichment). References that 

provide more details on these measurements are also 

included. Additional sections address protocols for 

quality assurance/quality control (QA/QC) for water 

chemistry sampling, laboratory analyses, data 

analysis, and protocols for sampling and analyzing 

aquatic biota. 

1.1.1 RESOURCES SENSITIVE TO ATMOSPHERIC DEPOSITION

AIR QUALITY-RELATED VALUES 

There are many approaches that can be used by the FS to assess the: 1) current condition of 

surface waters; 2) sensitivity of aquatic natural resources to potential degradation from

atmospheric deposition of S, N, or toxic materials; and 3) extent to which sensitive aquatic 

natural resources have been harmed in the past or might be harmed in the future under scenarios 

of future air pollution and atmospheric deposition. A number of approaches for monitoring 

aquatic resources are highlighted in this protocol; those expected to be most useful to the FS Air 

Resource Management (ARM) program are listed in Table 1-1. This protocol recommends that 

such approaches be routinely considered in making evaluations regarding atmospheric deposition 

sensitivity and/or effects. Site-specific studies can be further customized to fit particular regional 

or local ecosystem conditions and stressors. 

Nutrient Enrichment

In N-limited and co-limited aquatic 

systems, atmospheric N deposition 

can influence algal growth, trophic 

state, and the distribution and 

abundance of diatoms and other 

aquatic species. Lakes and streams 

that are wholly or partly N-limited are 

most likely to occur in remote regions 

with naturally oligotrophic surface 

waters that have not received high 

levels of atmospheric N deposition in 

the past. Within the United States, 

such lakes and streams are most 

common in the mountainous West.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

3

Table 1-1. Recommended sensitive receptors and indicators for air quality related values affected 

by atmospheric deposition of air pollutants. (Source: modified from Sullivan and Herlihy 

2007.)

AQRV Sensitive Receptor Indicator/Metric Potential Criteria*

Flora red spruce (East) growth 

decline

change in diameter

change in extent of damage

sugar maple (East) growth 

decline

change in diameter

change in extent of damage

lichens community composition loss of sensitive taxa

Soil soil chemistry base saturation

exchangeable Ca2+

exchangeable Ca2+ + Mg2+

C:N molar ratio

BS < 10%

% change over time

% change over time

C:N < 0.2

soil solution chemistry Ca:Al molar ratio

[Ca2+ + Mg2+ + K+]:Al molar ratio

NO3

- concentration

Ca:Al < 1.0

BC:Al < 1.0

NO3

- > 20 μeq/L during growing 

season

Water water chemistry ANC (acid neutralizing capacity)

NO3

- concentration

SO4

2- concentration

ANC < 50 μeq/L 

NO3 > 10 μeq/L 

change over time

water productivity chlorophyll a

clarity (lakes)

change over time

change over time

fish salmonid species presence

fish species richness

fish condition factor

fish Hg concentration

fish pesticides(s) concentration

loss over time

change over time

change over time

Hg > 0.3 ppm

above threshold values

zooplankton (lakes) total zooplankton richness

crustacean taxonomic richness

rotifer taxonomic richness

change over time

change over time

change over time

benthic macroinvertebrates (streams) mayfly taxonomic richness

Index of Biotic Integrity

loss of sensitive taxa

deviation from reference

diatoms community composition historical change from 

paleolimnological reconstruction

* Metrics can be represented in multiple ways, often as change over time detected in a monitoring program or as 

exceedence above or below a threshold value. Typically, multiple threshold values are possible. For example, 

surface water target ANC thresholds are commonly set at 0, 20, or 50 μeq/L to achieve different levels of protection. 

Atmospheric deposition can contribute to toxicity responses in aquatic systems in several ways. 

Water acidification entails chemical changes including reduced pH (increased hydrogen ion [H+

] 

activity), decreased acid neutralizing capacity (ANC), increased inorganic monomeric aluminum 

(Ali) concentration, and changed (increased or decreased depending on the extent of acidification) 

concentrations of calcium (Ca2+) and other base cations (BC). Hydrogen ion and Ali can be toxic 

to many aquatic species at sufficiently high concentrations. Other atmospheric pollutants of 

concern include mercury (Hg) and some pesticides. Atmospheric deposition is an important 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

4

component of Hg cycling and biogeochemistry. Mercury in its methylated form is known to 

bioaccumulate in aquatic organisms, reaching potentially high concentrations in larger 

piscivorous fish and species that consume them. 

Air quality related values (AQRVs) are resource elements that may be damaged by air pollution 

or atmospheric deposition. There are many possible sensitive receptors for each AQRV. Sensitive 

receptors monitored for the AQRV water often include its chemistry, which influence its 

suitability to support various aquatic species and life forms. Acid neutralizing capacity is one

indicator of change for the sensitive receptor water chemistry. There are also sensitive biological 

receptors, which reflect the suitability of the lake or stream water for supporting aquatic 

organisms that might be sensitive to acidification or eutrophication. These could include specific 

species of fish, zooplankton, macroinvertebrates, or diatoms. A sensitive receptor can be 

evaluated by measuring indicators of injury or ecosystem change. 

A limited list of key variables does not exist with which to measure ecosystem condition, or 

ecosystem response to stressors, such as those associated with atmospheric deposition (e.g.,

acidification, eutrophication, toxicity). Ecosystems are highly complex and cannot be represented 

by a handful of variables. Nevertheless, some variables are considered reflective of the general 

level of ecosystem harm associated with atmospheric deposition. This protocol proposes a set of 

consistent AQRVs and associated sensitive receptors for aquatic ecosystems that could be used 

by the FS nationwide for evaluation of resource sensitivity to, and effects from, atmospheric 

deposition (Table 1-1; Sullivan and Herlihy 2007). Individual forests may wish to augment these 

listed items to emphasize other AQRV receptors that are especially important to a particular 

forest or region, or for which that forest has specialized expertise. Nevertheless, the 

recommended sensitive receptors summarized in Table 1-1 are broadly applicable and reflect a 

range of aquatic effects of atmospheric deposition. Identification of these receptors and indicators 

helps to determine the protocols that will be needed by the FS ARM program for nationwide 

inventory and monitoring. 

Detailed protocols should be an important part of any resource characterization and/or monitoring 

program intended to evaluate atmospheric deposition impacts on AQRVs. Standard protocols 

help to ensure that measured differences among locations or changes over time at one location 

actually occur and are not a reflection of different methods, sampling personnel, or timing of 

sample collection. Protocols are necessary to ensure that the data collected are appropriate to the 

question(s) asked and are of sufficient quality to allow development of meaningful answers. It 

must, however, be recognized that there will not be a single appropriate protocol in every 

situation that will efficiently characterize an important attribute nationwide. In limited 

circumstances, some attributes and site characteristics are sufficiently variable from region-toregion so that supplemental or amended protocols may be justified. Nevertheless, adoption of 

national-scale procedures for data gathering, analysis, and required core data elements will allow 

data to be compared across forest- and regional-level boundaries and will provide information 

that is needed for national assessments and decision-making. Aquatic effects inventory and 

monitoring for atmospheric deposition effects on FS lands have historically focused on both lakes 

(mainly in the western United States and streams (mainly in the eastern United States). To the 

extent practical, this protocol describes attributes and methods that are applicable to both lakes 

and streams, and that can be applied throughout most or all regions of the United States. This 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

5

protocol addresses the sensitive surface water receptors, and associated field, laboratory, and data 

analysis approaches that are most useful for meeting FS air program objectives.

SENSITIVE CHEMICAL INDICATORS OF WATER QUALITY 

Sensitivity to acidification and its effects are commonly evaluated using several chemical criteria, 

especially ANC, pH, and Ali. Sensitivity of surface waters to chronic and episodic acidification 

depends on watershed soil characteristics, mineralogy, and hydrologic flow paths within the 

watershed (Chen et al. 1984, Cosby et al. 1985), as well as on the current and historic 

atmospheric deposition loadings of acids and bases. Surface water ANC provides an initial 

baseline point from which to quantitatively assess the status of stream or lake chemistry. 

Biological effects of acidification have been associated with a variety of ANC benchmarks, the 

most common of which are ANC’s equal to 0, 20, and 50 microequivalents per liter (μeq/L). 

Waters with ANC ≤ 0 μeq/L have no capacity to neutralize acid inputs; they are acidic by 

definition. Lakes and streams with ANC chronically less than 0 μeq/L are often fishless or 

contain few species of fish. Brook trout, which are relatively acid-tolerant, have been shown to be 

sensitive to episodic acidification1 when chronic stream ANC is less than about 20 μeq/L. A 

general benchmark for sensitivity of other types of aquatic biota is often established at ANC 

equal to 50 μeq/L (Driscoll et al. 2001). Some species may be affected at higher ANC values, 

even at levels of 100 μeq/L or above. Generally, surface waters with ANC ≤ 50 μeq/L are 

considered prone to episodic acidification (DeWalle et al. 1987, Eshleman 1988), especially 

where seasonal snowpack accumulations are substantial. Such low ANC waters may also be 

susceptible to future chronic acidification at current or increased rates of acidic deposition. 

Common reference values for pH are 5.0, 5.5, and 6.0. Such values are often used to evaluate the 

possible extent of adverse effects on fish and other aquatic organisms. Threshold pH levels for 

adverse biological effects have been summarized for a variety of aquatic organisms (Haines and 

Baker 1986, Baker et al. 1990). The effects of low pH are specific to the organism and region 

under consideration and depend upon the concentrations of other chemical constituents in the 

water, notably Ali and Ca2+. Lakes or streams having pH less than about 5.0 generally also have 

ANC less than 0 μeq/L and often do not support fish. Depending on the region, waters having pH 

greater than about 6.5 and ANC greater than about 50 to 100 μeq/L support large, but variable, 

numbers of species. Populations of salmonid fish are generally not found at pH levels less than 

5.0, and smallmouth bass (Micropterus dolomieu) populations are usually not found at pH values 

less than 5.2 to 5.5 (Haines and Baker 1986). A number of synoptic surveys indicate loss of 

species diversity and absence of many other fish species in the pH range of 5.0 to 5.5 (Haines and 

Baker 1986). Levels of pH less than 6.0 to 6.5 have been associated with adverse effects on 

populations of dace, minnows, and shiners (family Cyprinidae), and bioassays suggest that, given 

sufficient Ali concentrations, pH less than 6.5 can lead to increased egg and larval mortality in 

blueback herring (Alosa aestivalis) and striped bass (Morone saxatilis; Klauda et al. 1987, Hall 

1987).

Aluminum (Al) toxicity to aquatic organisms is caused primarily by inorganic forms of Al rather 

than organically complexed Al (Alo); Driscoll et al. 1980, Baker and Schofield 1982, Havas 

 1 Episodic acidification refers to the temporary (usually hours to days) decrease in lake, or especially stream, ANC 

that occurs in response to hydrologic events such as rainfall or snowmelt.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

6

1985). There is limited evidence of biological effects at Ali less than 50 μg/L (~ 2μM). Free Al 

concentrations (Al3

+

; roughly equivalent to Ali concentrations at pH values substantially below 

5.0) between 50 and 200 μg/L have been shown to reduce the growth and survival of various 

species of fish (Muniz and Leivestad 1980, Baker and Schofield 1982). Concentrations of Ali

greater than 200 μg/L are generally considered to have toxic effects to the majority of freshwater 

fish species (Table 1-2).

Table 1-2. General guidelines1 for evaluating the likelihood of aluminum toxicity in fresh water.

Concentration of Inorganic Monomeric Al Expected Response of Aquatic Biota

< 50 μg/L Biological effects not common in most forms of aquatic biota.

50 to 200 μg/L Reduced growth and survival of various species of fish, including 

brook trout, and likely other aquatic life forms.

> 200 μg/L Adverse effects likely for most freshwater fish species.

1 Variability is high with species, life stage, and various aspects of water chemistry, including Ca2+ concentration, dissolved organic 

carbon (DOC), and total fluorine (F) concentration.

Sensitivity of surface waters to eutrophication and the nutrient status of lakes and streams are 

typically evaluated on the basis of concentrations of N and P. These nutrients can be assessed as 

total N and/or total P, or as one or more of the various forms that commonly occur in surface 

waters, such as nitrate (NO3

-

) and soluble reactive phosphorus (SRP). The EPA has provided 

guidance to States for setting nutrient criteria for total N and P concentrations in U.S. lakes (U.S. 

EPA 2000a), and streams and rivers (U.S. EPA 2000b). Different nutrient criteria are being 

developed for each of 14 different nutrient ecoregions throughout the country. Nutrient 

ecoregions are based on aggregations of level III Omernik ecoregions. Draft nutrient guidelines 

are available on the Web at 

<http://www.epa.gov/waterscience/criteria/nutrient/ecoregions/sumtable.pdf>.

In some areas, the concentrations of potentially toxic substances in surface waters may be of 

concern. This issue is likely to be of greatest interest to FS staff in areas downwind of substantial 

emissions sources of pesticides, or where atmospheric deposition of Hg (or other trace metals) is 

known to be elevated. Monitoring of pesticides in surface waters may be advisable on FS lands 

directly downwind of intensive agricultural development. 

Studies of Hg concentrations in fish tissue may be warranted in areas that are downwind of 

known Hg emissions sources, especially where such areas co-occur with probable geologic 

sources of Hg. Regional Hg deposition attributable to long range atmospheric transport is also of 

concern. In general, this protocol does not recommend that the FS ARM program include routine 

monitoring of surface waters for Hg concentrations in water. Nevertheless, Hg is of interest to the 

ARM program because it is atmospherically deposited and it bioaccumulates in aquatic 

environments reaching potentially high concentrations in large piscivorous fish. Mercury can 

pose a health risk to humans or wildlife (e.g., bald eagle, osprey, loon, and river otter) that 

consume large quantities of mercury-contaminated fish. This protocol recommends that a more 

effective way to evaluate Hg contamination in surface waters is to analyze or monitor

concentrations of Hg in fish tissue, rather than in water. Of particular concern are the larger, 

older, piscivorous fish, such as bass, pike, and some species of trout.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

7

POTENTIAL CONFOUNDING FACTORS

In developing and implementing a national field sampling program, it is important to consider 

numerous potentially confounding factors. Some of the important considerations that can 

complicate aquatic inventory and monitoring assessments include:

• Low signal-to-noise ratio2

, especially for dilute aquatic ecosystems;

• Variation in watershed properties such as slope, aspect, underlying bedrock composition, 

extent and mineralogy of glacial till, depth and composition of soils, distribution of 

vegetative cover, role of ground water, and presence and hydrologic connectedness of 

wetlands;

• Interacting stressors especially climate, introduced species, and legacy effects of fire or past 

land use and exposure to pollutants;

• Constraints of sampling in designated wilderness areas where land-use rules prohibit access 

via mechanized equipment and installation of fixed equipment;

• Constraints regarding laboratory analytical holding times;

• National and regional applicability;

• Cost and training constraints; and

• Quality control issues and the need for peer review.

1.1.2 STUDY PURPOSE AND OBJECTIVES

Before selection of study sites and parameters to measure, it is important to determine the purpose 

of the sampling program. For example, the main purpose could encompass any of these:

• Evaluate nutrient limitation;

• Document temporal variability (i.e., diurnal, episodic, seasonal, annual, inter annual) in water 

chemistry;

• Evaluate spatial extent of acid-base status;

• Parameterize interpretive and predictive models3

;

• Determine sensitivity of resources to potential damage; or

• Estimate the magnitude of impact on water quality.

Variation in purpose dictates variation in general approach (Table 1-3), which in turn influences 

the selection of appropriate protocols.

 2 Natural and sampling variability are high relative to the magnitude of change that has occurred in response to 

atmospheric deposition. 3 To apply a process-based effects model to a particular site various input data are needed depending on the selected 

model. Such data might include characterization of soils, hydrology, vegetation, and/or historical documentation 

of land use or atmospheric deposition.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

8

Table 1-3. Common management issues for FS ARM program staff, with associated field study 

approaches.

Purpose General Approach

Determine whether one lake or stream or a 

group of lakes or streams is N-limited for 

algal growth.

Sample water and determine nutrient and chlorophyll a concentrations on multiple 

occasions (at least monthly during the snow-free season) during multiple years. 

Consider also nutrient (N, P) addition experiments in the laboratory and/or field 

enclosures.

Quantify episodic excursions from base-flow 

conditions in surface water chemistry (i.e., 

ANC, pH, Ali, NO3

- concentrations) during 

hydrologic events.

Sample water and measure full ion chemistry during rainstorms, snowmelt, and/or 

rain-on-snow events, with hourly to weekly periodicity. 

Determine the distribution of lake or stream 

water chemistry (i.e., ANC, pH, NO3

-

concentration) across a particular forest or 

wilderness.

Conduct a statistically based or systematic synoptic survey of lake or stream 

chemistry.

Quantify long-term changes in lake or 

stream ANC (or other variables) over time in 

a particular lake or stream.

Sample at least annually (preferably monthly or seasonally during the open water 

season) over a period of at least 8 years. Consider restricting sampling times to 

common hydroperiods or other approaches to standardize timing of sample 

collection among years. Length of time required to continue monitoring to document 

statistically significant changes will depend on temporal variability in water 

chemistry and extent of long-term changes that occur. In general, at least 8 years of 

data will likely be required. 

Determine to what extent air pollution is 

currently affecting the water resources in a 

particular forest or wilderness.

Multiple approaches can contribute to this evaluation:

1) Characterize index chemistry for multiple lakes and/or streams expected to be 

highly sensitive.

2) Conduct a synoptic survey (preferably using a stratified random selection 

process) of waters in the forest/wilderness.

3) Use a dynamic process-based watershed model to hindcast past changes in 

acid-base chemistry.

4) Collect and analyze diatom remains in a sediment core from the deepest part 

of one or more of the presumed most acid-sensitive lakes.

5) Use a steady state or dynamic process-based-watershed model to quantify 

the critical load of S and/or N deposition.

Evaluate whether the current condition of 

acid or nutrient sensitive waters warrants 

mitigation.

Multiple approaches can contribute to this evaluation:

1) Characterize index chemistry for multiple lakes and/or streams expected to be 

highly sensitive.

2) Conduct synoptic survey (preferably using a stratified random selection 

process) of waters in the forest/wilderness.

3) Use a dynamic process-based watershed model to hindcast past changes in 

acid-base chemistry.

4) Collect and analyze diatom remains in sediment cores from the deepest part 

of one or more of the presumed most acid-sensitive lakes.

5) Use a steady state or dynamic process-based-watershed model to quantify 

the critical load of S and/or N deposition.

6) Use a dynamic process-based-model to evaluate likely future responses to 

reduced levels of acidic deposition. 

The management needs that the field study is intended to address will help determine the type of 

field study that is most appropriate. The management needs will lead into a series of questions, 

which in turn will guide the sampling effort:
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

9

• What kinds of sampling are required to support the management needs?

• What are the protocols to meet those sampling requirements?

• What are the standard operating procedures (SOPs) to implement those protocols?

In designing the field study, there are 10 basic questions to consider. Each of these questions 

should be addressed to avoid the risk that the sampling program will fail to yield the data required 

to meet the program’s needs. This protocol provides guidance regarding how-to answer these 

questions (modified from <http://www.epa.gov/owow/monitoring>):

1. Why is the sampling taking place?

2. Who will use the resulting data, and how will that influence the level of quality 

assurance that will be required?

3. How will the data be used, and how will the intended use influence data requirements?

4. What parameters or conditions will be measured?

5. How good does the data need to be in terms of accuracy, representativeness, 

completeness, and intrasite and intersite comparability?

6. What methods should be used?

7. Where are the sampling sites?

8. When will the sampling occur?

9. How will the data be managed?

10. How will the program ensure that the data are credible?

The most important aspect of any inventory and monitoring plan is specification of the objectives 

and questions to be answered using the resulting data. Once the objectives and questions are 

conceived and refined and some preliminary data are collected with which to evaluate data 

variability issues, it is possible to specify a plan that will have a high probability of success. The 

greatest challenge in developing a monitoring or synoptic survey plan is asking the most 

appropriate questions. It is important to decide what you want to know and what uncertainty you 

are willing to accept in your answers. Many field sampling programs are compromised from the 

beginning because they were not specific about what questions the program was intended to 

answer. Specificity regarding the questions can lead to specificity regarding the sampling design 

and result in the collection of data suitable for providing the desired answers.

Because it is not possible to sample at all locations at all times for all parameters, it is important 

to consider in advance how to make the best choices regarding expenditure of limited funds for 

field sampling. The most important aspect of sampling design is setting specific objectives, and 

linking the objectives to specific questions. The questions should consider elements of subject, 

location, time, trend, and degree (Table 1-4).
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10

Table 1-4. Elements to be considered in formulating sampling questions.

Element Example

Subject Stream NO3

- concentration.

Location Spring Creek, 50 m below its confluence with Sparks Creek.

Time During spring snowmelt.

Trend Is stream NO3

- concentration increasing from year-to-year during the spring high flow period?

Degree Is it changing by a statistically significant amount, or a biologically meaningful amount?

Population of Interest First through third order (at 1:100,000 scale) streams in the Blue Ridge ecoregion in Virginia.

A well-conceived plan for water quality sampling should be:

• Relevant to the beneficial uses of the waters;

• Specific with respect to sampling locations, depths, parameters, schedule, and methods;

• Consistent with approved methods;

• Specific with respect to recommendations for quality assurance, data analysis, and reporting;

and

• Designed to maintain continuity to the extent possible with the existing sampling efforts, 

especially if trend analyses will be conducted using the data.

Within the context of characterization and monitoring studies to measure or document air 

pollution effects on surface waters, there is a multitude of questions that could be an appropriate 

focus for field studies. A partial list is given in Table 1-5. Selection of the most relevant questions 

depends to a substantial degree on location. Key questions can be influenced by the extent of 

historical acidic and nutrient deposition, inherent sensitivity of the resources present, hydrologic 

characteristics, types of aquatic resources of greatest interest (e.g., drainage lakes, seepage lakes, 

low-order streams, moderate-order streams), topography, as well as many other factors. 

Monitoring of lake and stream water quality is performed to provide resource managers with 

information on possible water quality problems that may require intervention to determine the 

susceptibility of lakes and streams to potential stressors and to document changes (improvement 

or deterioration) in key parameters of interest or in known problem areas. For example, FS 

managers should know surface water sensitivity to acidification when they review emissions 

permit applications. Information from a well-designed and properly-executed monitoring plan 

may also allow evaluation of the effectiveness of emissions controls or other best management 

practices and the potential need for other actions that might be warranted.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

11

Table 1-5. Examples of questions that could be used to guide inventory, characterization, and 

monitoring study design.

Subject Question

Inventory What is the distribution of lake water ANC (or alternatively pH, inorganic monomeric Al, Ca, NO3

-

, or 

SO4

2-

) across high-elevation lakes in a particular Wilderness area?

What is the annual average (or index) water chemistry of the most acid-sensitive streams in a particular 

National Forest (expressed as 5th percentile of sensitivity of the population of streams, or the five most 

sensitive streams known to exist in the forest)?

What are the concentrations of stream water NO3

- (or ANC, pH, Ali) during snowmelt at selected longterm monitoring locations in a particular National Forest, and how do they compare with summer or fall 

index NO3

- concentration in these streams? 

Characterization What is the extent of episodic chemical change (decrease in ANC, pH; increase in inorganic monomeric 

Al, NO3

-

) during the peak of snowmelt at selected long-term monitoring stream sites?

What landscape characteristics (i.e., lithology, soil type, elevation, ecoregion, stream order, etc.) are 

associated with the occurrence of streams having spring-base flow ANC below 50 μeq/L within the 

National Forests of NC, TN, and SC?

Monitoring What is the long-term trend in lake water NO3

- (or other variable) concentration for FS long-term 

monitoring sites in the Rocky Mountains over the period of monitoring since 1990, as measured during 

the summer index period, and what are the characteristics of the sites that show the largest positive 

trends?

Given the observed temporal variability in spring-base flow ANC in a particular stream, how long would 

monitoring need to be conducted to document a statistically significant increase in stream ANC if the 

average actual increase in ANC was 1 μeq/L/yr?

Do long-term trends in spring-base flow stream water Ca2+ concentrations in second- and third-order 

streams in XYZ Wilderness area since 1990 suggest the potential for Ca-deficiency in the soils of 

higher elevation forests in this wilderness? 

1.2 STUDY DESIGN

Water quality studies for evaluating aquatic effects of atmospheric deposition are most commonly 

designed as lake or stream characterization studies, synoptic surveys of the chronic chemistry of 

lakes or streams in a particular forest or region, characterization of episodic variations in 

chemistry in response to rain storms and/or snowmelt, or long-term monitoring studies to 

document and quantify changes in chemistry over time. Each type of design is described below. 

In selecting an appropriate design, determine in advance precisely what you would like to know. 

Subsequently, you can determine the type of study design that will be most useful (Table 1-6).

One of the most important, and most frequently overlooked, aspects of study design is that it 

should incorporate the data requirements of the statistical procedures that will be used to analyze 

the data. Consulting with a statistician can aid in the development of a study design. In addition, 

you should determine what you intend to do with the data before beginning study design. 

Study design should also coordinate with existing efforts by other FS programs, as well as other 

governmental agencies, where possible. In some cases, another FS program, the U.S. 

Environmental Protection Agency, the U.S. Geological Survey, or a State agency may have 

ongoing or planned sampling programs that overlap with the FS Air Program projects. This 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

12

coordination effort may be as simple as collecting some additional data that might be shared, or 

pursuing joint funding of a desired sampling effort. 

Table 1-6. General guidance regarding water quality study design.

If what you want to know is: You should consider the following kind(s) of study design:

Number of lakes, length of streams, or percent of the 

regional population of lakes or stream length that is 

above or below a particular criteria value (i.e., ANC 

≤ 50 μeq/L).

Some form of stratified random sampling that will allow extrapolation of 

results from individual sites to the larger area.

Status of the acid-base chemistry of the most (or 

some of the most) sensitive lakes or streams in an 

area.

Non-statistical survey of selected lakes and/or streams in portions of the 

study area and landscape positions expected to contain the most sensitive 

aquatic resources.

General assessment of lake or stream chemistry in 

an area, with identification of some of the more 

sensitive water bodies in the area.

Statistical or non-statistical screening of a relatively large number of water 

bodies across the expected gradient of sensitivity, measuring specific 

conductance and/or pH in the field for making a rough assessment of 

condition, and collecting samples for full laboratory analyses for a subset of 

those samples.

Estimate of seasonal or episodic variability in the 

chemistry of an acid-sensitive lake or stream.

Frequent interval sampling during the period of interest. Sampling can 

range from hourly to monthly during the season or period of interest and 

can include multiple years to capture the range of variation. 

Analysis of long-term changes in water chemistry

over time. 

Periodic sampling (usually monthly to annually) over a period of usually at 

least 8 years focused on an index period or standardized by hydroperiod. 

More robust studies (with greater statistical power to detect trends) will 

entail more frequent sampling (weekly to seasonally) and/or will extend for 

longer than 8 years. 

Assessment of temporal variability in water 

chemistry of a particular lake or stream.

Frequent interval (hourly to seasonally) sampling that captures major 

changes in hydrology during the season(s) of interest. Should include 

multiple years. 

Determination of whether and to what extent water 

resources in a particular forest or wilderness have 

been adversely affected by atmospheric deposition 

to date. 

Multiple designs will be needed using a weight-of-evidence approach. They 

might include:

• Synoptic survey (statistical survey preferred)

• Characterization of multiple representative sensitive lakes and/or 

streams

• Long-term monitoring

• Assessment of seasonal and episodic variability

• Hindcast chemistry using dynamic process based model(s).

Determination of the prognosis for future recovery of 

damaged aquatic resources or quantification of the 

atmospheric deposition levels that will be protective 

of sensitive resources.

Model scenario and critical loads analysis.

1.2.1 LAKE OR STREAM CHARACTERIZATION

There are multiple approaches to characterize the chemistry of a lake or stream. Some studies are 

based on only one or a few samples. Most commonly, these are collected as index samples. 

Decisions should be made concerning the frequency and timing of sampling. Springtime baseflow samples are often regarded as a good representation of annual average flow-weighted stream 

water quality in the southeastern United States when only single samples can be collected. 

Summer or fall index chemistry (commonly avoiding large rainstorm events) is often regarded as 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

13

a good representation of annual average lake-water chemistry. Lake sampling after fall overturn 

can yield results for fully mixed conditions, but may require measuring the lake temperature 

profile to verify that turnover has occurred. Selection of an index period has implications for the 

temporal stability of the water quality and for the degree of impact that might be revealed by that 

water quality. Water quality is more likely to be stable (and thus comparable among water bodies 

if a survey is conducted of multiple lakes or streams) during summer and fall. However, in many 

regions that contain acid-sensitive waters, the lowest pH and ANC and the highest Ali

concentrations are more likely to occur during spring.

Better representation of annual conditions in both streams and lakes can be obtained with 

seasonal or other periodic sampling, as opposed to collection of only one sample to represent a 

given year. Selecting a sampling period when flows are low and least variable may provide data 

that are generally comparable among a group of waters or in a given water body between years, 

but may not represent the conditions of interest. For example, surveys to assess the effects of 

acidic deposition on surface water chemistry can substantially underestimate impacts if low-flow 

periods are used for sampling (Lawrence et al. 2008b). Some studies endeavor to collect one or 

more samples during high-flow periods (heavy rain or snowmelt) to augment index chemistry 

sampling. High-flow periods tend to cause:

• Relatively high NO3

-

, Ali, and P concentrations;

• Relatively low pH and ANC; and

• Variable concentrations of SO4

2- and base cations (depending on local characteristics). 

Sampling to Support Modeling

If modeling is anticipated using the collected data, consider which water chemistry variables 

will be needed to calibrate the model. For example, application of the MAGIC model (Cosby 

et al. 1985), sometimes used in acidic deposition assessments, requires analyses of nine 

stream water constituents: 

• pH;

• Three acid anions: sulfate, nitrate, and chloride;

• Four base cations: calcium, magnesium, potassium, and sodium; and

• Ammonium.

It is desirable, but not necessary, to also provide Gran titrated ANC. If modeling is to be 

done to estimate future pH, then it will also be necessary to measure dissolved organic 

carbon, and if pH is below 5.5, to determine total monomeric aluminum and organic 

monomeric aluminum. If pH is below 5, measure total fluoride, which will be needed to 

calculate aluminum speciation. Thus, the additional analyses needed to support MAGIC 

modeling of future pH (in addition to those listed above as required) include:

• Gran ANC;

• Dissolved inorganic carbon;

• Total monomeric Al; and

• Organic (non-labile) monomeric Al.

These additional parameters are not needed to support MAGIC modeling of future ANC. 

Other models may require different or additional parameters.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

14

The most stressful conditions (to aquatic biota) generally occur during high-flow periods. The 

range of difference between high-flow and low-flow chemistry varies by region, with drought 

cycles, and by individual water body. In general, this protocol recommends that characterization 

of lakes and streams be represented by one or more index samples for a given year, plus at least 

two additional samples during snowmelt or rainstorms to partially characterize variability. It is 

preferable to collect data for at least 2 years and to document inter-annual variability associated 

with wet/dry cycles. 

1.2.2 SYNOPTIC SURVEY

Synoptic surveys of lake or stream chemistry within a forest or within a designated region are 

usually conducted at times expected to exhibit fairly stable water chemistry. For acidic deposition 

monitoring, this is usually spring base flow for streams in the Southeastern United States, and the 

summer or fall index period4 for lakes or streams in regions that typically develop substantial 

snow pack. For backcountry sampling within the FS ARM program, typically one sample with 

replicate(s) is collected for each lake or stream that was selected for sampling. Synoptic surveys 

are ideally, but not always, statistically based, which allows for extrapolation of sample results 

from individual water bodies to the regional population of interest. At the time of sample 

collection, standardize for consistent weather and runoff conditions as much as possible at each 

site. Periods of high temporal variability, such as heavy rain and periods of rapid snowmelt, or 

periods with heavy smoke from wildfires, are typically avoided during a synoptic survey. 

However, if assessment of acidic deposition effects is the goal, avoiding high-flow conditions can 

result in a substantial underestimation of the magnitude of impact. The extent of this 

underestimate can be quantified by conducting additional seasonal and episodic sampling for at 

least a subset of the sampling sites.

1.2.3 LONG-TERM MONITORING

Long-term monitoring of stream or lake chemistry usually involves collection of water samples at 

regular intervals from weekly to quarterly or even annually, with the primary purpose to detect 

trends that reflect an environmental change over time. How quickly a trend can be detected 

depends on the strength of the trend (the rate of change) and the amount of intra annual and inter

annual variability in the water chemistry. It is generally possible to detect a change of slight 

magnitude under conditions of low variability and with a longer period of record. The likelihood 

of detecting a significant trend in the concentration of a given water chemistry variable will be 

determined in large part by the length of the monitoring period. In the event of a small to 

moderate change in chemistry, it may take 10 to 20 years or more of monitoring data to document 

a significant change. 

An effective monitoring plan stems from a series of questions and constraints that sequentially 

focus the plan into specific elements that are well-defined and unambiguous. Because information 

is gained during implementation of a monitoring plan, it is often desirable to revisit plan elements 

to continuously refine and update the monitoring activities. In addition, external factors such as 

 4 Index period is a relatively narrow period of time for synoptic sampling (often a 2-month window during either 

spring, summer, or fall) intended to represent the lake or stream chemistry for that year. Typically, rain or 

snowmelt conditions are avoided when collecting index samples.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

15

changes in monitoring technology, analytical methods, and regulations will often impinge on the 

design and execution of the monitoring. For these reasons, routine (e.g., annual) reviews of the 

results and methods should be incorporated into the monitoring plan. However, if trend detection 

is one component of the plan, care should be exercised before making changes to the program 

that might compromise the integrity of the data and the ability to use earlier data to infer 

statistically significant changes in water quality.

1.2.4 CHARACTERIZATION OF STREAM CHEMISTRY DURING

HYDROLOGIC EVENTS

Hydrologic events, high flows caused by rainstorms and/or rapid snowmelt, are episodic and can 

last from a few hours to a few weeks. Although these events occur over a relatively small fraction 

of the year, they often represent the majority of total annual flow (Likens et al. 1977) and 

constituent flux. Events can happen throughout the year in most regions of the United States but 

are most common during seasons of high precipitation and during spring snowmelt in regions 

where snow accumulates and are generally least common during summer when high 

evapotranspiration reduces soil moisture. 

Stream-water chemistry during hydrological events is important to characterize because high 

flows often lead to extreme chemical conditions. Effects on stream biota from episodic variations 

in stream chemistry can be as severe as chronic effects associated with base-flow chemistry. For 

example, episodic acidification can result in the elimination of an annual age class of fish 

(McComick and Leino 1999) when the event occurs during sensitive life stages. Episodic 

acidification can also affect other forms of aquatic life, such as diatom communities, which have 

been found to be less diverse in an episodically acidified stream than in a nearby chronically 

acidified stream (Passy et al. 2006). 

Manual water sampling is generally not effective at characterizing chemical variability over the 

course of a hydrologic event because the timing and shape of the hydrograph is difficult to predict 

and may occur at inconvenient times for the sampler. Automated water sampling triggered by 

changes in water level, often referred to as stages, provides an effective solution to this problem. 

Automatic samplers can collect water at predetermined time intervals or at intervals based on the 

rate of change in water level. Samples collected during events can then be evaluated using the 

flow measurements recorded at the times of sample collection to optimize the selection of 

samples for chemical analysis. This approach offers the opportunity to greatly reduce analytical 

costs with minimal loss of information. However, the use of automatic sampling equipment is 

restricted in wilderness.

1.2.5 OTHER USES OF RESULTING DATA

Surface-water-quality data can also be used to support process-based modeling studies using a

watershed model such as MAGIC or PnET-BGC. Such models can be used to hindcast pre

industrial chemistry to determine whether and to what extent a lake or stream has acidified since 

the Industrial Revolution. A second general approach is to conduct scenario modeling to estimate 

future changes in water chemistry in response to one or more scenarios of emissions control and 

deposition. A third modeling approach is simulation of the critical and/or target loads of 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

16

atmospheric deposition to protect or restore acid-sensitive or nutrient-sensitive aquatic resources 

(see side box on Critical and Target Loads). All of these modeling approaches require 

compilation of model-input data. These can include, depending on the selected model, data on 

soil and water chemistry, hydrology, vegetative characteristics, and estimates of historic and 

current deposition.

1.3 WHERE, WHAT, AND WHEN TO SAMPLE

Data that provide information on the quality of surface water can be used to evaluate the 

following kinds of issues:

• Short-term episodic changes (scale ~hours to days) in water quality;

• Long-term chronic changes (scale ~years to decades) in water quality;

• Types of water quality changes;

• Likely cause(s) of water quality changes;

• Longitudinal variation of water quality along streams or depth variation within lakes;

• Status and extent of chemical and biological condition across populations of lakes and 

streams; and

• Biological effects of water quality changes.

The ability to assess these issues will be limited largely by the extent and intensity of the 

sampling effort. Three of the most critical aspects of water-quality-sampling design (cf., Green 

1979) include:

• Stating the questions to be addressed concisely and precisely;

• Conducting a preliminary pilot sampling; and

• Replicating sampling in time and in space.

The questions to be addressed will come from the project objectives, which should be agreed 

upon in advance of field sampling. 

Sampling can be conducted at different intensity levels depending on study design, questions to 

be addressed, and the intended use of the resulting data. The level of intensity will influence 

decisions about how, where, what, and when to sample and will affect associated quality 

assurance, laboratory analysis, data analysis objectives, and standard operating procedures. For 

example, if the objective is to gain a general understanding of the distribution of potentially acidsensitive streams within a particular wilderness, then a low level of sampling intensity may be 

acceptable. This may, for example, entail only a summer survey of specific conductance, pH, 

and/or ANC, with no additional measurements. If, however, the objective is to more fully 

characterize the acid-base chemistry of one or more streams or lakes or to quantify long-term 

trends in water chemistry, then a higher level of intensity will be required. Sampling that is 

intended to support regulatory decision-making or that will likely be used in permitting or 

litigation demands the highest level of intensity. There is not a one-size-fits-all approach to 

establishing water sampling protocols. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

17

The ability to determine the existence of a statistically significant trend in water quality over time 

is influenced by the: 1) magnitude of change that occurs in the parameter and water body of 

interest; 2) temporal variability that occurs in that water quality parameter and water body; and 3) 

the number and temporal distribution of samples collected. To design a monitoring plan to detect 

the existence of a statistically significant change in lake or streamwater chemistry parameters, the 

level of change that one wishes to be able to detect, in conjunction with the known or expected 

temporal variability of the parameter in that lake or stream must be considered. Before initiating a 

monitoring effort that is intended to evaluate change over time (trends detection), it is helpful to:

1) consult with the person who will be responsible for the eventual statistical analysis of the 

resulting monitoring data or someone knowledgeable in statistics and 2) conduct a pilot study to 

determine the temporal variability that occurs in the parameter(s) of interest in that water body or, 

at a minimum, in a water body thought to be similar in its chemistry and temporal variability. 

The overall quality assurance objectives for a water-quality-sampling project are 1) to implement 

quality control procedures and requirements for field sampling and laboratory analysis that will 

provide data of sufficient quality to achieve the program objectives and 2) to follow procedures 

that will provide data of known quality in terms of precision, accuracy, completeness, 

representativeness, and comparability. Quality assurance and quality control (QA/QC) issues are 

covered in detail in the QA/QC section (Section 3) of this report. It is important to note that 

certain aspects of the QA/QC protocols that are adopted for a particular study will affect choices 

made in designing the sampling program. In particular, it is important to determine, in advance of 

initiating field work, what the data quality objectives (DQOs)will be with respect to the selected 

Critical and Target Loads

Modeling of critical and/or target loads requires that a number of decisions be made before 

initiating modeling. These decisions determine what resources are to be protected, at what 

level, and over what time period. Sensitive resources to be protected by a given critical or 

target load can include fish or other types of aquatic biota, or various terrestrial resources 

such as vegetative or soil condition. 

To protect these resources, one or more chemical indicators are typically chosen. Often, ANC 

is used as the indicator for protecting aquatic resources from acidification. One or more 

critical ANC levels must be selected (e.g., ANC 20 or 50 μeq/L), typically in association with 

known or suspected dose/response relationships for various sensitive species. Different 

critical ANC levels are expected to protect different species of aquatic life. 

Finally, one selects a steady-state approach or specifies the time period over which the

sensitive resources are to be protected or over which the damaged resources are expected to 

recover. Steady-state critical loads are determined irrespective of time. Dynamic critical loads, 

or target loads, may be determined for various endpoint years, for example, 2050 and/or 

2100. 

Each decision that must be made to simulate critical or target loads has an influence on the 

resulting model-simulated values. A target load can be selected that is higher than the 

modeled critical load if the objective is to make some limited progress towards reaching the 

critical load. Conversely, a target load can be selected that is lower than the critical load to 

ensure that the sensitive ecosystem is fully protected given modeling uncertainty or to attain 

the targeted threshold chemistry more quickly for resources that have already been damaged.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

18

targets for analytical detection limits, precision, accuracy, and completeness. In addition, plans 

need to be made concerning how many and which of the field samples to be collected will be 

replicated in the field. Some sampling programs replicate all samples in the field. 

1.3.1 WHERE TO SAMPLE

SELECTION OF SAMPLING LOCATIONS

Selection of sites for water quality sampling should be based on systematic and documented 

criteria. One of the most important criteria is having a well-defined population of interest. The 

criteria should be chosen with consideration of watershed factors. These can include 

representation of bedrock, soil type, geographic distribution of surface waters, presumed 

sensitivity to stressor(s) of interest, elevation, watershed area, lake area or stream order, site 

accessibility, and avoidance of watersheds with other elements of human disturbance that might 

influence surface water composition. Approximate sampling site locations can be identified on a 

preliminary basis from examination of available mapped data before initial sampling trips, with 

specific site selection and further documentation developed in association with sampling. 

Features of Landscape Associated with Acid-Sensitivity of Surface Waters

It is not possible to define features of the landscape that will be most closely associated with 

surface water acid sensitivity nationwide. Such relationships are highly variable across the 

landscape and vary from region-to-region, and often from forest-to-forest. Nevertheless, for a 

given region or forest it is often possible to identify certain landscape features, such as lithology, 

elevation, watershed area, and watershed slope, which correlate with acid sensitivity or effects. 

Sometimes vegetation type (e.g., coniferous forest, alpine, and subalpine vegetation), soil type, or 

one or more regional soils variables (e.g., pH, depth, or percent clay fraction) may be helpful. 

Ecoregion designations incorporate many of these variables and can also be useful. In general, the 

most acid sensitive lakes and streams are expected to occur under the following conditions:

• Bedrock that is not basaltic and does not contain appreciable amounts of carbonate;

• Relatively high elevation;

• Steep terrain;

• Small watershed;

• Thin soils;

• Low soil clay content and low soil pH; and/or

• Flashy hydrology.

In the Southeastern United States, acid-sensitive streams are often associated with siliciclastic 

bedrock lithology (cf., Sullivan et al. 2007). In glaciated regions, the presence of varying amounts 

of glacial till can obfuscate relationships between lithology and surface water chemistry. Acidsensitive lakes and streams often, but not always, occur at relatively high elevation, on steep 

slopes, and in relatively small watersheds. Knowledge of such relationships, especially if 

regionally specific, can aid in selection of surface waters for inclusion in inventory and 

monitoring programs. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

19

Random Versus Non-Random Site Selection

One of the most important considerations in site selection is the determination of whether or not 

the sampling sites will be selected using a randomized sampling design. Streams or lakes should 

be randomly selected for sampling if the goal is to characterize populations of surface waters for a 

defined area too big or impractical to census. Doing so enables the statistics obtained for the 

sampled waters to be applied to the full population of waters in the designated sample frame 

within the area. 

For a statistically based survey of surface waters some form of stratified random sampling is 

generally used as this approach allows the sample population to be stratified such that the streams 

or lakes of greatest interest can be included in amounts that are disproportionate to their 

frequency of occurrence. Such a stratified random sampling process preserves the ability to make 

population-level extrapolations while 

maximizing the collection of data for 

the sites of greatest interest. A 

carefully targeted and stratified 

random sampling does not necessarily 

have to entail a large and expensive 

sampling program. 

Random surveys of aquatic resources 

conducted by EPA have often been 

large efforts that sampled hundreds to 

more than a thousand water bodies. 

These have included the Wadeable 

Stream Survey, National Lake 

Assessment, National Surface Water 

Survey, and various surveys 

conducted as part of the 

Environmental Monitoring and 

Assessment Program (EMAP). 

Nevertheless, smaller surveys could 

also be conducted using a random 

sampling structure, thereby allowing 

extrapolation to a population of waters 

of particular interest. 

If all streams or lakes are included for 

potential sampling, accessibility may 

complicate a totally randomized 

sampling design. Remote sites may 

require extended periods of time to 

reach, which lengthens the period 

over which the survey is conducted and may introduce complications regarding sample-holding 

times and costs. This can also be problematic because environmental sampling conditions (e.g.,

stream flow) may vary during the survey if some of the sites take several days to access. Data 

Nutrient Status

A rough evaluation of the nutrient status of a lake or 

stream can be made on the basis of the molar ratio of 

total N to total P in solution. This determination was 

formerly based on the Redfield N:P ratio equal to 16. 

More recent compilations of experimental data (cf., 

Elser et al. 2009) suggest a cutoff near 44 for N 

versus P limited fresh water lakes. 

If the ratio is above about 44, the water body is 

presumed to be P-limited, and further addition of N 

would not be expected to have a large effect on 

primary productivity. If the ratio is below about 44, the 

water body is presumed to be N-limited and therefore 

may be sensitive to nutrient enrichment effects from 

N addition. 

Such a rough evaluation should be based on multiple 

samples (at least 10) collected across multiple 

seasons (ideally spring through fall), as the nutrient 

status can change with season and/or with short term 

changes in flow or other conditions. Because this is 

an area of active research, such interpretation should 

be considered uncertain and subject to change. 

A more complete evaluation of nutrient status should 

be based on laboratory, and perhaps also in situ

nutrient addition experiments that add N and P, 

individually and combined, to laboratory flasks or in 

situ enclosures, and then measure nutrient 

concentrations and chlorophyll a or some other 

measure of primary productivity.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

20

collected from surface waters sampled during low-flow conditions are generally not comparable 

to those collected during high-flow conditions. 

Within a randomized sampling design, candidate sampling sites can be stratified according to 

accessibility. This can help ensure that a limited number of remote sites will be included. 

However, use of this approach can result in lower precision in quantifying conditions of remote 

sites. 

Selection of streams or lakes for sampling may be non-random if information is needed on 

specific waters or watersheds, rather than on a population of waters or watersheds. Non-random 

sampling may be acceptable for certain studies. Nevertheless, one should carefully consider that 

the gains realized in ease of sampling and/or availability of data collected previously in a nonrandom study must be weighed against the loss of the ability to extrapolate conclusions directly 

from the sampling sites to the population of interest. 

For non-random sampling, especially if the lake or stream is intended to be part of a long-term 

monitoring effort, it may be desirable to select water bodies that exhibit particular characteristics. 

For example, it makes little sense to spend years monitoring a body of water that is not acidsensitive if the objective is to evaluate acidification response. Thus, one might purposely select 

one or more highly sensitive sites for monitoring or for detailed study. Similarly, it would be 

illogical to focus a study of atmospheric nutrient N enrichment on a surface water body that is Plimited. 

For certain studies, it is logical to select sites that are highly sensitive to the stressor in question. 

Nevertheless, it can be difficult to interpret the results of such studies without first determining 

where the studied sites fall within the distribution of site sensitivities across the forest or across 

the region. Such distributions of regional site characteristics can sometimes be provided by 

various statistically-based large synoptic sampling programs such as EPA’s Wadeable Stream 

Survey, National Stream Survey, National Lake Survey, or EMAP. 

Statistically-based survey data can be used to aid in selection of sites for long-term monitoring. 

Long-term monitoring sites may be chosen at random from among randomly selected survey sites 

so that the resulting monitoring data will be representative of the entire population of interest. 

This approach was taken in EPA’s Temporally Integrated Monitoring of Ecosystems (TIME) 

project (Kahl et al. 2004). 

Connection of survey or monitoring sites to the broader regional population of lakes or streams is 

always highly desired. This connection allows extrapolation, whether statistically or semiquantitatively, of results to more bodies of water than just the one(s) sampled. Ideally, study sites 

should be statistically selected. If this is not possible, it may be possible to express the results for 

a given study site relative to the broader population by quantifying its chemistry relative to the 

population of lake or stream chemistry determined in one of the larger regional or national 

surveys, such as those conducted by EPA. Alternatively, the feasibility of conducting a synoptic 

survey targeted to the specific forest or region should be considered. Such a survey could range, 

depending on resource availability, from sampling a few variables to a study of full water 

chemistry. A screening survey to identify candidate sites for further study could be focused 

mainly on such parameters as specific conductance and field pH, with full laboratory chemistry 

conducted on only a subset of the sites. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

21

Candidates for Inclusion

Before conducting either a random or non-random survey of lakes or streams, determine what 

kinds (classes) of lakes or streams should be included as candidates for sampling. Pre-selection of 

classes of water bodies to include may change the population frame in a statistically based 

sampling or change the waters that are candidates for selection in a non-random design. 

Candidate lakes might be restricted by hydrologic type (to drainage lakes or seepage lakes, for 

example), lake size, topographic position, ease of access, or depth. Candidate streams might be 

restricted to certain stream orders5

, or otherwise constrained according to watershed area, 

elevation, presence/absence of fish, or presence or absence of watershed disturbance. For nonrandom surveys, intended to identify and characterize the most acid-sensitive surface waters in a 

particular region, forest, or wilderness, this protocol recommends focusing on the following types 

of lakes and streams:

• Perched, seepage lakes;

• Small (less than about 50 to 100 ha) drainage lakes occupying relatively high landscape 

position and having average depth greater than about 1 m6

; and

• Low-order streams (first through third order).

In some cases, a systematic approach to pre-selection of sampling sites may reduce the number of 

candidate sites to such a degree that all or most of the high-interest candidate sites can be 

sampled. 

Selection of Specific Sampling Sites

The sampling site in a lake is generally selected on the basis of logistical considerations. The 

preferred site is the deepest area of the lake, but this requires use of a boat, raft, or float tube. If it 

is not possible to sample at the deepest area of the lake, then use the lake outlet or, as last choice,

a shoreline location near the outlet.

A number of factors should be considered when selecting the specific sampling point. Any 

sampling point along a stream will be affected by features of the upstream watershed. The 

following is a list of landscape features that can affect water quality:

• Impoundment structure;

• Wetland;

• Tributary-stream junction;

• Dramatic change in slope;

 5 Stream order refers to a system of classifying streams based on their branching pattern. The smallest headwater 

streams are first order. When two first-order streams come together they form a second-order stream. As more firstorder streams flow into the second-order stream, its order is not affected; it is still second order. When two secondorder streams combine, it becomes third order. The process continues to progressively higher orders. The scale of 

the mapped data used to designate stream order has influence on the classification. Most acid-sensitive streams tend 

to be relatively low order (often first through third order at 1:100,000 scale or first through fourth order at higher 

resolution). 6 Lakes that are less than about 1 m deep grade into wetlands. Some studies of lakes only include those deeper than 

1 m.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

22

• Abrupt change in vegetation, soil type, bedrock type;

• Groundwater discharge (spring);

• Upslope disturbance (fire, mining, camping areas or trails, logging, heavy grazing, 

windthrow); and

• Upslope human activity (agriculture, residential development, road building).

Specific information on the watershed is necessary for determining if the stream is appropriate for 

sampling a particular sensitive receptor, and, if so, where the sampling point should be located on 

the stream reach. The upstream drainage should also be assessed to determine if disturbances,

such as fire, mining, or logging, have occurred or if the stream has been influenced by erosion 

from the stream bank or from adjacent roads or land disturbance. In addition, the presence of 

other human activity in the watershed, such as agriculture or residential development, may affect 

downstream water quality.

Proximity to trails or roads can be considered in selecting a sample location. Roads and trails 

provide accessibility and a bridge can be used for sampling larger streams and taking flow

measurements. If there is an existing stream gage in the area of interest, co-locating the stream 

water sampling site with the gage will provide flow data that would be valuable for interpreting 

the chemistry data.

As described above, the general sampling location can, and should, be specified in advance of 

sending sampling personnel into the field. However, the precise sampling location can be selected 

by the field personnel when a site is first sampled within a non-random survey or monitoring 

program. Random stream sampling requires that the crews sample at the specified random 

sampling point; if that is not possible, then the site is classified as Not Sampled and the portion of 

the population that it represents is categorized as Not Assessed. 

Any subsequent sampling of a given site should rely on global positioning system (GPS) 

coordinates, site photographs, detailed maps, and written description of the site location to return 

to precisely the same location each time that site is sampled. Where allowed, if a site is intended 

to be sampled repeatedly, the placement and documentation of uniquely numbered metal tags at 

the base of a tree or on a rock adjacent to the sampling site can provide confirmation of site 

location.

The field crew should follow these guidelines in selecting new non-random sites for repeated 

long-term sampling:

• The best point to sample a lake is over the deepest area of the lake. This requires use of a 

boat, raft, or float tube. If it is not possible to collect such an open-water sample, the next best 

option is to sample the largest flowing outlet from the lake; the outlet should be sampled, 

using stream sampling procedures, as close to the lake as is practical. The third, and least 

desirable option, is to sample the lake from the shoreline. Shoreline sampling should be 

conducted, if possible, from a large rock or by wading a short distance from the shore. Care 

must be taken to avoid disturbing the sediment in any way that could affect the quality of the 

sampled water. Proximity to logs and aquatic vegetation should be avoided. If possible, use 

wind currents to advantage by collecting the sample from an area that receives wind-driven 

surface water movement from the larger lake. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

23

• The best point to sample a stream is where the water is flowing fast or falling, where there are 

no eddies, and where the depth is at least 15 cm (6 in). Ideally, the sampling point is one that 

can be reached during most flow conditions while kneeling on the stream bank or on stable 

rocks downstream from the sampling point. Where possible, sites should be selected that 

allow the sampler to avoid standing or stepping in the water to reach the sampling point and 

to avoid any disturbance of the streambed upstream from the sampling location. Ideally, sites 

should be selected that allow the sampler to reach upstream to collect the sample; well 

upstream of his or her immediate location and well upstream of any location that has been 

disturbed. 

• Stream-sample sites should be identifiable by reference to semi-permanent landmarks such as 

confluence points of major tributaries, well-marked boundary lines, and stream crossings by 

permanent roads or well-marked trails if they occur in proximity to the selected site. 

• Stream sample sites should be selected to avoid direct runoff from roads and trails, as well as 

unmixed flow from tributaries, unless the goals of the sampling include those conditions. 

This will be achieved for most small streams by selecting sampling sites at least 50 m above 

road or trail crossings or 50 m above or below inflowing tributaries.

ESTABLISHING AND LOCATING SAMPLING SITES

For sites that are or will be subject to periodic or routine monitoring, a site information folder or 

report should be established for each site, and provided to the field crew in advance of each 

sample collection visit. The site information folder should contain: 

• Driving and site access directions;

• Maps, including U.S. Geological Survey (USGS) 1:24,000 quadrangle maps and site maps;

• Estimated travel time from the base location to the sampling site;

• Overnight lodging and/or camping information;

• Local contact personnel, if applicable;

• Data collection forms;

• Permission letters for access, if needed;

• Site coordinates and elevation;

• Site-tag numbers for long-term monitoring sites that are marked with a tree tag and locations 

(not allowed in wilderness);

• Site photographs; and

• Other relevant information.

Maps provided in the site information folder may also include forest recreation maps to help 

navigate to the area. Maps generated using geographic information systems (GIS) could also be 

included to show where the project manager has selected potential sites to sample, spatial patterns 

in the distribution of vegetation types or other landscape properties (e.g., soil or bedrock 

distribution), or locations where sites were sampled in a previous study.

Lake or Stream Sampling Record data sheets will serve for documenting site information, sample 

locations, and field measurements. These forms should be printed on waterproof paper. Copies of 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

24

these data sheets are provided in Appendix E and are also located on the FS website at 

<http://www.fs.fed.us/air>.

In addition to the material described above for inclusion in site documentation folders, site 

documentation materials can also include:

• Uniquely numbered aluminum tags (where allowed) for sites sampled repeatedly (i.e., 

monitoring sites) or for replacement of missing tags at previously established sites;

• Nails and a small hammer for tag placement;

• Flagging tape;

• A camera with a date/time stamp for site photographs;

• A GPS unit for determining geographic coordinates in decimal degrees; and 

• Waterproof pens for completing forms in the field.

Depending on the objectives of the field data collection, field crews may be collecting water 

samples as part of a synoptic survey, or they may be repeating sampling at the same locations in a 

monitoring effort to examine changes in water chemistry over time. The extent of documentation 

required by the field crew will depend on whether the site is new or previously established.

1.3.2 WHAT TO MEASURE

This report focuses on protocols for water-quality sampling to quantify the effects of atmospheric 

deposition on aquatic ecosystems. The atmospheric deposition pollutants addressed in this 

protocol are S, N, and, to a lesser extent, toxics. Toxics are those air pollutants that are known or 

suspected to cause cancer or other serious health effects, such as reproductive and birth defects. 

Straightforward guidance regarding sampling constituents associated with characterization or 

monitoring of the acidification and nutrient enrichment effects of S and N deposition is possible. 

However, the constituents to monitor for studies of the effects of toxics are more variable 

depending on the objectives of the particular study and are less subject to generalization. 

The primary water-quality variables to be sampled can include physical, chemical, and/or 

biological attributes. Choice of variables depends on the potential environmental risks, logistical 

issues associated with sampling, and costs. 

Water-quality survey or monitoring on National Forest System lands to evaluate sensitivity to or 

responses to atmospheric deposition can involve a number of parameters. The choice of 

parameters should clearly relate to the water-quality concerns and should be measurable in a 

routine sampling program. The challenge is to select parameters that are most important with 

respect to revealing key features of ecological integrity and that can be determined in a relatively 

straight-forward and cost-effective fashion. For some studies, samples are needed at sufficient 

frequency and temporal resolution to allow appropriate characterization or statistical trend 

detection. 

The choice of what to measure depends on the type of study: acidification, eutrophication,

bioaccumulation, and/or toxicity. Parameters to include in each of these kinds of studies are 

discussed below. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

25

ACIDIFICATION STUDIES

Atmospheric inputs of both S and N can cause acidification of soil, soil water, and fresh drainage 

water (lakes, streams). In most regions of the United States that have experienced acidification 

impacts from air pollution, the impacts have been primarily due to S deposition. There are also

some regions, especially in the Western U.S., where resources are more threatened by N than by 

S inputs. This is partially due to the very low levels of S deposition received in many Western 

locations. There are also regions (portions of the Northeast, West Virginia, high elevations in 

North Carolina and Tennessee) where both atmospheric S and N contribute substantially to the 

observed acidification in some lakes and streams. National forests occupy significant portions of 

these regions. 

Acidification from S and N deposition can have several important chemical and biological 

effects. In particular, changes in the acid-base status of surface and soil water can cause shortterm or long-term toxicity to aquatic or terrestrial biota. 

Watershed processes control the extent of ANC contribution from soils to waters as drainage 

water moves through terrestrial systems. These processes regulate the extent to which drainage 

waters will be acidified in response to acidic deposition. Of particular importance is the 

concentration of acid anions in solution, including sulfate (SO4

2-

), nitrate (NO3

-

), and organic acid 

anions. Naturally-occurring organic acid anions, produced in upper horizons of acid sensitive 

soils, usually are removed from solution as drainage water percolates into the deeper mineral soil 

horizons. In some regions, organic acids can dominate the acid-base chemistry of a lake or stream 

(as indicated by color and dissolved organic carbon concentration) due to the occurrence of 

hydrologically connected wetlands. Organic acids from wetlands, although they acidify a lake, 

also serve as buffers against further pH depression from acidic deposition. Acidic atmospheric 

deposition causes natural soil acidification, anion mobility, and cation leaching processes to occur 

at greater depths in the soil profile, allowing water that is rich in SO4

2- or NO3- to flow from 

mineral soil horizons into drainage waters. If these anions are charge-balanced by H+ and/or Ali

cations, the water will have low pH and could be toxic to aquatic biota. If they are chargebalanced by base cations, the pH of the water will be higher but the base cation reserves of the 

soil can become depleted over time.

Nitrate (and also ammonium [NH4

+

] that can be converted to NO3

- within the watershed) has the 

potential to contribute to acidification of surface waters. Additionally, N is also a limiting nutrient 

for plant and microbial growth in most terrestrial and some aquatic ecosystems. Atmospheric N 

deposition has the potential to contribute to increased productivity, eutrophication, and Nsaturation in some surface waters. This occurs in estuaries and near-coastal marine waters and in 

fresh waters in remote locations where historic atmospheric N deposition has been low. Many of 

these remote fresh waters are on National Forests. 

High concentrations of lake or stream water NO3

-

, which may indicate ecosystem N saturation, 

have been found at a variety of locations throughout the United States. Locations include the San 

Bernardino and San Gabriel Mountains within the Los Angeles Air Basin (Fenn et al. 1996), the 

Front Range of Colorado (Baron et al. 1994, Williams et al. 1996), the Allegheny Mountains of 

West Virginia (Gilliam et al. 1996), the Catskill Mountains of New York (Murdoch and Stoddard 

1992, Stoddard 1994), and the Great Smoky Mountains of Tennessee (Cook et al. 1994). 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

26

Sulfur deposition moves through watershed soils and into surface waters in anionic form as SO4

2-

.

Sulfate is the most important anion contributed by acidic deposition in most parts of the United 

States. In some regions, notably the glaciated Northeast, Upper Midwest, and West, much of the 

deposited S moves readily through soils into streams and lakes. Although SO4

2- has been 

classified as a mobile anion (Seip 1980), it is less mobile in some areas, most notably the 

unglaciated southeastern United States. Sulfate mobility is an important factor governing the 

extent to which S deposition contributes to soil and water acidification, base cation depletion, and 

Al mobilization, each of which can harm sensitive ecosystems.

Aluminum occurs naturally in soils and has a pH-dependent solubility in water. Solubility 

increases dramatically at pH values below about 5.5. One of the most important effects of acidic 

deposition on watersheds is increased mobilization of Al from soils to surface waters (Cronan and 

Schofield 1979). Aluminum concentrations in acidified drainage waters are often an order of 

magnitude higher than in more neutral waters. Effects of Al mobilization to surface and soil 

waters include toxicity to aquatic biota (Schofield and Trojnar 1980, Muniz and Leivestad 1980, 

Baker and Schofield 1982, Driscoll et al. 1980), toxicity to terrestrial vegetation (Ulrich et al. 

1980), alterations in nutrient cycling (Dickson 1978, Eriksson 1981), and pH buffering effects 

(Driscoll and Bisogni 1984). Inorganic monomeric aluminum concentrations often increase with 

decreasing pH, and reach potentially toxic concentrations (> ~2 μM) in surface drainage waters 

having pH less than about 5.5. 

SELECTION OF ANALYTES FOR ACIDIFICATION STUDIES

There can be substantial leeway in terms of selection of parameters to measure in a field study of 

surface water acidification. Analytical costs must be weighed against the value contributed by 

each constituent that can be analyzed in the laboratory. In general, this protocol recommends that 

the parameters in Table 1-7 be considered the core for inclusion in the suite of analytes to be 

measured in any study of surface water acid-base chemistry. When budgets allow, add DOC to all 

of the parameters listed in Table 1-8 of high importance. Although DOC should be measured in 

all acid-base chemistry studies, color could be substituted as an inexpensive alternative. If pH is 

below 5.5, this protocol recommends also analyzing 

for total monomeric aluminum (Alm) and non-labile 

(organic) monomeric aluminum (Alo). The 

concentration of the potentially toxic Ali is then 

obtained by subtracting Alo from Alm. Measurement 

of dissolved inorganic carbon (DIC) is optional; this 

measurement can be used in estimating bicarbonate 

(HCO3

-

) concentration, which is important as part of 

the charge balance. Some studies might choose to 

analyze silicon (Si) or total fluorine (F), but these are 

often not needed for a standard acid-base chemistry 

assessment. The concentration of Si can be useful in 

evaluating the extent of groundwater influence on 

surface water chemistry and in discriminating 

between perched and flow-through seepage lakes. It 

can also provide useful information when

Seepage Lakes

Seepage lakes are lakes that do not 

have either inlet or outlet streams. 

There are two general types: 

“perched” and “flow-through.” 

Perched seepage lakes are raised 

above the surrounding terrain, often 

by build-up of organic deposits; they 

are often precipitation-dominated in 

their hydrology. 

Flow-through seepage lakes receive 

considerable groundwater inputs and 

generally have higher ANC and Si 

concentrations than do perched 

seepage lakes. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

27

interpreting diatom data, as Si or can be limiting to diatom growth. Measurement of total 

dissolved fluorine (F) is needed to calculate the speciation of Ali into various components, such 

as Al(OH)2

+

, Al(OH)2+, AlF2

+

, Al(F)2+, Al3+, etc. This can be important because the Al-fluoride 

species are thought to be less toxic to aquatic biota than the Al-hydroxide species and free 

aluminum (Al3+).

Table 1-7. Parameters to consider for possible inclusion in surface water acidification studies.

Parameter Preferred Unit Importance Rationale

ANC μeq/L High ANC is the master acid-base chemistry variable in aquatic systems.

pH -- High Biota respond strongly to pH.

SO4

2- μeq/L High Usually the major acid anion from atmospheric deposition.

NO3

- μeq/L High Sometimes an important acid anion.

Ca2+ μeq/L High Usually the major base cation.

Mg2+ μeq/L Moderate Usually an important base cation.

K+ μeq/L Moderate Base cation, usually in low concentrations.

Na+ μeq/L Moderate Indicator of road salt contamination, geological sources, or sea salt 

inputs.

Cl- μeq/L Moderate Indicator of road salt contamination, geological sources, or sea salt 

inputs.

NH4

+ μeq/L Moderate Potential indicator of agricultural influence or anaerobic conditions.

Specific Cond. μS/cm Moderate Useful in QA evaluation of internal data consistency; potential general 

screening variable to identify low ionic strength waters.

DOC μM Variable Indicator of organic acidity.

Alm μM Variable Used with Alo to estimate potentially toxic Ali.

Alo μM Variable Used with Alm to estimate potentially toxic Ali.

Si μM Variable Potential indicator of lake hydrologic type and groundwater inputs; may 

explain some patterns in diatom presence and abundance.

DIC μM Low Used to estimate HCO3

- concentration.

Total dissolved 

F

μM Low Used for Ali speciation.

Important aspects of the water chemistry QA/QC evaluation include determining the charge 

balance and comparing measured and calculated conductivity, the sum of anions and cations, and 

comparing titrated and calculated ANC.

Charge balance calculations can also be used to determine the charge density (organic anion 

concentration per mole of DOC) of DOC in surface waters. To permit these QA/QC checks to be 

conducted, all parameters listed in Table 1-8 are required except Si. Thus, the full list of

parameters should be analyzed if funding permits. It is possible to perform these evaluations 

without a measurement of total dissolved F if one is willing to make certain assumptions about 

the Ali speciation.
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Table 1-8. Parameters to consider for possible inclusion in studies of atmospheric nutrient 

nitrogen enrichment of fresh waters.

Parameter Preferred

Unit

Importance Rationale

Total N μM High Reflects all forms of N in the system.

NO3

- μM High Biologically available form of N.

NH4

+ μM High Biologically available form of N.

Dissolved organic N (DON) μM Moderate Potentially available form of N.

Total P μM High Reflects all forms of P in the system.

Soluble reactive P (SRP) μM High Biologically available P.

Chlorophyll a μg/L High Reflects primary productivity.

Fe μM Variable May bind with P, influencing its bioavailability and transport.

Total Al μM Variable May bind with P, influencing its bioavailability and transport.

Ca2+ μM Variable May bind with P, influencing its bioavailability and transport.

Si μM Variable May be limiting to diatoms under some conditions.

Dissolved oxygen (DO) mg/L Variable May decrease to biologically stressful levels under extreme 

conditions of nutrient inputs (under most conditions of 

atmospheric nutrient deposition, decreased DO is not an 

important issue).

Total suspended solids (TSS) mg/L Variable May be an erosional source of P to streams.

Turbidity standard units Variable May be used to estimate TSS.

Secchi depth m Variable Can reflect algal abundance in lakes.

SELECTION OF ANALYTES FOR EUTROPHICATION STUDIES

Eutrophication, or nutrient enrichment, is a potential consequence of N deposition to aquatic 

ecosystems. Many freshwater ecosystems are P-limited and would not be expected to increase 

primary productivity in response to increased atmospheric inputs of N. However, there are also 

many examples of fresh waters that appear to be N-limited or N and P co-limited (e.g., Baron 

2006, Elser et al. 2009). In such aquatic systems, atmospheric inputs of N would be expected to 

increase productivity and/or alter biological communities such as phytoplankton. 

Atmospheric deposition of N is expected to increase in the future in remote areas that are situated 

down-wind from centers of agricultural and/or human population growth. Surface waters in such 

areas may be N-limited. As a consequence, N additions can contribute to nutrient enrichment 

(eutrophication), including changes in algal species distribution and abundance. In particular, 

high-elevation areas in the Sierra Nevada and Rocky Mountains, and perhaps portions of the 

Cascade Mountains, are susceptible to such increases in nutrient N deposition (Fenn et al. 2003, 

Sickman et al. 2003b). In some areas, atmospheric N deposition has been linked with 

eutrophication of high-elevation lakes (cf., Sickman et al. 2003a, Melack et al. 1989). 

Estuaries and other coastal ecosystems are also susceptible to nutrient enrichment, especially 

from N. Estuarine and marine waters tend to be N-limited. Land clearing, agricultural land uses, 

sewage treatment discharge, and atmospheric deposition can all result in high loadings of N to 

coastal zones. Excessive N inputs can contribute to a range of impacts, including enhanced algal 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

29

blooms, decreased distribution of seagrasses, and decreased dissolved oxygen (DO) concentration 

(Valiela et al. 1992, Nixon 1995, Borum 1996, Bricker et al. 1999, Kopp and Neckles 2004). 

Because of human population growth and the great popularity of coastal areas, there is substantial 

potential for increased N loading to coastal ecosystems. Atmospheric deposition of N contributes 

to that load, but is generally not the major source of estuarine N. Air Quality Related Values for 

protection of estuarine ecological conditions are beyond the scope of this protocol and are not 

addressed. 

In a study of potential eutrophication of lake or stream water, a variety of measurements may be 

useful (Table 1-8). In general, measures of N, P, and chlorophyll a are of greatest importance. 

This protocol recommends, at a minimum, that water samples be analyzed for total N, NO3

-

, 

NH4

+

, total P, soluble reactive phosphorus (SRP), and chlorophyll a. Dissolved organic N (DON) 

may also be of interest. Measurement of SRP is intended to reflect the forms of P in surface 

waters that are most readily available to aquatic biota. Nevertheless, P forms are interchangeable 

within the water column and stream/lake sediment. Therefore, measured total P, which includes 

both soluble and particulate forms, is also of interest in evaluating potential nutrient limitation 

and growth responses. NH4

+ and NO3- are primarily biologically available forms of N. 

Nevertheless, DON may be converted to NH4

+ and NO3- or used directly by some producers and 

thus, measured total N is also of interest. 

Additional physicochemical parameters that can be useful in evaluation of nutrient status include 

iron (Fe) (and perhaps other metals), Si (lakes only), DO, total suspended solids (TSS), turbidity, 

calcium (Ca2+), total Al, and Secchi depth (a physical, rather than a chemical, measurement). 

Iron, Ca2+, and Al can bind to P and influence its cycling between sediment and water and also its 

bioavailability. Silica can be limiting or co-limiting, along with P and N, to diatom productivity. 

High productivity in response to nutrient enrichment can lead to reduction in DO. This effect is 

generally associated with rather extreme eutrophication, well above the levels that might be 

expected to occur in response to atmospheric deposition inputs to fresh waters on National Forest 

System lands. Total suspended solids concentration is useful because eroded sediments, 

especially the smaller clay-sized particles, can be relatively enriched with adsorbed P, depending 

on local geology and land use. Thus, eroded sediments contribute to the total P in surface waters, 

especially in streams during high-flow periods. 

It is therefore helpful to evaluate human activities throughout the entire watershed, especially 

land-use actions that contribute substantial erosion to surface waters. Turbidity can sometimes be 

used, along with the appropriate training set that includes simultaneous measures of TSS and 

turbidity, to estimate TSS in stream water. It is possible to rely upon routine measurement of 

turbidity and infrequent measurement of TSS (a more expensive laboratory analysis), along with 

an empirical relationship between the two to estimate TSS. Depending on the stream, this 

approach can yield relationships that are more or less robust. Secchi depth provides an indication 

of relative algal density in lake water, which is one biological response variable.

BIOACCUMULATION AND TOXICITY STUDIES

Atmospheric deposition can contribute to toxicity responses in several ways. The atmospheric 

pollutants of concern, in addition to H+ and Ali associated with acidification toxicity, are 

primarily Hg, pesticides, and trace metals. 
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Mercury

Atmospheric deposition is an important component of Hg cycling and biogeochemistry. Mercury 

is naturally occurring and is found throughout the environment. Mercury is also present in fossil 

fuels, is released into the atmosphere during combustion, and is subsequently deposited to the 

Earth’s surface. Mercury enters lakes and rivers from atmospheric deposition emitted by air 

pollution sources and also from nonpoint sources via erosion and runoff. 

Mercury is known to bioaccumulate in aquatic organisms, reaching potentially high 

concentrations in larger, piscivorous fish and the species that consume them such as loons and 

river otters. Mercury causes physiological and central nervous system effects on piscivorous 

birds, piscivorous mammals, and humans who consume large quantities of fish or seafood that 

have accumulated high levels of Hg. Fish consumption advisories for various lakes and rivers 

have been issued in most States throughout the United States. 

Monitoring studies to evaluate the extent to which Hg deposition affects aquatic ecosystems 

could focus on the concentrations of total and/or methyl Hg in water, or in tissue of invertebrates, 

fish, or piscivorous birds and mammals. Alternatively, methylation rates within different 

environmental compartments might be quantified. 

This protocol does not recommend that the FS ARM program initiate widespread efforts to 

measure and/or monitor Hg concentrations in surface waters. Rather, focused studies are 

recommended in areas where atmospheric deposition of Hg is known or suspected to be high. 

Such studies should begin by investigating Hg concentrations in muscle tissue of large 

piscivorous fish as these data would likely be more useful than measurement of Hg 

concentrations in water for preliminary judgments regarding Hg cycling and toxicity issues. 

Furthermore, measurement of ambient Hg concentrations in water is technically difficult and 

requires advanced training of field crews. 

Pesticides and Other Toxics

Pesticides and other toxics can also be deposited from the atmosphere and some of these can 

bioaccumulate in predator species. The degree of bioaccumulation is generally a function of the 

age of the organism and its position in the food web. In general, older individuals at the top of the 

food web have bioaccumulated more toxic materials than have younger individuals nearer to the 

bottom of the food web.

Pesticides applied to agricultural crops can become volatized or suspended in the atmosphere 

with dust particles, and eventually be transported by wind to remote areas. Organophosphates 

have been detected in precipitation at elevations up to 1,920 m in Sequoia National Park (Zabik 

and Seiber 1993) and measured in plant foliage across a range of elevations (Aston and Seiber 

1997). Although the effects of atmospheric deposition of pesticides at remote locations are poorly 

known, there is particular concern that fungicide deposition could harm sensitive lichen species 

(McCune et al. 2006). 

A variety of other toxic chemicals can be atmospherically deposited, some of which have the 

potential to bioaccumulate. These include trace metals, polychlorinated biphenyls (PCBs), and 

some fire retardant chemicals.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

31

1.3.3 WHEN TO SAMPLE

There is no one-size-fits-all answer to the question of when to collect samples of surface water for 

evaluating potential effects of atmospheric deposition. The answer depends on the type of study 

and its specific objectives. A suggested breakdown of sample timing is given in Table 1-9. It is 

fairly standard procedure to target sampling to a particular season under base flow conditions. 

Lakes are often sampled during summer or fall. Streams are often sampled during spring base 

flow for acidic deposition research and summer for nutrient work. Streams in the southeastern 

United States, where snowmelt is not a major hydrologic factor, are often sampled during spring. 

Because of the possibility for either episodic or incremental degradation of water quality, it may 

be important to implement a program for monitoring both short- and long-term changes.

Table 1-9. Suggested timing of surface water samples for evaluation of sensitivity to and effects 

from atmospheric deposition.

Type of Study Suggested Timing of Sample Collection

Lake or stream characterization Index period – at least one sample each year for at least 3 years, and

High-flow period (snowmelt and/or rainstorm) - At least two samples, if 

possible.

Synoptic survey Index period – At least one sample at each site within a relatively narrow 

time period; avoid high-flow conditions.

Characterization of episodic chemistry during 

hydrologic events

High-flow period – At least three samples during each of at least three 

hydrologic events, including at least one large storm (1-year storm or 

larger) and (if applicable) one substantial snowmelt event.

Long-term monitoring Acceptable Approach 

Index period – At least one sample per year within a relatively narrow time 

period or hydroperiod; avoid high-flow conditions.

Preferred Approach

Index period – At least one sample per season during the open-water 

seasons, within relatively narrow time periods or hydroperiods.

Modeling with MAGIC or PnET-BGC model Acceptable Approach

Index period – At least one sample.

Preferred Approach 

Index period – At least one sample during each of the open-water seasons 

during at least 3 years.

For some studies, it may be desirable to avoid sampling during abnormally low or high runoff 

conditions. Other studies may be focused on extreme flow conditions. USGS discharge data (for 

example, North Carolina data are found at: <http://nc.water.usgs.gov/info/h2o.html>) can be 

examined before going into the field to evaluate current stream flow from stream gages in the 

general area of the sample site. This precaution is more important when sampling stream

chemistry (as opposed to lake chemistry) and when water characterization will be based on a 

single sample rather than multiple samples collected at different times. It is also important to 

consider the potential influence of climatological wet/drought cycles on the chemistry of surface 

waters.
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LAKE OR STREAM CHARACTERIZATION

The water chemistry of lakes or streams is often characterized on the basis of a single sample, 

collected during the index season (spring or summer for streams; summer or fall for lakes). 

However, it is preferable to base surface-water characterization, assessment, or modeling on 

multiple samples collected over several years either throughout the annual cycle or during the 

index season.

SYNOPTIC SURVEY

The time at which the water sample is collected during a synoptic survey can influence the 

resulting chemistry and the ways in which the data can be used. Stream-water chemistry, and to a 

lesser extent lake-water chemistry, can vary diurnally (Burns 1998), seasonally (Lawrence et al. 

2004), and annually (Murdoch and Shanley 2006). Stream-water chemistry can also vary hourly 

in response to changes in flow (Lawrence 2002). If the objective is to conduct a synoptic survey 

to compare measurements among different streams or lakes within the same region (a spatial 

assessment), all sites should be sampled in as short a time period as possible. Even this approach 

might not ensure the same sampling conditions if a localized storm was affecting only part of the 

study region. Nevertheless, approaches can be adopted to minimize the effects of temporal 

variability when conducting synoptic sampling.

The major causes of temporal variability are generally flow and weather. Diel changes in pH and 

metal concentrations due to patterns of photosynthesis and respiration can also be important. 

Climate is particularly important in regions where snow accumulation occurs in winter, and water 

chemistry is affected by snowmelt (Campbell et al. 1995, Lawrence et al. 2004). Seasonal effects 

can be addressed by restricting the sampling to a period that falls within a single season. 

However, the choice of season can also affect the frequency and magnitude of flow variations. 

For example, in some regions, summer is the season that typically has the lowest flows and the 

fewest rain- or snowmelt-driven hydrological events. Scheduling a synoptic survey during a 

period of stable flows in such regions is therefore more easily accomplished during the summer 

as compared with other seasons. 

The choice of sampling season may necessitate collection of samples when flow variations are 

relatively frequent and substantial. Extreme chemical conditions are often associated with 

extreme flows. This is the case with some acidic deposition effects, which tend to be most severe 

in some regions during the high, fluctuating flows of spring snowmelt or the large rainstorms 

associated with hurricanes or other major storm systems. Although these are the conditions most 

difficult to characterize, they are often highly relevant for assessing biological impacts. 

Two approaches can be used in synoptic studies to address flow variations within a season, but 

each requires stream-flow gages within the sampling region to monitor flow conditions during the 

sampling period. In the first, sampling of each stream is repeated multiple times during the season 

under a variety of flow conditions. If the mean and distribution of flows at the collection times are 

not statistically different among the streams, the mean or median chemical concentration at each 

site can be used as the representative value for comparing streams. This approach was used 

successfully in the first large-scale stream survey to assess acidic deposition effects in the United 

States (Colquhoun et al. 1984).
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

33

The second approach involves collection of samples at all sites during a period of time that has 

limited variation in flow. This approach requires close monitoring of weather conditions coupled 

with the ability to initiate or interrupt the sampling on short notice. This approach was 

successfully used in two snowmelt surveys by Lawrence et al. (2008b) in the Adirondack region 

of New York. Collection of all stream samples was done over 3 days when flows were elevated 

but stable (higher than 90% of the year in one survey and higher than 84% of the year in a second 

survey). This approach can be challenging to implement with stream sampling sites that are 

difficult to access or far apart.

Neither of these approaches is likely to collect samples during the most extreme conditions. 

However, the stream gage(s) flows can be used to determine how the flow conditions during the 

sampling window relate to conditions for the overall year and previous years. If information is 

available to show that the chemical measurement of interest is statistically related to flow, then it 

is possible to estimate the measurement for more extreme flow conditions based on this 

relationship and available flow data. For streams where flow measurements are not available, 

chemical concentrations for flows higher than those at the time of sampling can be approximated 

from an index stream where flow and chemical concentrations are monitored on a regular basis 

throughout the year. Examples of this approach are given in Eshleman (1988) and Lawrence et al. 

(2008b).

CHARACTERIZATION OF EPISODIC CHEMISTRY DURING HYDROLOGIC EVENTS

Characterization of episodic chemistry (generally of streams, occasionally also of lakes) during 

hydrologic events is challenging under the best of circumstances. Given the additional 

complications of access difficulties for remote sites, episodic sampling in backcountry settings is 

seldom attempted. Nevertheless, such sampling can yield important information to aid in 

interpretation of surface water acid-base and nutrient chemistry. 

Because of the transient and unpredictable nature of hydrologic events, precise timing of samplecollection occasions is generally not possible. In particular, it cannot be assumed that such 

samples necessarily capture the most extreme chemical conditions. For that reason, multiple highflow samples should be collected, if possible, and they should be distributed across multiple 

years. 

LONG-TERM MONITORING

If a long-term monitoring program for lakes is intended to represent a particular time of year, 

inter-annual variation in lake hydroperiod (periodicity in lake conditions that reflect the changes 

of the seasons, including water and air temperature, snowmelt, and vegetative development) can 

introduce substantial variability. This can be especially problematic for high-elevation lakes. The 

chemistry of such lakes can change, sometimes abruptly in response to spring snowmelt or fall 

overturn. Such changes are largely governed by the depth of the snowpack, patterns of rainfall, 

and temperature. Thus, a program that entails, for example, sampling the first week of July each 

year, while reducing some aspects of inter-annual variability, may still yield considerable year-toyear variability as a consequence of inter-annual differences in snowmelt hydrology. One 

potential solution is to target sampling to a specific degree day, which is calculated based on 

maximum and minimum daily temperature (cf. <http://pnwpest.org/wy/index.html>). 

Standardization of sampling timing on the basis of degree day can partially adjust for inter-annual 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

34

differences in snowmelt and the transition to summer weather. It should therefore eliminate some, 

but not all, of the variability associated with standardization based on the calendar for certain 

types of lakes. Such an approach should not, however, ignore the influence of flow conditions. 

One can quantify changes over time in the concentration-discharge relationship. Using this 

approach, one compares differences between the concentration-discharge relationship determined 

during one period of time (several months to several years) and the same relationship determined 

during a later period of time of similar length. If, for example, the concentration of stream ANC is 

higher at a given flow condition this year than it was several years ago, the change may suggest 

that ANC may be increasing at times represented by that flow regime, independent of any 

changes in flow. 

Analysis of trends is often done on an annual basis using one of several approaches that 

incorporate seasonal effects (Helsel and Hirsch 1992; Lawrence et al. 2004). These approaches 

are most effective if multiple samples are collected for each season. Weekly sampling provides a 

sufficient number of samples to account for within-season variability and is likely to enable a 

trend to be detected sooner than data collected at longer intervals. Annual or quarterly sampling is 

less expensive than weekly sampling, but does not account for within-season variability and can 

substantially increase the length of time needed to detect a trend (Murdoch and Shanley 2006). 

However, annual or quarterly sampling may free-up resources to monitor more sites to get a 

better picture of regional patterns. Thus, the intended eventual use of the data is important for 

making sampling decisions, as is the length of time one is willing to wait before being able to 

document with statistical certainty that a change has taken place. 

Because long-term monitoring of surface-water chemistry is usually based on sampling at a 

constant frequency, most samples are typically not collected during high-flow periods. However, 

long-term trends in stream chemistry may first become apparent during high flows. An approach 

for separate trend analysis of high, medium, and low flows has recently been developed 

(Murdoch and Shanley 2006). This method uses annual or grouped years of data to develop 

concentration-discharge relations that enable concentrations to be predicted for various flow 

conditions throughout the year. An annual value can then be derived for upper, medium, or low 

flow ranges so that long-term trends can be determined for each specific flow range. This type of 

approach requires that: 1) flow is monitored for the stream site of interest; 2) the solute of interest 

is statistically related to flow; and 3) sufficient data are available to develop the concentrationdischarge relations.

1.4 FIELD METHODS

The methods outlined here are appropriate for analysis of low ionic-strength stream and lake 

waters associated with forested and alpine watersheds in lands that are sensitive to acidification, 

toxicity, and/or nutrient enrichment impacts from atmospheric deposition. Because stream and 

lake waters in these sensitive areas can be extremely dilute and easily contaminated, care must be 

taken in all phases of sample collection and analysis to ensure that data will be of sufficient 

quality to support the intended assessment purposes. Each of the important aspects of field 

sampling is discussed here, with an explanation of the reasons why certain steps should be taken 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

35

or avoided in the sampling program. The intent is to provide a general understanding of sampling 

issues. The specific, step-by-step instructions to the field personnel should be adapted by the 

project manager from the SOPs included in the appendices for the relevant protocol. The protocol 

provides core attributes and standard sampling design and procedures across the country, which 

should then be used by local FS personnel to develop a site-specific project plan that includes the 

SOP with the specific steps to be followed by field staff. The project plan and SOPs may vary 

from study to study, but the overall protocol remains constant. General SOPs are provided in 

Appendix D, which should be modified as needed. 

1.4.1 PRE-TRIP PREPARATIONS

A field data sheet should be prepared in advance for each sampling site. The lake and stream 

sampling record data forms are provided in Appendix E. The appropriate data forms should be 

completed to the extent possible before the sampling trip. For previously established sites, the 

available site information, site tag number, and description of the tag tree (where applicable) 

should be filled out on each form.

Pre-trip preparation should include an evaluation of sample-holding-time issues. Determine in 

advance what parameters will be analyzed in the laboratory, and then check the laboratory 

protocol to determine the holding-time requirements of these measurements. The requirements

may influence sample-collection scheduling and/or in-field sample aliquot preservation decisions. 

Coordinate with the laboratory regarding the sample delivery timing. In general, sampling should 

not be done late in the week or in advance of a holiday unless arrangements have been made with 

the laboratory to receive samples on weekends or holidays. 

The most important issues to consider before going to the field can vary depending on the type of 

study. A checklist of important issues to consider in advance of initiating and implementing an 

inventory or monitoring field effort is provided in Table 1-10. Additional issues to consider 

before initiating a long-term monitoring effort are listed in Table 1-11. Table 1-12 provides a list 

of issues to consider when conducting a study that will involve critical load or emissions scenario 

modeling.
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Table 1-10. Key issues to consider when initiating and implementing an inventory of lake or stream 

water chemistry to assess the effects of atmospheric deposition.

Issue Question

Site-Selection Is the lake/stream representative of other lakes/streams in the wider regional or National Forest System land 

population? How sensitive to the stressor(s) of interest is it expected to be?

Is the sampling site and its upstream drainage basin reasonably free of disturbances other than atmospheric 

deposition (i.e., acid mine drainage; geological S; fertilizer application; heavy livestock influence; riparian, inchannel, or shoreline disturbance)?

Is the sample site representative of the body of water being sampled? In other words, is the lake sampling 

site in the deepest part of the lake or in the outlet stream? Is the stream sampling site in the thalweg well 

below the nearest up-stream confluence?

Implementation Have arrangements been made with the laboratory? Are any required permits or property access 

permissions obtained?

Have issues associated with laboratory holding times and length of time needed for -site access and delivery 

of samples to the laboratory been addressed?

Have clean, appropriately sized bottles and (if required) syringes been obtained?

Have decisions been made about field QA/QC activities including use of sample replication and field blanks?

Have all required sampling equipment and supplies been assembled and checked?

Have all safety procedures been reviewed?

Is the timing of sample collection standardized and appropriate to the research questions? For example, is 

sampling focused on a summer or fall index period for lakes? Is sampling linked to seasonal climatic shifts? 

How many samples are collected from each site each year, and how are they distributed in time?

Table 1-11. Key issues to consider when initiating and implementing a long-term monitoring effort 

to document and quantify trends in lake or stream chemistry over time in response to 

inputs of atmospheric deposition. Issues include all of those listed in Table 1-10, plus 

the following.

Issue Question

Expertise Has a statistician or someone knowledgeable about statistics been consulted in planning the monitoring effort?

Temporal 

variability

Has temporal variability in the lake or stream been characterized before including that body of water in the 

monitoring program? This might include, for example, collection of weekly or seasonal samples within the index 

period during 1 or 2 years.

Has an analysis been conducted to determine, given the amount of temporal variability documented in this 

water body, how large a change over what period of time would allow unambiguous, statistically significant 

demonstration of change over time in key water chemistry parameter(s)?

Table 1-12. Key issues to consider when conducting modeling using the MAGIC model (Cosby et 

al. 1985) to estimate critical load or to calculate changes in lake/stream chemistry in 

response to future emissions controls. Issues include all those listed in Table 1-10, 

plus the following.

Issue Question

Soils Are soils data available for the subject watershed? Although soils protocols are beyond the scope of this 

document, MAGIC requires soil chemistry data from the upper mineral B soil horizon (often the top 20 cm of 

the B horizon). Two to three soil pits per watershed are generally recommended. Soil parameters needed for 

MAGIC include pH; cation exchange capacity (CEC); exchangeable Ca, Mg, K, Na, and Al; exchangeable 

acidity; bulk density; loss on ignition; and an estimate of soil depth. 

Discharge Is annual discharge available from a stream gage at or near the location of sample collection? If not, can 

discharge or runoff be estimated from regional data?

Deposition Are estimates available for total (wet plus dry plus occult) deposition of S, oxidized N, and reduced N?
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

37

Table 1–12. Continued.

Issue Question

Models If models other than MAGIC are to be used, have the required inputs been determined and are appropriate 

input data available?

If critical and/or target loads are to be modeled, have decisions been made regarding the resource(s) to be 

protected, chemical indicators of biological effects, critical levels of those chemical indicators, and time period 

at which protection is to be evaluated?

PERMITS AND ACCESS

Sampling in designated wilderness areas may require obtaining special permission or permits. 

Typically, if the monitoring plan involves installation of equipment or motorized access, special 

authorization will have to be obtained. It may not be possible to obtain permission to install 

equipment, such as a stream gage, in the most highly protected areas. Early coordination with 

forest managers and staff regarding all monitoring programs is recommended.

Sampling on lands outside National Forest boundaries or obtaining access across other lands may 

require additional coordination and planning. Make sure you plan well ahead to obtain any 

permits or permissions necessary to conduct the project before finalizing your plans. Consider 

selecting back-up sample locations to be used only if access is not available to the intended 

sampling site(s). Such an approach may be useful if, for example, road or trail access is blocked 

by late snowpack, road washout, avalanche, land slide, or other impediment to access of remote 

sites.

LABORATORY AND SAMPLE BOTTLE ARRANGEMENTS

Appropriate agreements should be made or contracts established with a qualified water chemistry 

laboratory well in advance of field sampling. If waters are expected to be dilute, the laboratory 

must be able to implement low-ionic strength methods to achieve the necessary data quality 

objectives. The laboratory, or another entity, should prepare and provide sample bottles, insulated 

shipping containers, and refrigerant. Historically, the FS Air Resource Management Laboratory 

(ARML) in Fort Collins, Colorado, has performed many of the analyses on water chemistry 

samples collected on National Forest System lands to evaluate sensitivity to and effects from 

atmospheric deposition. 

This protocol recommends that plastic ware and plastic aliquot bottles should be high density 

polyethylene (HDPE), low density polyethylene (LDPE), or polypropylene. The sample bottle 

must be made of a material that is nonreactive with the chemical constituents to be measured. 

Polyethylene and polypropylene are commonly assumed to be essentially inert with respect to 

most dissolved substances. Harder plastics, such as polycarbonate, tend to be less reactive but 

will crack more easily than softer plastics. Glass is generally preferred for measurement of low 

concentrations of dissolved carbon. Teflon®, the most inert plastic but also the most expensive, 

can be used for measurements of trace concentrations of highly reactive substances.

New bottles should be soaked in deionized water (DIW) before use. Samples can also be 

collected into previously used bottles that have been rinsed with a dilute wash acid (e.g., 

hydrogen chloride [HCl] 2%) and soaked in DIW for at least 24 hr. The laboratory should follow 

a procedure to check acid-washed bottles to ensure that all traces of the acid are undetectable in a 
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38

chemical analysis. The ARML does not acid wash bottles. Instead, bottles are washed and 

subsequently checked with DIW blanks to ensure that cleaning is adequate. 

Generally, the laboratory is responsible for providing contamination-free bottles for the sampling. 

Sample bottle preparation should involve a triple-rinsing of each bottle with DIW. The bottles 

should then be stored overnight or longer filled with DIW, followed by another rinse with DIW. 

Ideally, each bottle should then be filled with DIW (which can be poured out after 24 hours or at 

the time of sample collection). Treating the sample bottles in this manner will help ensure a 

contamination-free sample. Laboratory conductivity analyses of blank samples typically employ a 

standard acceptance criterion of less than about 1.2 to 2 μS/cm.

Note that, if water samples are to be collected for analysis of Hg concentrations, sample bottles 

will need to be Teflon or glass (with Teflon-lined caps), and special bottle cleaning procedures 

will need to be followed, including prolonged heating in an acid solution. Check with the 

analytical laboratory for specific requirements. 

The following steps need to be completed before going to the field:

• Obtain the necessary sample bottles and (if required) syringes for each site to be sampled. 

Depending on the intended laboratory analyses and sample replication requirements, this can 

range from one to several bottles and syringes per site. Preprocessed bottles, often filled with 

DIW, with a preprinted or blank label tape affixed, should be provided by the analytical 

laboratory. It is a good idea to carry a few extra bottles beyond those needed for the intended 

sampling. 

• Field studies often, but not always, include some sample replication (often 5 to 10% of 

samples) for quality assurance purposes. Replication is generally desired to assess the 

Sample Bottles and Labels

The size of sample collection bottles can vary depending on the parameters to be analyzed, 

but should usually be 500 ml or 1000 ml (large enough to allow reanalysis, if necessary), 

wide-mouth HDPE or LDPE bottles. For some studies, it may be possible to use smaller 

bottles; some studies collect 2 L of water, but a full bottle weighs over 4 pounds (2 kg). 

Pre-printed labels with prompts for all required information associated with the sample should 

be affixed to each sample bottle as part of the bottle preparation (see Appendix F). At a 

minimum, this information should include: 

1. collection date and time; 

2. lake or stream name and site location (inlet/outlet/deep); 

3. name of the person who collected the sample (first initial and complete last name); 

4. Sample ID/Barcode; and 

5. whether the sample is a regular, replicate or blank sample. 

The field crew must take precautions to ensure that no bottle mouth is contaminated with 

leaking refrigerant, tap water, dirt, handling contact or other foreign substance. Package each 

processed bottle individually in a zipper-lock type plastic bag. Refrigerant should be in double 

zipper-lock bags. This precaution is especially important if the laboratory analysis indicates 

that sample contamination has been an issue in the past.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

39

repeatability of the sampling procedure, sample holding and treatment, and laboratory 

analysis. The amount of replication will be dependent on the sampling design and the QA/QC 

program. Some studies, including some ARM programs, replicate all samples, but may only 

pay for analysis of a portion of these, keeping the others as “insurance” against 

contamination, leakage, or shipping problems. This should be done in areas where access to 

the site is difficult and travel to the site is the biggest expense involved in the sample 

collection and analysis.

• Obtain ice blocks and/or frozen refrigerant. Ice blocks generally work better for shipping 

samples to the lab unless a large number of refreezable packs are used. If using refrigerant, be 

sure that it has been in the freezer at least two days before the day of sampling. If using block 

ice, it must be placed in two securely sealed plastic bags to prevent leakage. The outer bag 

should be clearly marked “ice”. If using refrigerant, place each refrigerant container in two 

securely sealed zipper-type plastic bags to prevent sample bottle contamination in the event 

of leakage. Place the refrigerant containers into insulated container(s) that will be used for 

sample holding and transport. Provide enough refrigerant to keep the samples cold until 

delivery to the lab or until placement in a refrigerator if samples are to be stored at a staging 

area before shipping. 

• Transport the sample bottles and syringes, including the replicate bottles and process blank 

bottles (if applicable), in the cooler that will be used to store the samples. Bottles can be 

transferred to a small cooler, suitable for carrying in a backpack, before departing from the 

trailhead to access a site. 

Note that, if sample filtering is to be performed in the field (not recommended in this protocol), 

then field blanks should be transported to the field, filtered in the field, and returned to the 

laboratory for analysis due to the increased potential of contamination with field filtering.

ACQUISITION OF EQUIPMENT, SUPPLIES, AND DATA FORMS

Each person or sample-collection team should typically be provided the following equipment and 

materials:

• Site information folders (including maps and stream water field data forms);

• Site documentation materials;

• Sampling protocol (this document);

• SOPs document(s) for sampling and sample handling;

• Sampling bottles (with label affixed, in a zipper-lock plastic bag) and syringes, if applicable

(in a light-weight plastic box with snap-on lid, large enough to hold multiple syringes with 

plunger pulled three-fourths of the way out);

• Plastic gloves, stored in a secure plastic bag;

• Insulated containers, refrigerant, and backpacks;

• Thermometer appropriate for use in air or water;

• Wrist watch;

• Survey-grade global positioning system and compass;
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40

• 50- or 100-meter tape (and, if available, laser rangefinder [optional]) to measure distances;

• Waterproof labels and markers;

• Notebooks and number 2 pencils or write-in-rain-type pens;

• Digital camera and extra memory cards and battery;

• Heavy-duty aluminum tags, aluminum nails, and a hammer if the sites are being established 

for long-term monitoring of water chemistry and this type of marking is permitted;

• Backpack with waterproof cover (if the site is not accessible by vehicle);

• Van Dorn sampler if sampling a lake;

• Cable and instrumentation for lake “at-depth” measurements;

• Raft or float tube for in-lake sampling;

• First aid kit; and

• Locally appropriate safety equipment.

Depending on the study, other materials may also be required. 

Sufficient time should be allocated in advance of field work for assembling and checking all 

equipment needed for the sampling program and to make sure that field personnel are thoroughly 

familiar with all field equipment. Arrangements will need to be made in advance for a vehicle 

that is suitable for carrying capacity needs (people and equipment) and anticipated road 

conditions. 

Supplies must be assembled for sample collection and transport. These include sample bottles 

(usually provided by the laboratory), sample syringes (if applicable), refrigerant, coolers or other 

sample containers for transport of samples from the field to the vehicle, coolers for transport of 

samples from the vehicle to field staging areas, and packaging materials (including refrigerant) 

for shipping samples to the laboratory. 

PLAN FOR STAFFING

Field sampling staffing needs should be determined well in advance of sampling activities, 

allowing an adequate time buffer for possible extension of the sampling effort in the event of 

inclement weather or unforeseen circumstances. Field efforts frequently require more time than is 

initially estimated. In addition, it can be advantageous to identify at least one back-up field person 

in the event of sickness or injury. 

Field staff should be current on first aid training and local emergency procedures before heading 

to the field. Lead time of several months may be required for staff to obtain proper first aid 

training.

1.4.2 SAMPLE COLLECTION

Sample bottles should be labeled using label tape and indelible ink (Appendix F). Information on 

the bottle label should also be recorded on a chain-of-custody record (Appendix E), along with 

information about the desired analyses and the identity (ID) of the sample collector. A field 

logbook should be kept in which station identification codes, dates and times of sampling, and all 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

41

field data are recorded. Notes on any unusual conditions at the sample sites or any circumstances 

that may have caused deviation from normal procedures should be recorded on the Lake or 

Stream Sampling Record Form and described in the field data logbook. 

Each sample should be labeled uniquely with site identifier (site name and ID number), date, and 

unique barcode or ID in the field. An additional lab number is typically added at a later time to 

each aliquot in the laboratory. The sampler should fill out the bottle label at the time of sample 

collection. The Chain of Custody Record Form should include the sample name, date, time of 

day, analyses requested, comments, and appropriate signatures. Collection time should include 

whether the time recorded was in Daylight Savings Time or Standard Time. (Most electronic data 

recording, including stream stage, is recorded in Standard Time, year round. Therefore, this 

information is needed to relate the water sample to the electronic data.) A chain-of-custody form 

is shown in Appendix E. 

This protocol recommends that aliquots of samples be collected in syringes or glass bottles with 

septum caps along with the standard bottles and that the syringe or septum-capped samples be 

used for analysis of pH and DIC. This precaution is considered more important for streams than it 

is for epilimnetic samples from lakes. If samples for these analyses are collected into bottles (with 

or without septum caps) in the field, it is especially important that no head space be left in the 

bottle (i.e., that the bottle is filled completely to the top) and that laboratory procedures limit the 

opportunity for CO2 degassing in the laboratory before and during analysis of these parameters. 

Filled syringes should be transported from the field to the lab in plastic containers that minimize 

disturbance of the seals of the syringes and protects them from breakage. 

Dissolved inorganic carbon (DIC) concentrations and pH are the measurements that are typically 

analyzed without contact with the atmosphere. Water sample aliquots for DIC and pH from 

streams should be collected in the field in sealed syringes or glass bottles with septum caps for 

some analytes to minimize contact with the atmosphere in the event that the dissolved carbon 

dioxide partial pressure is considerably higher than that of the atmosphere; in these cases, the 

concentration of some analytes can change if the water sample equilibrates with atmospheric 

carbon dioxide partial pressure before analysis. This is a common occurrence in surface waters. 

Collection into a bottle having a septum cap is done by immersing and capping the bottle 

underwater. Throughout the water chemistry sampling process, it is vital to take precautions to 

avoid contaminating the sample. Many surface waters in regions of the United States considered 

sensitive to effects of atmospheric deposition have low ionic strength (i.e., low levels of chemical 

constituents). Samples from such waters can be contaminated quite easily by perspiration from 

hands, sneezing, smoking, suntan lotion, insect repellent, fumes from gasoline engines, or 

chemicals used during sample collection.

For quality assurance, sample collection should be routinely replicated so that the variability 

introduced by the collection process can be quantified. Although duplicate collection of samples 

from a subset of the sampling sites is sometimes done, the collection of three replicate samples 

from a subset of pre-specified sites is an alternative approach for characterizing variability. The 

entire collection process should be repeated for the duplicate pairs or triplicates so that either two 

or three sample bottles representing the same sample location and taken at approximately the 

same sample time are returned to the laboratory. The frequency of replicate sampling is 

dependent on the overall structure and requirements of the quality assurance program. Some 
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42

studies replicate all samples in the field but analyze only a subset of the replicates in the 

laboratory. This protocol recommends collection and laboratory analysis of duplicate pairs for 5%

to 10% of the field samples in a given study. 

Water temperature should be measured at the location of sample collection. Place the 

thermometer in the water at the sampling point and wait for the reading to stabilize. If this is not 

practical, temperature can be measured in a sampling bottle designated for this purpose and 

labeled as such. In the latter case, cool the bottle to ambient stream temperature before filling it 

with stream or lake water for temperature measurement. An example localized SOP illustrating 

surface water chemistry sampling procedures for lake and streams can be found in Appendix C.

COLLECTION OF STREAM WATER

The stream water sample should be representative of stream water at the location of interest with 

respect to the measurement(s) of interest. Collection of water at a single point will provide a 

representative sample of the channel cross-section if the stream is uniformly mixed. The mixing 

of stream water increases with increasing flow velocity and roughness of the channel bottom. 

Streams are generally well mixed with regard to dissolved substances if flow is turbulent and 

there are no close upstream tributaries or nearby point sources of contamination. To verify that 

the stream is uniformly mixed, sampling can be done for measurement of specific conductance 

(for example, using a meter in the field) and perhaps other parameters at several points along the 

cross-section and at different depths. If the measurements do not vary beyond the expected 

analytical variability, sampling at a single point can be done thereafter. If the required sampling 

location is not well mixed along the cross-section, depth-integrated samples will have to be 

collected at multiple points along the cross-section or the sampling site might be moved to a 

different location. In general, stream sampling within the FS ARM program will involve 

sampling relatively fast-flowing small streams free of point source impacts. The water in such 

streams should generally be well-mixed. This protocol therefore recommends sampling at a single 

point in the main area of flow across the stream cross-section unless local conditions suggest the 

likelihood of incomplete mixing of water in the stream. 

A stream sampling training video has been produced by the Forest Service. Field personnel 

should review this video as part of their field training program. It is available on the Web via

<http://www.fs.fed.us/air>.

Manual Sampling

At many sites, the sample bottle may also serve as the collection device by simply dipping the 

sample bottle into the stream by hand. This avoids the need for a collecting device, thereby 

reducing equipment needs and the chance for sample contamination. At stream sites that are 

hazardous to access because of steep banks and/or high flows, a sampling pole (a long pole with a 

bottle attachment point on the end) or a weighted bottle holder can be used so that the collection 

bottle can be extended out to the stream or lowered into the stream from above. 

With the weighted bottle approach, an open sample bottle is placed within the weighted holder 

and lowered into the water with a handline. Discrete volume samplers (such as a Van Dorn 

sampler) can also be used to collect the water sample, but are usually not necessary in relatively 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

43

small streams with well-mixed flows where the objective is measurement of dissolved 

constituents in stream water. 

There are many acceptable methods of collecting water samples. Some, such as flow integrated 

stream samples, are complex and beyond the scope of this protocol; studies requiring such 

sampling should consult appropriate references (e.g., Wilde et al. 1999). Methods likely to be 

used to collect samples for most stream studies of atmospheric deposition effects are described 

below. The information in this section is taken largely from the recommendations of Turk (2001). 

Grab/Hand Samples 

Once the sampling site has been selected, bottles should be assembled and necessary information 

added to the labels to unambiguously identify the samples. If the bottles contain DIW, this should 

be discarded away from the shore so it does not disturb the sampling site.

Avoiding disturbance that can affect the water being sampled is especially important. For grab 

samples, the most likely disturbances are stirring up sediment or incorporating surface debris into 

the sample; each can significantly change analytical results. 

Personnel falling into the stream is not only a major disturbance of sediment but can also pose a 

safety risk; thus, selection of a stable place to wade or a shore location from which to reach the 

sample location is critical. Suitable sampling sites are often slippery due to water, ice, algae, mud,

or an unstable substrate, such as loose boulders or poorly stabilized logs. If the sampler tries to 

use both hands for handling bottles while leaning over the water, sudden loss of balance can 

occur. 

The sample should not be collected where the sampler has waded or fallen. If the sampler is 

holding the bottles in hand, powder free gloves can minimize contamination from sweat, etc. 

Laboratory gloves generally cover hands to the wrist, but longer gauntlet-style gloves cover to the 

elbow and should be used if the sample is collected by hand at a depth greater than a few inches. 

In addition to salts in sweat, common contaminants include sunscreen and insect repellent. All of 

these potential contaminants can be minimized by thoroughly rinsing hands and arms before 

collection and also rinsing the gloves at a site far enough away (and downgradient) such that the 

sampling site itself is not contaminated by the rinse water.

For cleaning before sample collection, bottles should be individually uncapped underwater, 

partially filled with stream water, capped and shaken, and the rinse water discarded away from 

the location where the samples are to be collected (e.g., onshore or downstream). Rinse water 

should be poured over the cap as the water is being discarded. Three rinses for each bottle are 

required unless protocols specify otherwise. The bottles are then filled completely and capped

underwater. The sampling depth should be consistent for all samples and documented. In general, 

for streams that are less than about 2 m deep, where possible, sample at a depth of about 0.3 m or 

mid-way between the water surface and the water/sediment interface, whichever is closest to the 

surface. It often is impractical and/or unsafe to collect samples deeper than about 0.3 m without a 

sampling device of some kind. 
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Shallow Samples

If the water at the site is very shallow, which may be the case for many small streams and the 

outflow of some lakes, it may not be possible to sample very far below the surface. Very shallow 

streams and seeps may require creative approaches to collecting samples without disturbing 

sediment. It may be necessary to create a small dam that allows water to drop into the bottle. The 

bottle cap, pipettes, syringes, or even plastic basters used for cooking can be cleaned and used to 

transfer samples from the stream to the bottle in extreme cases. One option, using a syringe (a 

new syringe at each sampling site), is as follows: 

• Rinse the syringe three times with stream water, downstream of the sample site, as usual.

• Use the syringe to put stream water in the sample bottle and rinse the sample bottle three 

times.

• Finally, use the syringe to fill the bottle to the brim with stream water at the sample site. Cap 

the bottle and proceed as normal.

Pole Samples

An alternative to collecting grab samples by hand is the use of a bottle attached to a pole made of 

non-contaminating material such as smooth fiberglass or painted aluminum. This approach is 

safer than leaning over the water surface or wading and often allows the sample to be collected 

farther from shore and at greater depth than can be done by hand. This approach is not suitable 

for streams with significant velocity because of excessive drag from the assembly. 

The collection bottle can be larger than the sample bottle and can therefore contain sufficient 

water to rinse and fill it. Alternatively, the sample bottle itself can be directly attached to the pole. 

Because of buoyancy and leverage, it may be impractical to use a bottle larger than about 500 ml 

capacity and a pole longer than about 3 to 4 m. The bottle can be attached to the end of the pole 

with stainless steel hose clamps or laboratory three-finger style bottle clamps. To minimize the 

possibility of contamination with surface debris or floating slush and to allow collection at a 

specific depth, the bottle can be plugged with a non-contaminating silicone stopper attached to a 

line that the sampler pulls when the bottle is at the proper depth. The depth can be estimated or 

can be measured with a simple float and line attached to the pole near the bottle. Care must be 

taken to avoid introducing soil or other debris that may have accumulated on the pole into the 

sample; the pole should be rinsed in an area away from the sampling site before use.

Deep Samples

If the stream is deeper than about 2 m, it is recommended that samples be taken about 0.5 m 

below the surface if logistics allow or that a shallower sampling site be selected a short distance 

further upstream or downstream. Sample collection at a depth of 0.5 m can sometimes be 

achieved using a pole sampler (described above) or a Van Dorn sampler (described in the section 

on the collection of lakewater samples). The choice will depend on site location and sample 

collection logistics. Use of a Van Dorn sampler requires low to moderate stream velocity and a 

stable position from which to collect the sample, such as a bridge, raft, or float tube. 

The water sample should be collected from a point where flow velocity is high relative to other 

points along the cross-section at the sampling location and water depth is sufficient to submerse a 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

45

collection device without disturbing bottom sediments. Side pools with low velocity or eddies 

should not be used for sampling. Powderless latex, polyethylene, or nitrile gloves should be worn 

while handling sampling equipment and collecting the sample to reduce the chance of 

contamination. Care must be taken to avoid touching potential contaminating surfaces while 

wearing the gloves. For personal safety reasons, field personnel should be alert to the possibility 

that some individuals are allergic to contact with these glove materials. 

A stream water sample can be collected from a deep stream at a specific point along the channel 

cross-section by 1) lowering a weighted collection bottle with a handline, 2) collecting the sample

with a discrete-volume water sampler, or 3) drawing the water sample with a suction pump 

through a tube that is lowered into the water. The sample bottle should be rinsed with stream 

water three times by partially filling the bottle, capping and shaking it, and dumping the rise 

water away from the sample site. If wading is required and if it is both practical and safe at the 

sampling location, the sampler must stand downstream of the point of collection and avoid 

collecting particulates resuspended by wading or bumping the streambed with the collector. 

Sample collection with a tube and peristaltic suction pump can be useful when large sample 

volumes are needed, but the tube and pump must be well rinsed before actual collection of the 

sample. 

Auto Sampling

If high-flow events need to be sampled in a non-wilderness setting, in most cases, auto-samplers 

should be used. However, if auto sampling is done during the winter at a location that experiences 

below-freezing temperatures, the auto-sampler should be kept in a heated shelter to prevent 

collected samples from freezing. In addition, the sampling tube should be buried between the 

auto-sampler house and the stream to prevent formation of ice plugs in the tube. 

Auto-sampling can be done with one of several types of commercially available auto-samplers. 

All operate similarly and consist of a controller, peristaltic pump, sample tubing that extends into 

the stream, and space-efficient custom-shaped sample bottles. The water sample is drawn through

a weighted suction head attached to the end of the tubing. The auto-sampler can be set to collect 

at specific time intervals. This can reduce the frequency of site visits. However, to collect samples 

timed to the hydrograph, the auto-sampler must be controlled by a programmable datalogger that 

monitors water level changes. 

Water level is most often measured by a pressure transducer installed in a deep portion of the 

stream so that it will remain below-water during low flows. A large number of pressure 

transducers are commercially available and vary in design and price. The water level measured by 

the pressure transducer is typically recorded by the datalogger at 15-minute intervals. The 

datalogger transfers this information to a data storage module for retrieval during site visits. The 

datalogger can be programmed to trigger the auto-sampler based on the rate and direction of 

change of the water level so that samples can be collected on ascending and descending limbs of 

the hydrograph as well as at the peak. Programming the datalogger for this type of sampling will 

require some knowledge of flow variability in the stream being monitored. A weatherproof box is 

required to protect the datalogger, storage module, and battery needed to run the datalogger and 

auto-sampler. Auto samplers are usually kept in a shelter to limit the risk of vandalism.
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For automatic collection of samples, the auto-sampler should be placed on the bank where there is 

no risk of its being washed into the stream during high flows. This also allows the sample tube to 

drain freely after sample collection. Auto sampling requires that the sampling tube extend into the 

stream and that the suction head at the end of the tube is anchored in a deep, well-mixed portion 

of the channel where it will not be easily dislodged by high flows. The suction head should be 

positioned so that it will not draw in sediment from the bottom. The entire section of the sampling 

tube under water also needs to be well anchored. The pressure transducer and its line to the 

datalogger should be similarly anchored on the stream bottom. 

Flow-activated auto-samplers should be visited promptly after hydrological events to retrieve the 

samples and reset the auto-sampler. Disposable, powderless latex or nitrile gloves should be worn 

while handling sampling equipment to reduce the chance of contamination. Care must be taken to 

avoid touching potential contaminating surfaces while wearing the gloves. 

Upon arrival at the site, bottles in the auto-sampler that contain samples should be capped and 

labeled with the date, site, and their position number in the auto-sampler. These bottles can then 

be removed and replaced with clean, empty bottles with caps removed. The auto-sampler is then 

prepared for sampling by resetting the counter and sampler spout to sample position number one. 

Data can then be downloaded from the datalogger. These data provide the date, time, and water 

level associated with each collected sample. This information can then be used to select samples 

for chemical analysis.

COLLECTION OF LAKE WATER

Lake water sampling should typically be done by boat, raft, or float tube over the deepest part of 

the lake. Such a sample is intended to represent average lake chemistry. If boat-based sampling is 

not possible, an alternative protocol is to sample the outlet stream, if one is present, close to the 

lake. Outlet stream chemistry should closely approximate average lake chemistry unless there are 

major perturbations in the vicinity of the outlet stream. Note that, even though this sample is 

actually collected from the outlet stream, it is intended to represent the chemistry of the lake. 

Thus, the sample should be labeled and documented as a lake sample rather than a stream sample. 

In general, this protocol recommends that lake samples should not be collected from the shore 

line for the purpose of characterizing overall lake chemistry as the chemistry of this water may be 

different from the lake average, due in part to differences in temperature and biological 

productivity. Nevertheless, if there is no outlet present or if the outlet is not flowing at the time of 

sample collection, an alternative, less desirable approach is to collect the sample from the 

shoreline. For this approach, a shoreline sample collection location should be selected that 

satisfies as many of the following criteria as possible: 

• As close to the outlet as possible;

• Near the lowest point of land around the perimeter of the lake;

• From a bedrock outcropping or otherwise rocky area; and

• From the deepest accessible point.

Water must be deep enough so that surface scum and sediments are not collected into the bottle, 

preferably in a wind-exposed area so that the water is relatively well-mixed. Avoid sampling in 

locations having emergent vegetation and/or downed logs or other woody debris. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

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Lake samples collected from the deepest lake location are normally collected using a Van Dorn 

sampler at a pre-specified depth. The Van Dorn sampler and any associated tubing should be 

rinsed three times with lake water, emptied, and then lowered to the specified sampling depth for 

sample collection. The sampler should be held at the sampling depth for approximately 1 minute 

to allow equilibration with the water at that depth. A weighted messenger is used to trigger 

closure of the sampler doors for retrieval of the sample. Water from the Van Dorn is then used to 

rinse the sample bottle and cap (and syringe[s] if applicable) three times before filling the bottle. 

Sample depth for lakes should be standardized to the extent possible across all lakes included 

within a particular program. For lakes deeper than 2 m, a sample depth of 1.0 or 1.5 m is 

commonly specified. For lakes less than 2 m deep and for lakeshore sampling (where required), 

the single-depth sample should be collected at 0.5 m depth. An alternate approach (not detailed in 

this protocol) is to collect a depth-integrated sample using a 2 m-long tube. The EPA’s National 

Lake Assessment project collected depth-integrated samples of the euphotic zone, estimated as 

two times the Secchi depth, to a maximum of 2 m. 

Normally, a surface sample (i.e., 1.5 m depth) from the deepest part of the lake, with or without 

replicate(s), is used to characterize lake chemistry at the time of sampling. For some studies, 

additional samples may be required. These might include samples of particular portions of the 

lake, littoral zone samples, or samples at different depths. Even if samples are not collected at 

different depths, lake sampling should ideally be accompanied by measurements of the 

temperature profile in order to determine if the lake is stratified and to characterize the location 

and depth of the thermocline, the epilimnion, and the hypolimnion. See the SOP for depth 

measurements in Appendix D. If samples are collected at different depths, the water temperature 

should be measured at each sampled depth in conjunction with measuring the temperature profile. 

Water samples cannot be determined to be within a specific littoral zone unless a depth profile is 

taken. Surface samples without temperature profile should be labeled as water surface samples 

rather than epilimnetic samples to avoid possible confusion. 

Point samples are those collected at a specific depth. Van Dorn cylinders and Kemmerer bottles 

are the most common point samplers for lakes. The Van Dorn has some advantage in allowing 

better circulation of water through the sample container. In general, use of a Van Dorn sampler is 

recommended for lake sampling. In either case, the sampler is difficult to keep clean unless it is 

kept in a plastic bag between sites. The sampler should be soaked in the lake before use. At the 

sample site, the sampler should be raised and lowered several times just below the surface to 

further rinse the container. It is then lowered to the desired depth, held to stabilize it, and 

triggered, usually with a weight that slides down the line holding the sampler. When sampling the 

hypolimnion, care should be taken not to touch the bottom as this will disturb sediment that could 

contaminate the sample.

Because of water drag on long lengths of rope and on the sampler itself when submerged, both 

Van Dorn and Kemmerer samplers are prone to sampling at shallower depths than indicated by 

the length of the rope. If the boat is drifting due to current or wind, deep samples may be in error 

for reported depth. This error can be avoided by anchoring the sampler or tying the rope to a fixed 

buoy. These samplers also tend to plane while being lowered; allowing the rope to straighten 

before triggering the bottle can help minimize this error. 
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Pumps and tubing are sometimes used to collect point samples from lakes and streams or to 

integrate samples from lakes. The use of tubing offers the ability to collect an integrated sample 

of the water column in lakes by lowering the tubing at a constant rate of travel while collecting 

samples downward to near the bottom and then back up to the surface. In general, however, lake 

water sampling for inventory and monitoring of the effects of atmospheric deposition does not 

require collection of depth-integrated samples with a tube.

The primary concern in using these devices is keeping the tubing clean. It is impossible to 

thoroughly clean the inside of a tube in order to eliminate bacterial growth. Sampling tubes can 

generally be kept relatively clean by storing them filled with DIW and in a black plastic bag (to 

reduce light and inhibit bacterial growth) and avoiding disturbance and uptake of sediment while 

sampling near the bottom. The tubing also can be easily blocked with slush during freezing 

conditions. 

At the time of this writing, a lake sampling video is being prepared for use in field-staff training. 

All field personnel who will be involved in lake sampling should view this video, which can be 

found on the FS ARM program website: <http://www.fs.fed.us/air>.

1.4.3 ON-SITE MEASUREMENTS

EVALUATION OF SITE CHARACTERISTICS

The field notebook and all field forms should be filled in while personnel are at the sampling 

location so the sample can be accurately linked to field data and observations. Field notebooks are 

very helpful at sites used for long-term monitoring to provide easy access to location information 

and maps, historical information on site characteristics, and field data collected during previous 

years.

Observations and impressions made by the field teams at the sampling location and elsewhere on 

the target stream or lake are extremely useful for ecological value assessment, evaluation of 

general water body condition, and data verification and validation. Thus, it is important that 

observations made by the field team about lake, stream, or watershed characteristics and 

conditions be recorded while the field personnel are in the field. Field data forms and field 

notebooks are provided for this purpose. The forms are designed as a guide for recording 

pertinent field observations. Field data entry forms are never considered to be comprehensive;

any additional observations made by the field crew that might eventually be useful in making a 

site condition assessment should be recorded in the "Comments" section or the field notebook. 

Team members complete the form at the end of the sampling, taking into account all observations 

made while on site.

STREAM STAGE AND DISCHARGE AND LAKE LEVEL

Stream Stage and Discharge 

The most valuable non-chemical measurement for interpreting stream chemistry data is often 

stream flow (also referred to as discharge), which is measured as volume of water per unit time. 

Variations in stream flow reflect precipitation and the different pathways water takes to reach the 

stream channel. 
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During low flow conditions, water discharging into the stream channel has usually had 

opportunity to pass well below the surface soil into deeper soils, till, and/or bedrock. Such deeper 

flow paths provide greater contact between water and the soils and geologic materials in the 

watershed. As a consequence, base flow commonly receives larger quantities of weathering 

products that can buffer acidity, raise pH, and increase concentrations of base cations in solution. 

During high flow periods, some water enters the stream channel through shallow flow paths that 

more clearly reflect the chemistry of upper soil horizons. Shallow flow paths tend to result in 

lower concentrations of base cations in drainage water, less acid neutralization (sometimes 

reflected in increased water acidity), and higher concentrations of DOC than deeper flow paths.

The United States Geological Survey (USGS) is the recognized leader in development and 

implementation of flow measurements. USGS protocols and recommended equipment for 

measuring flow are detailed in Rantz (1982). The following provides a synopsis of this material. 

In addition, USGS has produced a training video for measuring discharge. It is available on the 

Web at <http://wwwrcamnl.wr.usgs.gov/sws/SWTraining/WRIR004036/Index.html>. Field 

personnel should review this video as part of their field training program. 

To measure stream flow, some type of channel control is necessary. This control may be 

constructed as a temporary feature, such as a weir or dam, or a natural control, such as a bedrock 

outcrop or channel-width restriction. An effective control provides a predictable relationship 

between water level (stage) and flow that does not change over time. A pressure-transducer 

installed in the deepest part of the stream channel, just upstream of the channel control, can be 

used to record the water level (commonly at 15 minute intervals). A line must be secured in the 

stream to transmit the response of the pressure transducer to a datalogger. The pressure transducer 

measures changes in stage; these stage measurements then must be converted to estimates of flow

through the channel control. Alternatively, stage can be measured using a measuring rod or 

yardstick held vertically in place at a specific location. 

To establish the relationship between stage and flow (referred to as a rating curve), simultaneous 

stream stage and flow measurements are needed over as wide a range of stream flows as possible. 

To conduct the stream flow measurements, a cross-section is chosen in the general vicinity of the 

location where the water level measurements are taken. The ideal cross-section chosen for 

measurement should provide a regular cross-sectional channel shape that provides laminar flow 

throughout the channel. The more closely these conditions are met, the more accurate will be the 

resulting estimates of discharge. 

Stream flow measurement determined with a single stream velocity measurement is not sufficient 

for obtaining an accurate representation of discharge. The cross-section is divided into intervals 

such that at least one pair of depth and stream velocity measurements can be made in each 

interval. The number and width of the intervals are dependent on the shape of the cross-section. 

An alternative, common method of estimating stream velocity relies on measuring the velocity of 

a neutrally buoyant object, such as a small orange, traveling downstream. This approach can 

provide grossly inaccurate flow estimates as the object can follow preferentially rapid flow paths 

or, conversely, be temporarily impeded by stones or wood in the channel. This protocol does not 

recommend use of this method for estimating stream discharge. 
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The stream velocity can vary considerably through the cross-section by course and depth, 

requiring a number of velocity measurements to obtain an accurate flow measurement. A large 

variety of stream velocity meters with varying precision and accuracy are available commercially. 

The velocity of each interval is multiplied by the cross-sectional area of that interval and the 

products of all intervals are summed to provide the estimate of stream flow. 

The salt dilution gaging method also provides an approach for estimating stream flow at locations 

where reliable measurements with flow meters are not possible. Such conditions occur in streams 

that are highly turbulent or have irregular channels,, in small streams, and during low flow under 

conditions when a large part of the flow travels through gravel and rocks in the stream bed. For 

such conditions, salt dilution gaging provides a more reliable method for discharge measurement. 

Salt dilution gaging involves the addition of a known quantity of salt upstream of the gaging site, 

either by a single addition or by continuous injection. Discharge is computed based on the 

concentration or dilution of the salt, determined by conductivity measurements, as it passes the 

gaging site. 

Making flow measurements during periods of high flow may not be safe or it may not be possible 

to wade into the stream under such conditions. If there is any doubt about the safety of wading 

under the existing flow conditions, field staff should not enter the stream. 

If information on flow is needed to aid in the interpretation of stream chemistry measurements 

but neither installation of a stream gage nor collection of flow measurements are feasible, water 

level can be manually recorded from a staff gage at the time that the water sample is collected. 

This will provide data on stage, though not on discharge. For some research or monitoring project 

objectives, relative differences in stream stage may be sufficient in place of the more quantitative 

discharge data. 

The staff gage should be located just upstream of an effective control. The staff gage is usually 

comprised of a pressure-treated post or metal fence post anchored in the stream with a large ruler 

attached. The elevation of the ruler should be surveyed in reference to an object near the bank that 

would be considered immovable. This enables future verification that the staff gage has not 

moved. If the chemical concentrations that are being measured are statistically related to flow, 

they are also related to stage, although the relationship can differ. Changes in chemical 

concentrations of flow-dependent constituents can be estimated from stream stage measurements 

in a manner similar to that done with flow measurements.

Another approach to estimating discharge is to characterize regional flow conditions based on 

nearby gages on similar watersheds. Using this approach, it is possible to obtain a general idea of 

the likely flow conditions at the sampling site at the time of sample collection without the need 

for site-specific measurements. It is important to note that, although discharge can be a very 

useful parameter in evaluating the effects of atmospheric deposition, it is not absolutely essential. 

One should not choose to avoid sampling a site for water chemistry simply because it is 

impractical or impossible to collect parallel data on discharge.
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51

Lake Level

For interpreting lake chemistry data, the lake level can be especially useful. Of particular 

importance is the likelihood that lake chemistry will vary with precipitation cycles. During 

drought periods, a higher proportion of inflowing water may follow relatively deep flowpaths, 

allowing for greater acid neutralization and base cation mobilization. During wet periods, 

drainage water may preferentially follow shallow flowpaths, allowing less contact with soils and 

geologic materials and therefore limited acid neutralization. Seepage lakes may receive 

proportionately greater inflow of groundwater (which, depending on geological and soil 

conditions, may be rich in base cations) during drought periods. It is also possible that some 

seepage lakes might lose their connection with the groundwater during drought, causing the 

opposite effect. 

The extent of such influences on hydrology and consequent acid neutralization are expected to be 

region- and watershed-specific. Measurement of lake level at the time of sampling can provide 

critical data to help identify such effects. The simplest way to collect such data is to install a fixed 

staff gage in the lake and record the lake level at each time of sampling. In wilderness settings or 

other locations where installation of a staff gage is not allowed or is impractical, relative lake 

level can be documented by measuring the vertical height of the lake surface below a fixed 

landmark: for example, a large shoreline rock or tree root.

ANCILLARY DATA

Ancillary data is chemical and physical information that is not critical, but may be useful for 

analysis. This includes both chemical and physical data.

Chemical Data

Physico-chemical data can be collected on site with field equipment for measurements such as 

pH, specific conductance, DO, and turbidity. This information can be useful for reconnaissance 

work and sample site selection or for other investigations in which real-time information is 

needed to direct field activities. In general, however, this protocol recommends that assessments 

or monitoring of acid-base chemistry or nutrient status should be made using chemical analyses 

conducted in the laboratory rather than the field. Commercially available field equipment can 

produce data of quality similar to that of laboratory equipment for some variables; however, 

reproducing the clean, controlled environment of a laboratory in the field is difficult. Therefore, if 

real-time data are not required in order to satisfy the objectives of a particular study or other 

requirements7

, conducting the chemical analysis in the laboratory is recommended to ensure high 

data quality. If chemical analysis will be required in the field, pre-mobilization and field 

calibration checks should be conducted. Other types of chemical measurements can also be made 

in the field with commercially available analysis kits, but the data obtained using these methods 

typically only provide a rough approximation that may not be sufficiently accurate or precise for 

inventory or monitoring. 

 7 For example, in situ measurement of pH is sometimes required for data used by state 303(d) water quality 

assessment programs to determine waters to be classified as water quality limited according to the Clean Water 

Act.
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Meters with probes that continuously monitor pH and specific conductance are also available. 

These probes effectively characterize temporal variability but lack the precision and accuracy of 

laboratory measurements. The need for temporal resolution should be weighed against data 

quality objectives, logistics, and costs to determine if in situ monitoring is advantageous.

Physical Data 

Interpretation of water chemistry data can be significantly aided by ancillary physical 

measurements, such as air and water temperature, weather conditions, recent precipitation, and 

snow water equivalence (see the list of Natural Resource Manager Air application [NRM Air]

minimum database requirements in Section 1.4.5, Sample Documentation, below). Additional 

data might also be collected at the sampling site, depending on the study. 

For example, it can be useful to develop lake thermal profiles to evaluate the extent of lake 

stratification. For stratified lakes, it can be useful to collect water samples from the hypolimnion. 

Such data can be used in evaluating sulfur reduction in the lake sediment, hypolimnetic DO 

depletion, or the dynamics of phosphorus retention and release in lake sediments. Unless detailed 

analyses of sulfur or phosphorus cycling are to be conducted, however, hypolimnetic samples are 

generally not needed. 

Studies of temporal trends in surface water quality or characterization of water quality conditions 

within a specific lake or stream or across a forest can be designed to assess a variety of 

parameters and changes in those parameters over time. For this to be successful, information is 

often needed on watershed aspects that can impact or aid in interpretation of water quality. These 

can include, for example, the variables listed in Table 1-13.

The ecological significance of aquatic ecosystem degradation and loss due to physical habitat 

alterations can exceed the effects of atmospheric deposition or human activities on water 

chemistry. Therefore, physical habitat surveys of lake shore areas, littoral zones, stream channels, 

and riparian zones can be useful in conducting overall habitat condition assessments and in 

interpreting water chemistry data. Habitat information is helpful in estimating what lake or stream 

biological assemblages "should" be in the absence of many types of anthropogenic impacts. The 

physical evaluation can provide a reproducible, quantified estimate of habitat condition, serving 

as a benchmark against which to compare future habitat changes from anthropogenic activities or 

extreme events. Furthermore, habitat information can aid in the diagnosis of probable causes of 

ecological impairment in lakes or streams.

In addition to information collected in the field by the shoreline, stream channel, or littoral zone 

surveys, the physical habitat description of each lake or stream can include many map-derived or 

measured variables, such as lake surface area, shoreline length, stream width-to-depth ratio, and

habitat integrity or complexity. Furthermore, ancillary information, including watershed 

topography and land use, supplements the physical habitat information. The shoreline, channel, 

and littoral surveys concentrate on information best derived "on the ground." As such, these 

survey results provide part of the linkage between large watershed-scale influences and those 

forces that directly affect aquatic organisms day-to-day. Together with water chemistry, the 

habitat measurements and observations describe the variety of physical and chemical conditions 

that are necessary to support biological diversity and foster long-term ecosystem stability. 
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

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Table 1-13. Ancillary measurements that may help in interpretation 

of lake or stream water chemistry data.

Location Measurements

Streams Discharge and/or stage

Water temperature

Air temperature

Snowpack depth and snow water equivalence

Precipitation

Watershed morphometry

Fish stocking and management

Watershed disturbance1

Lakes Secchi disk transparency

Thermal profile

Chlorophyll a

Level (if lake is to be sampled multiple times)

Hypolimnetic water samples

Littoral zone water samples

Dissolved oxygen

Presence of inflowing or outflowing streams

Watershed morphometry

Fish stocking and management

Watershed disturbance

1 Watershed disturbance can be evaluated by field reconnaissance, 

examination of aerial photos, regional land cover data sets, etc.

Habitat surveys should not be considered a necessary component of inventory and monitoring. 

They require an additional commitment of time and resources. Nevertheless, the data collected in 

such surveys can be very helpful in the subsequent interpretation of effects, especially if the 

documentation or quantification of effects relies on collection of biological (i.e., phytoplankton, 

zooplankton, benthic invertebrates, and fish) as well as chemical variables. 

The shoreline and littoral habitat surveys conducted by EPA in the EMAP program employed a 

randomized, systematic design with 10 equally spaced observation stations located around the 

shore of each sample lake. Teams went to the field with pre-marked lake outlines showing the 

station locations. The observations at each station included quantitative and semi-quantitative 

observations of vegetation structure, anthropogenic disturbances, and bank substrate. In-lake 

littoral measurements and onshore observations dealt with littoral water depth, bottom substrate, 

near-shore fish cover, and aquatic macrophyte cover. With quantifiable confidence, investigators 

condensed these observations into descriptions applicable to the whole lakeshore and littoral 

zone. For example, team observations led to quantitative descriptions, such as the mean canopy or 

aquatic macrophyte cover along the lakeshore, the extent of shoreline disturbed by various human 

activities, and the dominant littoral substrate in the lake. There are similar physical habitat 

evaluation procedures for streams developed for EPA’s national surveys, such as the Wadeable 

Stream Survey.
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1.4.4 POST COLLECTION SAMPLE PROCESSING, DOCUMENTATION,

AND CLEANUP

In general, this protocol does not recommend filtering lake or stream samples in the field except 

where immediate filtering is required for a particular measurement. To avoid the possibility of 

sample contamination, it is generally preferable to perform this step within the controlled 

conditions of a laboratory. Similarly, measurement of pH in the field is not recommended for 

most studies. This measurement is best performed in the laboratory under controlled conditions. 

For most chemical constituents of interest for atmospheric deposition studies, sample preservation 

in the field is not necessary. Types of sample preservation may include addition of chemicals or 

filtering to remove particulates. Preservation procedures are generally done for a specific 

measurement and usually render the sample unusable for other measurements. Therefore, if 

preservation in the field is needed for a specific measurement, an aliquot will need to be removed 

from the sample before preservation. The volume of the aliquot will be dependent on the 

analytical requirements of the measurement. 

Most samples contain dissolved and/or particulate organic matter and associated microbes that 

can change sample chemistry through decomposition and assimilation. All samples should 

therefore be placed out of the sunlight and in a cooler with ice as soon as possible for transport 

back to the laboratory, where they can be refrigerated. This procedure is usually sufficient to slow 

biological processes enough to prevent measureable changes in chemical concentrations. 

As a rule of thumb, samples should be returned from the field as quickly as possible to enable 

processing in the laboratory, and filtration (if needed) should be performed in the laboratory. An 

exception applies to the collection and analysis of samples for measurement of chlorophyll a:

those samples are most commonly filtered in the field and the filter (not the filtrate water) is 

transported in a zipper lock bag on ice in the dark to the laboratory for analysis. It is essential to 

record in the Notes section of the sampling record forms the volume of water that was filtered for 

chlorophyll a measurement.

1.4.5 SAMPLE PACKAGING AND TRANSPORTATION

Filled sample bottles should be placed in zipper lock bags before transport from the field site. 

Syringe stopcocks should be taped shut to keep from leaking, and syringes should be secured 

from damage (e.g., placed in a paper towel roll) and packed in a plastic box with snap-on lid for 

transport and shipping. Each syringe box needs to be long enough to hold syringes that are 2⁄3 to 3⁄4 

filled with sample water and wide enough to hold multiple syringes. The bagged sample bottles 

and boxed syringes should be packed with double-bagged ice or frozen refrigerant for transport

(Figure 1–1).
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

55

Figure 1–1. Sample syringe packaging for transportation to the lab. Top left: stopcock taped; top 

right: syringes and packing tubes; bottom: packing tubes packed in a shipping box.

Insulated containers, with double-bagged chemical refrigerant (“blue-ice”) or (preferably) with 

double-bagged ice blocks, are needed for transport of collected samples between the field and 

other staging locations and, eventually, the laboratory. Ice works better for shipping unless large 

numbers of chemical refrigerant packs are used. Small insulated containers that will fit into 

backpacks can be used to carry and protect the samples in the field. Coolers can be used for 

assembly and transport of samples in vehicles. 

Chemical refrigerant containers should be packaged in two zipper-lock plastic bags to minimize 

the possibility of sample contamination through refrigerant leakage. Field crews will need to 

make sure that the chemical refrigerant was placed in a freezer at least two days before sampling

to completely freeze the refrigerant.

SAMPLE DOCUMENTATION 

Sample documentation should be completed in the field. Documentation includes completing and 

affixing all sample labels, completing all field and chain-of-custody forms, and recording field 

observations and site condition information. A list of minimum database requirements for the 

FS’s NRM Air database are:

• Forest name;

• Wilderness name (if applicable);

• Official USGS lake/stream name;

• Site name;

• Site ID;

• Field team leader and contact information;

• Latitude and longitude (decimal degrees);
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• Datum used (use NAD83 if possible);

• Monitoring project name;

• Date of visit;

• Time samples were collected;

• SampleID;

• Barcode;

• QA sample type (regular, duplicate, blank, split, etc.);

• Sample measurement location (e.g., inlet, outlet, deep, shore (lake), bank (stream)); and

• Sample method (e.g., grab, pole).

There may be other requirements according to the individual study. Note that this protocol does 

not require collection of temperature or stream flow data, although it does recognize that such 

data can be very useful in interpreting the results of chemical analyses. 

Documentation should also include a review of sampling procedures used, labeling, and 

photographic/written documentation. Key issues include:

1. Were the samples collected at the designated depth and free of influence from any sediment 

that could be disturbed during sample collection?

2. Were all sample bottle and sample syringe labels fully completed with all required 

information?

3. Was the sampling fully documented, including site photographs (if appropriate), completed 

field and chain-of-custody forms, and field notes?

4. Were any conditions or circumstances noted that could potentially compromise or influence 

the chemistry of the sample?

POST-SAMPLING EQUIPMENT CLEAN-UP 

Clean-up is important to minimize the possibility of transporting pathogens, noxious species, or 

invasive species from one sampling location to another. Risks vary from region to region and 

location to location. Field personnel should consult with regional and local FS offices for specific

problem identification and appropriate precautions and cleaning protocols. A variety of forest 

pathogens and aquatic and terrestrial invasive species may be of concern, depending on location. 

Personnel and their boots, vehicles, boats, and equipment can serve as transport vehicles for 

problematic species; thorough cleaning of equipment, boots, etc. should be done before leaving 

the site. Wherever a risk exists, field personnel should take additional site-specific appropriate 

risk-management precautions.
SECTION 1. WATER CHEMISTRY FIELD SAMPLING PROTOCOLS

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1.4.6 SEQUENCE OF FIELD ACTIVITIES

The recommended sequence of field activities to be conducted by field personnel at the sampling 

site is as follows:

1. Select/verify sampling site location.

2. Take photographs.

3. Fill out and affix the label for each sampling bottle to be filled at the site.

4. Evaluate and document site conditions.

5. Determine how many and what kinds of samples will be collected (e.g., QA replicates, 

single versus integrated samples, or special aliquots to be collected into glass bottle or 

syringe).

6. Collect water sample(s).

7. If required, preserve or filter (if necessary) or transfer selected sample aliquots to glass 

bottles or syringes (as appropriate).

8. Place collected samples in cold, dark storage container.

9. Collect any needed ancillary data.

10. Determine (if appropriate) stream discharge or stage.

11. Complete all site documentation and chain-of-custody forms.

12. Record all field observations in a field notebook.

13. Clean equipment, clothing, and boots to prevent spreading invasive species to another site.

1.4.7 SAFETY IN FIELD ACTIVITIES

KEY SAFETY CONSIDERATIONS 

For safety reasons, an emergency contact individual who is not part of the field crew should 

always know where the field crew is going each day and by what route. This person should be 

contacted by the field crew immediately upon return from the field each day. 

For sampling remote locations, safety equipment should include, but should not necessarily be 

limited to, the following: 

• Two-way radios and/or cellular telephones (if cell phone access is available in the study 

area);

• extra batteries for GPS units and radios;

• Rain gear;

• Emergency shelter blankets;

• An adequate supply of drinking water or an appropriate water filtration system;

• Sunscreen;

• First aid kit; and

• Locally required safety equipment.
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All field personnel should have current first aid and CPR certificates. Field personnel should 

never enter a deep or fast-flowing stream without wearing a personal floatation device (PFD). A 

PFD should also be worn when working close to a stream during high flow conditions. 

When sampling a lake at the mid-lake location it is advantageous for one person to remain on 

shore to provide logistical support, to record data, and serve as a safety precaution. If necessary, a 

two-way radio can be used to facilitate communication between mid-lake and shore personnel. 

Sampling from a boat typically requires two people in the boat, one maintaining the position at 

the sampling site and the other collecting the samples and measurements. The sampler can 

monitor present position by keeping two appropriate on-shore landmarks in line. The on-shore 

person should continuously monitor the float tube location in a similar way. 

JOB HAZARD ANALYSIS 

Field personnel should review the Job Hazard Analysis (JHA) before going into the field and 

construct a field itinerary before field work. The itinerary should include:

• Departure date and time;

• Expected return date and time; and

• Expected route of travel (roads, trailheads, trails, and destinations).

An example JHA is provided in Appendix I. A location-specific JHA should be created and 

periodically updated by the project.
SECTION 2. LABORATORY PROTOCOLS

59

SECTION 2. 

LABORATORY PROTOCOLS G.B. Lawrence, J.R. Webb, T.J. Sullivan, A.T. Herlihy

2.1 INTRODUCTION

Surface water samples collected in watersheds on forest lands with minimal recent influence from 

development or agriculture tend to have low concentrations of nutrients. Because these types of 

watersheds are often located in upland areas with rocky, infertile soils, their surface waters may 

have low ionic strength, which means most other dissolved constituents are also low in 

concentration. Chemical analysis of dissolved materials at low concentrations sometimes requires 

modification of the standard methods used for sample preparation and analysis. Therefore, the 

laboratory selected for analyzing low-nutrient, low-ionic strength surface water samples should 

not only be experienced with these types of samples, but should 1) analyze them on a routine 

basis, and 2) provide data with reporting limits sufficiently low for project needs. 

To produce high-quality data, a laboratory should have effective procedures in place for each of 

the following elements:

• Bottle cleaning;

• Sample processing; and

• Chemical analysis.

Each of these elements is described in the sections that follow. A fourth element, QA/QC, is 

covered in the QA/QC section of this document (Section 3). In addition, the laboratory should 

follow effective procedures for documentation of method implementation and method changes. 

This report provides information on all major aspects of successfully operating a laboratory for

low-nutrient, low-ionic strength water analysis.

2.2 LABORATORY PREPARATION PRIOR TO SAMPLE

ANALYSIS

2.2.1 Bottle Cleaning

All bottles used for sample collection and partitioning the sample into aliquots for transport to the 

laboratory must be clean and free of any contamination. Generally, it is the responsibility of the 

laboratory to provide clean bottles to the field crew in advance of field sampling. Low-nutrient, 
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60

low-ionic strength samples can be easily contaminated by improperly washed bottles because 

their low concentrations can be measurably altered by trace amounts of contaminants. Therefore, 

rigorous cleaning procedures must be used, which are often specific to the intended use of the 

bottle. Laboratories experienced with low-nutrient, low-ionic strength surface water samples have 

adopted various methods for cleaning laboratory plasticware and glassware. Washing with dilute 

acid (usually 1 or 2%) is generally preferred to solubilize potential contaminants that can then be 

removed with multiple DIW rinses. Acid washing should be done in a dedicated room with a 

negative pressure ventilation system. 

Rinse water should meet the specifications of Type III water (American Society for Testing and 

Materials [ASTM] 1984). A typical water deionization system that produces Type III water 

includes, in sequence, a carbon removal tank, 1 μm filtration, and two mixed-bed cation-anion 

removal tanks. To ensure consistent water quality, specific conductance (or resistance) of the 

water should be monitored between the two primary treatment modules, in this case the mixedbed tanks. If the specific conductance exceeds the preset limit of 1.0 μS/cm, an indicator light 

(which is normally on) is deactivated. The system is checked daily, and if the indicator light is 

off, the first tank is removed and replaced by the second tank. A new tank is then placed in the 

second position. By monitoring the specific conductance of the two tanks, the initial tank can 

serve as a first level treatment and the second tank as a polisher. By switching the polisher to the 

first position when the first level tank no longer meets the standard, you can extend the lifespan of 

the tanks and, most importantly, prevent substandard water from being used for sampling. The

functionality of the tanks degrades gradually, so the two-stage approach is needed to maintain a 

consistent level of water quality. The filter is replaced with every tank change, and the carbon 

tank is replaced in response to the volume of water being treated and the organic carbon 

concentration of the influent. Once every six months is a typical replacement frequency8

.

To ensure that the wash acid does not itself become a contaminant, repeated rinsing is followed 

by leaching the bottle with DIW. This is done by filling the rinsed bottle with DIW and storing it 

for 24 hours or longer (Table 2-1). This step is necessary because acid and other contaminants 

(such as those from a previous sample) can migrate into the plastic matrix of the bottle wall. Over 

time, the contaminants can slowly leach out and affect the sample concentration. The level of 

contamination caused by this process is usually low but can be sufficient to cause measureable 

increases in low-nutrient, low-ionic strength samples. Simple rinsing does not necessarily 

eliminate this type of contamination. 

Finally, each bottle is filled with DIW before shipping the bottles to the project location. 

Measurement of specific conductance of DIW stored in sample bottles, with an acceptance 

criterion of < 1.2 μS/cm9

, provides quality assurance for this procedure. Specific conductance 

testing can be done on selected sample bottles that are treated as sample blanks. 

Aliquot bottles that have had acid added for sample preservation are more likely to have 

contamination from bottle leaching than those that are in contact with wash acid for 30 minutes or 

 8 Note that appropriate methods for tank replacement are to some degree equipment-specific. The procedure 

described here is one possible approach to producing thoroughly rinsed bottles. Other procedures may also be 

acceptable.

9 We recommend using an acceptance criterion of ≤ 1.2 μS/cm, but we recognize that some laboratories use a 

different criterion: as high as about 2.0 μS/cm.
SECTION 2. LABORATORY PROTOCOLS

61

less. Therefore, preserved sample aliquot bottles require longer periods of DIW leaching. Plastic 

bottles should not be used if the caps have liners because the liners can become a source of 

contamination. Glass bottles, such as those needed for DOC aliquots, usually have caps with 

removable plastic liners. The plastic liners must be removed from the caps for washing and be 

soaked in DIW for the same length of time that the bottle is soaked before being replaced in the 

cap. 

Because the required cleaning procedure depends on the specific use of the bottle, a set of 

cleaning procedures needs to be developed and documented by the laboratory. Appropriate 

procedures to clean the various types of collection and aliquot bottles are listed below and 

summarized in Table 2-1. Although different laboratories may use different or modified 

procedures, it is critical to document the specific procedures used and to provide assurance that 

sample bottle contamination has been avoided.

Additional precautions may be needed when washing with acid solutions such as hydrochloric 

acid (HCl). Local regulations may require separate disposal of acid (or basic) waste products 

(depending on the concentration), rather than pouring them down the sink. The possible need for 

special waste disposal procedures may affect the laboratory budget and may influence decisions 

regarding bottle washing procedures.

Table 2-1. Summary of a typical bottle rinsing protocol for low-nutrient, low-ionic strength 

samples.

Bottle Acid-Wash? Rinses DI Soak Period

60 mL plastic (cations, acidified) Yes 4

1 wk, 2X rinse, 24 hr 

soak

30 mL plastic (anions) No 6 24 hr

250 mL plastic (pH, ANC,...) Yes 4 24 hr

30 mL plastic (total Al, acidified) Yes 4

1 wk, 2X rinse, 24 hr 

soak

30 mL plastic (NH4 or DON) No 6 24 hr

40 mL glass (DOC) Yes 4 24 hr

DOC caps and liners Yes 6 24 hr

1 L plastic (field, manual collection) Yes 4 24 hr

500 mL plastic (field, autosample) Yes 4 24 hr

1L plastic (field, autosample) Yes 4 24 hr
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CLEANING BOTTLES USED FOR SAMPLE COLLECTION IN THE FIELD 

1. Remove sediment and other particles from the bottle with tap water, using a soft plastic 

brush10 if necessary, then rinse once with deionized (DI) water.

2. Fill with 2% HCl11 and let stand for 15 to 30 minutes in a separate bottle-washing room, 

away from analytical instrumentation.

3. Pour out the HCl and rinse thoroughly four times with DIW.

4. Fill with DIW and store for at least 24 hours.

5. To prepare for transport to the field, empty and rinse once with DIW, then fill with DIW for 

transport to the field.

6. The bottle should subsequently be rinsed three times in the field with the sample that is 

being collected before filling the bottle.

CLEANING BOTTLES USED FOR GENERAL LABORATORY USE, UNACIDIFIED 

ALIQUOTS

1. Empty any remaining sample.

2. Fill with 2% HCl and let stand for 15 to 30 minutes.

3. Pour out the HCl and rinse thoroughly four times with DIW. (HCl can be reused for 

multiple washings but should be replaced and properly disposed of when it becomes 

discolored.)

4. Fill with DIW and store for at least one week.

5. To prepare for use, empty and then rinse twice with DIW.

CLEANING BOTTLES USED FOR GENERAL LABORATORY USE, ACIDIFIED 

ALIQUOTS

1. Pour out any remaining sample.

2. Rinse four times with DIW.

3. Fill with 2% HCl and let stand for 15 to 30 minutes.

4. Pour out the HCl and rinse thoroughly four times with DIW.

5. Fill with DIW and store for at least 24 hours.

6. To prepare for use, empty and rinse once with DIW.

Some labs avoid the acid wash step for aliquots to be analyzed for Cl

-

, NH4

+

, and/or DON, and 

instead rely on DIW leaching to remove any contaminants from the bottles. Depending on the 

intended use of sample aliquot bottles, the recommended number of rinses can vary. These 

differences are summarized in Table 2-1. As indicated above, specific conductance should be 

 10 Note that use of a stiff bottle brush can scour the inside of the bottle, allowing contaminants to more easily adsorb 

to the bottle wall. 11 Note the need to use HCl in a separate bottle washing room, away from analytical instrumentation. HCl can 

become volatilized and thereby contaminate nearby samples and, eventually, damage equipment.
SECTION 2. LABORATORY PROTOCOLS

63

measured for DIW stored in sample bottles as a quality assurance measure. Such analyses should 

be conducted, at a minimum, on a subset of bottles before field use and on sample blanks during 

laboratory analysis.

2.2.2 Sample Processing, Preservation, and Storage

When samples arrive at the laboratory, they need to be accompanied by proper documentation 

using a chain-of-custody form. This form provides field information that includes project 

identification: when, where, how, and by whom the sample was collected; and information on the 

chain-of-custody that was followed. This form will need to have been checked by the field 

sampler against the information written on the sample bottle label to ensure that the information 

matches. A format for recording sample information in the field has been developed (Appendix 

E). The transfer of custody from field personnel to lab personnel must be documented by dated 

signatures on this form. A copy of the signed form should be kept by both project and laboratory 

personnel. This procedure is needed to ensure that samples were collected and transferred to the 

laboratory. Samples can get misplaced before arriving at the laboratory or within the laboratory 

before processing, particularly if there was a sample labeling error. 

Prior to the start of sample processing in the laboratory, a unique code or sample serial number 

(SSN) is typically assigned by the laboratory and added to the laboratory data sheet. A single 

person (plus a trained backup) is generally assigned the responsibility to ensure that an SSN is not 

accidently used for more than one sample. If the SSN is assigned in the field, it is important that it 

is unique. The SSN will be used by the laboratory to track the sample through the steps of sample 

processing, chemical analysis, and data management. The information on the chain-of-custody 

form and laboratory data sheet will be entered into an electronic database through the use of a 

Laboratory Information Management System (LIMS). A variety of LIMS software is available 

commercially, though some laboratories develop their own database system.

Each sample will typically be analyzed for a variety of constituents that are chosen to meet stated 

project goals and require specific processing procedures. Different results may be obtained from 

the same analysis if samples are prepared for analysis using different procedures. For example, 

results may differ if samples are filtered with filters of different pore sizes or of different 

materials. Sample processing can involve both preparation for analysis and preservation of the 

sample, and therefore, varies among analyses. The details of processing and analyses must be 

established to ensure that both sample processing and analyses done by the laboratory meet 

project needs.

Processing is generally accomplished by dividing the sample into several aliquots, each with its 

own filtration/no filtration, preservation, storage, and process and handling time requirements. 

For example, analysis for concentrations of base cations (calcium, magnesium, sodium, and 

potassium) may require filtration through a 0.45-μm polycarbonate filter, whereas analysis of 

DOC may require filtration through a glass fiber filter in order to avoid possible organic 

contamination from the polycarbonate filter. Preservation of samples for analysis of base cation 

and other metals that could form precipitates at non-acidic pH values usually involves the 

addition of nitric acid. However, addition of chemicals to samples should be avoided unless 

necessary to reduce the potential for sample contamination or alteration.
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64

All samples that require filtering to remove particulate matter are normally filtered as soon as 

possible after arrival at the laboratory. Prompt filtration after sample collection is normally done 

for the purpose of removing bacteria, which can alter sample chemistry through their metabolic 

processes. Samples should be chilled as soon as possible after collection and kept refrigerated up 

to the time of processing and preservation in order to retard microbial activity. Some aliquots will 

continue to be refrigerated until analysis (Table 2-2). Freezing is not necessary for most analytes 

but is recommended for DON and NH4

+

. 

The containers used to store aliquots before analysis also vary by analyte. Aliquots for base cation 

analysis are generally stored in polyethylene or polypropylene bottles, which are economical and 

considered sufficiently inert with respect to base cations, whereas aliquots for DOC analysis are 

usually stored in glass bottles to avoid organic contamination from plastic. An example of a 

typical sample processing schedule for low-nutrient, low-ionic strength samples is shown in 

Table 2-2. It is important to establish this type of schedule with laboratory personnel to ensure 

that all samples from a particular project will receive timely processing. In this example, 

reminders are included to tape bottle caps and not fill bottles completely for aliquots that are 

preserved by freezing.

Table 2-2. Example laboratory aliquot schedule for a particular project.

Aliquot1 Container Filter Treatment Storage

A 250 mL polyethelene None None Refrigerator (4oC)

B 30 mL polyethelene 0.45 μm polycarb. None Refrigerator (4oC)

C 40 mL glass2 Glass fiber filter (GFF) None Refrigerator (4oC) 

D 60 mL polyethelene 0.45 μm polycarb. 0.3 μL HNO3 Room temp.

E 30 mL polyethelene (taped) GFF - fill 2/3 full None Freezer (label DON)

F 30 mL polyethelene (taped) None - fill 2/3 full None Freezer (label NH4)

3

1 A - pH, ANC, specific conductance, Alm, organic monomeric Al

B - Sulfate, nitrate, chloride

C - Dissolved organic carbon

D - Calcium, magnesium, sodium, potassium, silicon

E - DON

F – NH4

+

2 Use of glass for storing samples in the laboratory before DOC analysis is preferred but not essential.

3 Ammonium concentrations (although typically very low in natural waters) are very unstable. The sample should be analyzed 

immediately upon arrival at the laboratory or frozen until time of analysis. 

Recommended laboratory holding times are given in Table 2-3. Those laboratory holding times 

should be considered guidelines. Measurement of a sample analyte past the holding time is not 

justification for excluding that concentration value from the database. Nevertheless, sample 

measurements taken beyond the specified holding time should be flagged as such in the database. 

In general, laboratories should strive to complete analyses within the holding time windows, or as 

soon as possible thereafter. 
SECTION 2. LABORATORY PROTOCOLS

65

Table 2-3. Recommended laboratory holding times. (Source: USGS Troy Laboratory.)

Constituent Holding Time

pH 2 weeks

conductivity 2 weeks

ANC 2 weeks

ammonium 3 months1

dissolved nitrogen 3 months1

Alm 2 weeks

organic monomeric aluminum 2 weeks

calcium 6 months

magnesium 6 months

silicon 6 months

sodium 6 months

potassium 6 months

chloride 1 month

nitrate 1 month

sulfate 1 month

DOC 2 weeks

turbidity 2 days

1 Samples for NH4+ and dissolved N are preserved by freezing and 

analyzed in batches.

The temperatures of refrigerators and freezers that are used for sample storage must be 

electronically monitored around the clock to ensure that malfunctions do not result in temperature 

increases that compromise the samples. Various types of temperature monitors are commercially 

available to notify laboratory personnel via cell phone or computer that the temperature of a 

refrigeration unit has exceeded a preset threshold. With this notification, samples can then be 

quickly transferred to another unit until repairs can be completed.

Each laboratory should have detailed documentation of the steps used in sample processing. An 

example list of sample processing steps follows:

1. Obtain chain-of-custody forms that have assigned SSNs and aliquot labels that correspond 

to the processing selected for the project. Initiate laboratory data sheet. 

2. Retrieve clean aliquot containers and place appropriate numbered dots and/or label the tape 

on them. Put on gloves, empty the containers, and rinse them with DIW, if applicable.

3. Retrieve the field samples to be filtered from the refrigerator. If there is not sufficient 

sample volume to prepare all of the required aliquots, be sure to follow the procedure for 

low-volume samples.
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

66

4. Shake the field sample bottle. Rinse the aliquot bottles with a small volume of the sample. 

Fill aliquot containers with raw sample for the aliquots that do not require filtering. Fill in 

the letter code that corresponds to the aliquots on the laboratory data sheet to document the 

processing method.

5. Retrieve the filtering apparatus from the DIW soak and rinse it well with DIW. Set-up the 

filtering apparatus on the vacuum manifold.

6. Place an appropriate filter (handling the edge of the filter only) on the filter apparatus with 

tweezers that have been rinsed with DIW. For 0.4 and 0.1 micron filters, place the shiny 

side up when appropriate (filters are sometimes packaged shiny side down). Filter 10 mL of 

DIW through the apparatus and into a waste container, then filter 10 mL of the sample 

through the apparatus and into a waste container. Discard the filtrate.

7. Place proper aliquot container under filtering apparatus. Filter 5-10 mL of sample into

container, rinse, and then discard the filtrate.

8. Filter appropriate amount of sample into container. If another aliquot of the same sample 

requires the same filter, repeat starting at step 7. Fill in the letter code that corresponds to 

the aliquots on the laboratory data sheet. Discard the used filter and rinse the filtering 

apparatus with DIW.

9. Repeat steps 6 through 8 for each sample aliquot. If the filter clogs, replace with a new 

filter following step 6, then go to step 8.

10. After samples have been processed, rinse the filtering chambers with DIW and place them 

in DIW soak buckets. Replace the DIW in the buckets weekly.

11. Store the remaining sample volume for possible re-analysis, at least until QA/QC analyses 

have been completed. Once it has been determined that the analysis meets data quality 

objectives (DQOs), discard remaining sample in field sample bottles and bring the bottles 

to the bottle washing room.

12. Aliquots that require acidification should be acidified in the hood using the appropriate acid 

dispenser. 

13. Aliquots should be stored in the appropriate places as described in the specific project 

sample processing schedule.

14. Date and processor’s initials must be recorded on the laboratory data sheet. Completed 

chain-of-custody forms and laboratory data sheets should be filed in a safe location. 

Information on these forms will need to be entered into the LIMS. 

In some situations, it is possible that the sample bottle was not completely filled in the field. If 

there is insufficient sample volume for all aliquots, analysis of the sample volume available must 

be prioritized based on the objectives of the project. If it is anticipated that some low-volume 

samples will be collected, a low-volume schedule should be prepared and made available to 

laboratory personnel. An example of a low-volume schedule is provided in Table 2-4.
SECTION 2. LABORATORY PROTOCOLS

67

Table 2-4. Example low-volume sample schedule.

Priority Aliquot Type1

1 B - 15 mL

2 D - 15 mL

3 A - 50 mL

4 C - 15 mL

5 H - 15 mL

6 G - 20 mL

1 See Table 2-2 for the description of aliquot types. Fill aliquot bottles from the available sample 

volume to the appropriate aliquot volume in the listed order of priority.

2.3 CHEMICAL ANALYSIS

The constituents that need to be measured in a water sample will be determined by the specific 

objectives of the project. Each method of chemical analysis (method) will provide a constituent 

concentration with a certain level of accuracy and precision over a finite concentration range that 

is specific to that method. Low-nutrient, low-ionic strength waters generally require methods that 

are effective at the lowest concentration ranges. A variety of methods are usually available to 

determine the concentration of a given constituent, even at low concentration ranges. The method 

selected must 1) be appropriate for the expected concentration range, 2) provide the data with the 

accuracy and precision necessary to successfully achieve the data quality objectives specified in 

the Quality Assurance Plan, and 3) not exceed logistical limitations with regard to sample 

collection, sample preparation, or laboratory capabilities. For example, a method for determining 

ammonium concentrations might provide data over the necessary concentration range with a 

sufficiently high level of accuracy and precision but may require that the analysis be done within 

12 hours of collection. Such a short holding time might not be feasible for samples collected from 

remote sites or might be beyond the processing capabilities of the laboratory. 

Currently, the ARML performs most of the chemical analysis for the stream and lake water 

samples collected in the FS ARM program (see <http://www.fs.fed.us/air>). The following is a 

summary listing of instrumentation and techniques employed for sample analysis. Equivalent 

instrumentation and techniques can be substituted. Results should be reported by the laboratory 

using raw instrument units (which must be clearly specified) and converted to the recommended 

units (μeq/L for most analytes) at a later time. These raw instrument units will most commonly be 

μS/cm for conductivity, μeq/L for ANC, μg/L for Al, standard pH units, and mg/L for other 

analytes. 

pH (hydrogen ion) 

• Method: SM 4500-H+ B/EPA 150.1 

• Instrumentation: Metrohm/Brinkmann Titrator 

• Technique summary: Standard pH electrode

• Reporting units: standard pH units 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

68

Acid Neutralizing Capacity 

• Method: USGS NFM 6.6.4.C

• Instrumentation: Metrohm/Brinkmann Titrator

• Technique summary: Gran analysis technique (Gran 1952)

• Reporting units: μeq/L

Conductivity 

• Method: SM 2510 B/EPA 120.1

• Instrumentation: Metrohm/Brinkmann Titrator

• Technique summary: Electrometric (APHA 1998)

• Reporting units: μS/cm

Sulfate, Chloride, Nitrate 

• Method: SM 4110 B/EPA 300.0

• Instrumentation: Metrohm_MetrosepA

• Technique summary: Ion Chromatograph (IC) with separator column for anions (APHA

1998) 

• Reporting units: mg/L (must specify whether units are reported as S versus as SO4

2- and as N 

versus NO3

-

)

Calcium, Magnesium, Potassium, Sodium

• Method: ASTM D6919 - 03

• Instrumentation: Metrohm IC_Cation_M/D

• Technique summary: Ion Chromatograph (IC) with separator column for 

monovalent/divalent cations 

• Reporting units: mg/L 

Fluoride

• Method: SM 4110 B/EPA 300.0

• Instrumentation: Metrohm_MetrosepA

• Technique summary: Ion Chromatograph (IC) with separator column for anions (APHA 

1998)

• Reporting units: mg/L 

Neither monomeric Al nor DOC is currently analyzed at the ARML. Appropriate techniques for

those analyses include the following:

Aluminum, Total Monomeric and Nonlabile Monomeric 

• Method: McAvoy et al. (1992)

• Instrumentation: Lachat Flow Injection Analyzer

• Technique summary: Colorimetric detection with open-system samples by pyrocatechol 

violet technique. Fractionation with ion-exchange resin (Driscoll 1984, McAvoy et al. 1992). 
SECTION 2. LABORATORY PROTOCOLS

69

Labile (inorganic) monomeric Al is determined by subtracting the nonlabile monomeric Al 

concentration from the total monomeric Al concentration. 

• Reporting units: μg/L 

Dissolved Organic Carbon

• Method: U.S. EPA (1987)

• Instrumentation: Dohrmann Carbon Analyzer 

• Technique summary: Persulfate/UV oxidation with infrared detection (U.S. EPA 1987).

• Reporting units: mg/L 

The above methods will be suitable for water quality studies in a number of National Forests. 

Additional methods for sample processing and analysis for low-nutrient, low-ionic strength 

waters have been described by U.S. EPA (1987), Morrison (1991), Paulsen (1997), and Eilers 

(2007).

Other methods can be adopted depending on the type of water that is studied and the program 

objectives. For example, it may be important to obtain water quality data that contribute to state 

water quality management programs in order to ensure that National Forest waters are included in 

regional water quality assessments and remediation efforts. In many cases, this will require 

laboratory adherence to methods specified by the U.S. EPA for use in state implementation of 

Clean Water Act programs. These methods, however, may apply to standards or criteria 

associated with water quality issues and sample volumes that differ from particular National 

Forest concerns and water quality monitoring objectives. 

Selection of a method that is capable of meeting the required data accuracy and precision 

specified in the DQOs does not ensure that this level of data quality will be achieved. Rigorous 

QA/QC procedures must be followed as part of the method implementation. In this usage, quality 

control refers to procedures that identify results during chemical analysis that do not meet DQOs, 

thereby triggering immediate corrective action that usually involves reanalysis of that sample. 

Data quality objectives are generally based on the precision and accuracy levels required by the 

project and the laboratory and the analytical limits of the methods used. A key component of QC 

is the introduction of artificial samples of known concentration, which are associated with a 

specific set of project samples. Quality control procedures are generally focused on instrument 

performance.

Additional procedures related to QA, which are also evaluated by DQOs, are used to document 

laboratory performance through the introduction of artificial and natural samples that are not 

associated with a specific set of project samples but reflect the accuracy and precision of sample 

preparation and analysis, including instrument performance. Protocols for QA and QC are 

described in the QA/QC section of this report. 

Each method used to determine a chemical concentration involves a complex set of procedures, 

reagents, and instrumentation. Any variation in these factors can potentially change the result, 

yielding a different concentration value. Therefore, each method requires that the analyst adhere 

to a strict SOP each time that the method is implemented. All details of the method must be 

documented in the SOP, which must be available to any potential user of the data for review. 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

70

Each SOP must be dated, signed, and approved. Any change to an SOP must also be dated, 

signed, and approved.

We have not attempted to recommend specific SOPs for implementation of analysis methods

here. Prescription of specific detailed SOPs is neither desirable nor practical for long-term 

monitoring programs. SOPs are specific to individual laboratories and instrumentation. Analytical 

methods, and thus SOPs, will inevitably change as instrumentation and technology improve. 

Moreover, as indicated above, the specific analyses, methods, and details of SOPs should be 

determined by program objectives and DQOs. Rather than recommending specific SOPs for the 

FS ARM program, we recommend that each National Forest monitoring program adopt and 

adhere to SOPs that assure and document attainment of appropriate DQOs through all phases of 

data acquisition, including sample collection, handling, analysis, and reporting. Selection of 

DQOs and development of quality assurance plans is discussed in the QA/QC section of this 

report.

Example SOPs for laboratory analysis of low-nutrient, low-ionic strength surface waters are 

provided in Appendix G. Alternative SOPs may be used so long as QA/QC objectives are 

satisfied.
SECTION 3. QUALITY ASSURANCE/QUALITY CONTROL PROTOCOLS

71

SECTION 3. 

QUALITY ASSURANCE/QUALITY 

CONTROL PROTOCOLS 

A.T. Herlihy, T.J. Sullivan, G.B. Lawrence, and J.R. Webb

3.1 INTRODUCTION

The constituents to be measured in a water sample will be determined by the specific objectives 

of the project. Each method of chemical analysis (method) will provide a constituent 

concentration with a certain level of accuracy and precision over a finite concentration range that 

is specific to that method. Low-nutrient, low-ionic strength waters, such as those commonly 

included in acidic deposition or nutrient enrichment studies, generally require methods that are 

effective at the lowest concentration ranges. A variety of methods are usually available to 

determine the concentration of a given constituent, even at low concentration ranges. The method 

selected must 1) be appropriate for the expected concentration range in the water bodies of 

interest, 2) provide data with the accuracy and precision to successfully achieve project 

objectives, and 3) not exceed logistical limitations with regard to sample collection, sample 

preparation, or laboratory capabilities. For example, a method for determining ammonium 

concentration might provide data over the necessary concentration range with a sufficiently high 

level of accuracy and precision but may require that the analysis be done within 12 hours of 

collection. Such a short holding time might not be feasible for samples collected at remote sites or 

might be beyond the capabilities of the laboratory. 

Often, a variety of laboratory methods and instruments are capable of providing data of suitable 

quality for a particular study. Nevertheless, selection of a method that is capable of meeting the 

required data accuracy and precision for a given project does not ensure that this level of data 

quality will be achieved. Rigorous QA/QC procedures must be followed as part of the method 

implementation. Application of QA/QC procedures will provide the basis for determining the 

quality of the resulting data. In the absence of appropriate QA/QC procedures, it is impossible to 

judge whether the data are of adequate quality to meet the needs of the project. 

In this usage, QC refers to procedures that identify data during the chemical analysis that do not 

meet DQOs, thereby triggering immediate corrective action that usually involves reanalysis of 

that sample. Data quality objectives are generally based on the precision and accuracy levels 

required by the project and the laboratory and the analytical limits of the method used. A key 

component of QC is the analysis of synthetic samples of known concentration, which are 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

72

analyzed along with a specific set of project samples. Quality control procedures are primarily 

focused on instrument performance.

Additional procedures, referred to as QA, are used to document laboratory performance through 

the introduction of artificial and natural samples that are not associated with a specific set of 

project samples, but reflect the accuracy and precision of sample preparation and analysis, 

including instrument performance. Thus, QA and QC procedures are important parts of any field 

and laboratory sampling program.

There are three primary components to QA for the project laboratory:

• Routine evaluation of laboratory analytical performance relative to DQOs.

• Strict adherence to project SOPs including sample bottle preparation, sample collection, 

sample processing, and analysis methods.

• Submission of measurement data QA results along with reported analytical data.

3.2 ATTRIBUTES OF DATA QUALITY

The goal of any field monitoring project is to produce sound analyses and high quality data. 

Establishment of DQOs and development of a QA plan are important to ensure that data meet the 

established objectives for precision, accuracy, representativeness, completeness, and 

comparability. Each type of QA/QC sample or process is generally associated with a DQO. The 

value of the DQO for each analyte is set by project objectives and is usually method-specific. If 

the range of acceptable values measured for a sample of known concentration and defined by a 

DQO is exceeded, the method is considered to be out of control limits and remedial action must 

be taken in the laboratory. The various attributes of data quality and how they are evaluated are 

described below.

3.2.1 Method Detection and Reporting Limits

METHOD DETECTION LIMIT

For chemical measurements, requirements for the method detection limit (MDL) must be 

established. The term “detection limit” has been used in various ways when referring to the lower 

limit of a method concentration range. This lower limit can be a function of instrument capability, 

chemical reactions that are part of the method, or both. The most basic definition of a detection 

limit is the threshold below which measured values are not considered statistically different from 

a blank value (Helsel 2005). Blank values are measurements of samples of DIW (water 

containing no other ions). Thus, measured concentrations below the detection limit are not 

statistically different from zero. 

The repeated measurement of a sample with a known concentration that is at or near the detection 

limit will exhibit considerable variability, which is assumed to be normally distributed. Therefore, 

in this range of measurement, separating a true concentration value from a value resulting from 

analytical noise is problematic. This is of particular consequence for research or monitoring 

objectives that involve the detection of trace contaminants, such as Hg or organic contaminants 

and low levels of nutrients or other analytes in dilute waters.
SECTION 3. QUALITY ASSURANCE/QUALITY CONTROL PROTOCOLS

73

The MDL is defined as the lowest level of analyte that can be distinguished from zero. The first 

step in determining the detection limit is to make a best estimate of the value of the MDL. A set 

of standards is then defined with sequentially decreasing concentrations that extend above and 

below the estimate of the MDL. Each concentration should be analyzed seven times to provide a 

mean and standard deviation for each concentration value (Helsel 2005). 

The true MDL will occur at a concentration that is not statistically different from the next lowest 

concentration. This is determined by running t-tests between the paired concentrations, starting 

with the two highest concentrations. A t-test is run between the second highest concentration and 

the third highest concentration, working downward until a pair of concentrations that are not 

statistically different are reached. For example, it may be found that the fourth and fifth lowest 

concentrations were not statistically different (the method could not detect the difference between 

these two concentrations); the third lowest concentration would then be the true MDL. 

Determination of the method detection limit is demonstrated in the following example.

1. Assume that the best estimate of the MDL for a particular method equals 0.01 μg/L.

2. Make up a set of solutions with the following concentrations that bracket the estimated 

MDL, numbered from highest concentration to lowest concentration:

1. 0.06 μg/L

2. 0.04 μg/L 

3. 0.02 μg/L

4. 0.01 μg/L

5. 0.005 μg/L

6. 0.003 μg/L

3. Analyze each solution seven times. Calculate a mean and standard deviation from the seven

measured values obtained for each solution. The mean values might look like the 

following:

1. 0.055 μg/L

2. 0.039 μg/L 

3. 0.022 μg/L

4. 0.011 μg/L

5. 0.014 μg/L

6. 0.013 μg/L

4. Run t-tests sequentially between each pair of concentrations (1 versus 2, 2 versus 3, 3 

versus 4, etc.). 

5. Suppose that results for solutions 1 versus 2, 2 versus 3, and 3 versus 4 are statistically 

different, but results for solutions 4 versus 5 are not statistically different: this means that 

the concentration of solution 3 (0.02 μg/L) is your MDL.
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A variety of less rigorous methods for determining detection limits are also used. One of the most 

common methods determines the MDL by multiplying the standard deviation of repeated 

measurements of the estimated MDL by a factor of three. The accuracy of the detection limit 

determined in this manner will depend on the accuracy of the estimated MDL, which is unknown. 

Therefore, we recommend the stepwise determination described above.

The MDL can vary from run to run and over time in response to such issues as a change in 

analyst, new instrumentation, or the aging of instrumentation. Therefore, the initial analysis to 

determine the MDL should be repeated three times over several weeks and at least annually

thereafter for constituents with concentrations in water samples that commonly occur near or 

below the MDL. Laboratories that focus on the measurement of trace contaminants may 

determine MDLs more frequently, but otherwise, some unmeasured variation in the MDL won’t 

negatively affect data quality. If a new analyst is appointed or equipment is replaced, a new MDL 

value should be determined regardless of the length of time since the last MDL was determined. 

The MDL is not to be confused with the upper limit of the concentration range of a particular 

method. At concentrations above the method range, the relationship between measurements of 

standards and the known concentrations of these standards can change, thereby requiring a 

different standard curve. 

Measurements that fall below the MDL are considered to be non-detects and are often set to zero 

in the database (Helsel 2005). Establishing an MDL addresses the problem of distinguishing 

between false positives and real values at concentrations near the MDL.

REPORTING LIMIT

The reliability of measurements that are above but near the detection limit is lower than the 

reliability of measurements at higher concentrations. The concentration above which 

measurement variability becomes acceptably low defines the threshold referred to as the 

“reporting limit.” Like MDLs, reporting limits are low concentrations, but they are always higher 

than the MDL, at least by a small amount. At measured concentrations above the reporting limit,

the method is considered reliable and therefore subject to DQOs established for precision and 

accuracy. 

Measurements that fall in the narrow range below the reporting limit but above the MDL may not 

consistently meet the DQOs for reproducibility or accuracy and should be flagged in the database. 

We recommend retaining these values in the database because they indicate low, non-zero 

concentrations and therefore provide information that could be useful. The flag should caution the 

user that the precision and accuracy of these low measured values is uncertain and likely to be 

higher than measurements that fall above the reporting limit. 

Reporting limits are determined for each chemical analysis by establishing the precision and 

accuracy of measurements in the lower portion of the method concentration range. To determine 

the reporting limit for a particular analysis, the steps outlined below should be followed:
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1. Select a relative DQO (± %) for both precision and accuracy. A value of 10% is commonly 

used for most analytes.

2. Make an estimate of the concentration that defines the reporting limit. The values for 

reporting limits listed in Table 3-1 can be used to provide these estimates. From Table 3-1, 

for example, the estimated reporting limit for nitrate analysis is 0.1 mg/L.

3. Create three solutions of known concentrations that are higher than the estimated reporting 

limit, one solution with a concentration equal to the estimated reporting limit, and three

solutions of known concentrations that are lower than the estimated reporting limit but 

higher than the MDL (listed as 0.03 mg/L for nitrate in Table 3-1). For example, the nitrate 

concentrations in the test solutions might be as follows:

1. 0.6 mg/L

2. 0.4 mg/L

3. 0.2 mg/L

4. 0.1 mg/L 

5. 0.08 mg/L

6. 0.06 mg/L

7. 0.04 mg/L

4. Analyze each solution five times and calculate the mean and standard deviation of the five

values at each concentration level.

5. Using the mean values, calculate the accuracy (expressed as % error) and precision 

(expressed as the coefficient of variation or CV) following the procedures given below in 

the next section. Resulting data may look something like this:

1. 0.6 mg/L; % error = 5.4 precision = 6.8

2. 0.4 mg/L; % error = 6.2 precision = 4.9

3. 0.2 mg/L; % error = 7.5 precision = 6.3

4. 0.1 mg/L; % error = 7.2 precision = 7.5

5. 0.08 mg/L; % error = 8.1 precision = 9.5

6. 0.06 mg/L; % error = 21.8 precision = 25.0

7. 0.04 mg/L; % error = 45.6 precision = 39.3

6. Determine the test concentration above which the measurements of error and precision are 

both less than or equal to 10%. For this example, the concentration is 0.06 mg/L. The nexthighest test concentration in the series is then designated as the reporting limit. Based on 

these data and a DQO of 10% for both accuracy and precision, the reporting limit would be 

0.08 mg/L.
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Table 3-1. Recommended DQOs for detection limits, precision, accuracy, and completeness1.

Variable or Measurement

Method 

Detection Limit

DQOs for Precision and Accuracy

Reporting 

Limit Completeness

Relative 

DQOs

(± %)

Absolute DQOs

(± concentration 

value)

Oxygen, dissolved NA NA 0.5 mg/L NA 95%

Temperature NA NA 1 oC NA 95%

pH, closed system and equilibrated NA NA 0.15 pH units none 95%

Acid Neutralizing Capacity NA 15% 6 μeq/L none 95%

Carbon, dissolved inorganic, closed system 0.10 mg/L 10% 0.1 mg/L 0.5 mg/L 95%

Carbon, dissolved organic 0.1 mg/L 10% 0.1 mg/L 0.5 mg/L 95%

Conductance NA 10% 1 μS/cm none 95%

Aluminum, total dissolved, total monomeric, 

and organic monomeric

10 μg/L 10% 0.02 mg/L 27 μg/L 95%

Major Cations:

Calcium

Magnesium

Sodium

Potassium

0.02 mg/L

0.01 mg/L

0.02 mg/L

0.04 mg/L

10%

10%

10%

10%

0.02 mg/L

0.02 mg/L

0.02 mg/L

0.04 mg/L

0.08 mg/L

0.02 mg/L

0.03 mg/L

0.05 mg/L

95%

Ammonium 0.02 mg/L 10% 0.02 mg/L 0.04 mg/L 95%

Major Anions:

Chloride

Nitrate

Sulfate

0.03 mg/L

0.03 mg/L

0.05 mg/L

10%

10%

10%

0.03 mg/L

0.03 mg/L

0.05 mg/L

0.1 mg/L

0.1 mg/L

0.2 mg/L

95%

Silica 0.05 mg/L 10% 0.05 mg/L 0.4 mg/L 95%

Phosphorus, total 1 μg/L 10% 0.002 mg/L 4 μg/L 95%

Nitrogen, total 0.07 mg/L 10% 0.03 mg/L 0.15 mg/L 95%

True Color NA 10% 5 PCU none 95%

Turbidity NA 10% 2 NTU none 95%

Total Suspended Solids 0.1 mg 10% 1 mg/L 0.4 mg/L 95%

1 NA = not applicable. DQOs for precision and accuracy are expressed two ways: in relative terms (± % of measured value) and in absolute 

terms (± actual measured concentration). The DQO is considered to be met if either of these criteria is satisfied.

3.2.2 Precision and Accuracy

Precision and accuracy are estimates of random and systematic error in a measurement process. 

Together they provide an estimate of the total error or uncertainty associated with an individual 

measurement. Precision is measured by repeated analysis of a single sample. The variation of 

these measurements indicates the level of method precision. Accuracy is an indication of how 

closely the measurements match the true concentration of the sample. An illustration of the 

distinction between precision and accuracy is shown in Figure 3-1. 
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Figure 3-1. Schematic illustration of precision and accuracy.

Accuracy can be determined from measurements of solutions of known composition or from the 

analysis of samples that have been fortified by the addition of a known quantity of analyte. 

Accuracy is quantified by relating the measured value of a QC sample to the known value of that 

QC sample. It is usually expressed as a percent error. For QC samples, the DQO objective is 

defined as the value of the percent error. If the measured concentration is greater than the known 

value plus the DQO or lower than the known value minus the DQO, the method is considered to 

be out of the control limits. The percent error is calculated as shown below: 

(1)

Accuracy can also be quantified through analysis of interlaboratory reference samples. The USGS 

Standard Reference Program provides a most-probable value that can be used to calculate a 

percent error in the same manner that known values for QC samples are used. The same approach 

can be used with the Environment Canada’s National Water Research Institute Program (NWRI),

which provides a most probable value, D as follows: 

D = [(AV – MCN)/MCV] x 100 (2)

where AV = analyzed value, and

MCV = mean concentration value (most-probable value for source material)

Method precision is evaluated by analyzing multiple, often duplicate or triplicate, project 

samples. Ideally, each time a sample is reanalyzed, the same concentration value should be 

reproduced. Precision is typically quantified by the CV. The DQO is defined as the CV above 

which the method is out of control. The CV is calculated as:

(3)

Where s is the standard deviation, and is the arithmetic mean of replicate samples. 

It should be noted that relative precision (e.g., CV) is not independent of concentration. For low 

concentrations, criteria for both bias and precision are typically expressed in terms of absolute 

rather than relative error.

3.2.3 Completeness

Completeness requirements are established and evaluated as the percent of valid data obtained 

versus the amount of data expected. Thus, completeness quantifies the extent to which data are 

missing. Completeness objectives are usually designated as over 95% for each variable.
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3.2.4 Comparability

Comparability is defined as the confidence with which one data set can be compared to another. 

Comparability is enhanced by the use of standardized field and laboratory sampling procedures. 

Comparability of data is also facilitated by implementation of standardized QA and QC 

techniques. For all measurements, reporting units and formats are specified in advance and 

recorded on field forms and laboratory databases in these units and formats. Comparability is also 

addressed by providing QA data on detection, precision, and accuracy and by conducting 

methods comparison studies when necessary and participating in interlaboratory performance 

evaluation studies, such as those conducted by the USGS and NWRI. In order to provide 

estimates of trends in any analyte or indicator, data collected each year must be comparable to 

data collected in all prior and subsequent years. Comparability can be quantified through 

comparison of precision and accuracy estimates obtained from QA samples.

3.2.5 Representativeness

Representativeness is the degree to which the data accurately and precisely represent the 

environmental attribute of interest. Although representativeness is not a laboratory QA/QC issue, 

it is affected by problems in all other attributes of QA. A representative sample requires that the 

sample site be reflective of the study population of interest and that the sample itself is 

representative of the system of interest (for example, that the water sample collected in the field 

reflects the condition in the subject lake or stream). Representativeness is ensured by following 

all field and laboratory sampling procedures and holding time requirements to ensure that 

analytical results are representative of the conditions at the time of sampling. Use of QA and QC 

samples similar to the type of environmental samples being analyzed provides estimates of 

precision and bias that are applicable to the collected data.

3.2.6 Recommended Laboratory Data Quality Objectives 

Each laboratory must also have its own set of DQOs that pertain to the quality of the analytical 

data produced by the laboratory. Projects also have data quality requirements that are based on 

the objectives and resources of the project. Therefore, the laboratory DQOs must be evaluated to 

ensure that the laboratory is capable of delivering the accuracy and precision that the project 

requires. In general, we recommend DQOs for detection, accuracy, and precision as specified in 

Table 3-1. These DQO values are used by the USGS New York Water Science Center Water and 

Soil Analysis Laboratory, in Troy, NY, which specializes in the analysis of low-ionic strength 

waters for air pollution effects research projects. 

These recommended guidelines for precision and accuracy DQOs given in Table 3-1 may not be 

appropriate for all projects. Forest Service staff might determine, for a specific analyte and 

project, that one or more recommended guideline(s) can be relaxed, especially if the laboratory is 

unable to achieve the recommended level of data quality and if the project does not require such 

high levels of precision and accuracy. Conversely, FS staff might determine that a particular 

project requires higher standards of precision and accuracy. In general, the values presented in 

Table 3-1 should satisfy the needs of most anticipated FS ARM program water quality sampling 

projects. 
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As represented in Table 3-1, we recommend application of DQOs for precision and accuracy that 

are calculated two ways: based on relative percent variation and based on absolute variation. A 

given DQO can be considered to be met if either of these two conditions is satisfied. In general, 

conformance with the DQO for accuracy and precision will be determined by evaluation of 

relative variation. However, at low concentration values, the relative DQOs can be difficult or 

impossible to achieve. For example, if the ANC of a particular stream is 10 μeq/L, the relative 

DQO for precision and accuracy of the ANC measurement is 15% (Table 3-1), or 1.5 μeq/L. 

There is no laboratory that can achieve that level of accuracy and precision in measuring ANC. 

For a sample having such low ANC, however, the absolute DQO (6 μeq/L, Table 3-1) is 

considered to be achievable. As long as the absolute DQO criterion is satisfied, the DQO for 

precision and accuracy is considered to be met.

For most analytes, our recommended relative DQO for precision and accuracy is 10%. 

Nevertheless, most laboratories should be able to do better than that. A good target DQO in most 

cases is ±5%; this is the level of precision and accuracy that the laboratories and projects should 

strive for.

3.3 QA/QC SAMPLE TYPES

The following sections describe the various types of samples and DQOs that are typically used 

for QC and QA in laboratories that specialize in analysis of low-nutrient, low-ionic strength 

waters. There is no definitive rule regarding how many QA/QC samples should be included in a 

given project. This will be determined, in part, by the intended use of the data and the available 

budget. In general, we recommend that at least 30% of the samples analyzed in the laboratory for 

a given project be QA or QC samples, distributed among the types of samples discussed in the 

sections that follow. 

Quality-control samples are used to measure the accuracy of an instrument’s calibration and to 

detect variations in instrument response within an analytical run. Types of laboratory QC samples 

are summarized in Table 3-2. These samples are made up in the laboratory using Type I DIW and 

purchased chemicals. Source material for all QC samples is either obtained from a manufacturer 

other than the producer of the source material used to make calibration standards, or is obtained 

from a lot other than the source material used to make calibration standards. 

Quality control-high and QC-low samples are analyzed within a given laboratory run for most 

constituents. Exceptions are ANC, pH, and specific conductance. Either the QC-high sample or 

QC-low sample is analyzed within an ANC, pH, and specific conductance run, depending upon 

the expected concentration range of the environmental samples. This reduces the chance of 

carryover from a low pH (or low ANC or specific conductance) QC sample to a high pH project 

sample through the transfer of the electrode between samples. 

We recommend that QC samples be analyzed immediately after instrument calibration, once after 

every 10 project samples, and at the end of each run. QC samples that do not meet DQOs for 

accuracy are rerun. If the value is then acceptable, the run is continued. If the rerun QC sample 

value is unacceptable, the project sample data preceding it are considered to be out-of-control: the 

data are rejected, and the instrument is recalibrated. Only accepted QC-sample and project sample 
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data are entered into the database. The analytical results of QC samples should be recorded to 

indicate the frequency of out-of-control data that are not rerun and biases and trends of control 

data.

Table 3-2. Recommended laboratory quality control samples.

QC Sample Type (Analyte), and 

Description Frequency

Acceptance

Criteria Corrective Action

Laboratory Blank: (all analyses except 

pH and total suspended solids [TSS])

Reagent Blank: (DOC, Al [total, 

monomeric, and organic monomeric], 

ANC, NH4

+, SiO2)

Once per batch 

before sample 

analysis

Control limits < ±MDL 

or < 1 μM, whichever 

is least restrictive

Prepare and analyze new blank. Determine and correct problem (e.g., 

reagent contamination, instrument calibration, or contamination 

introduced during filtration) before proceeding with any sample 

analyses. Reestablish statistical control by analyzing three blank 

samples.

Filtration Blank: (all dissolved analytes, 

excluding syringe samples)

ASTM Type II reagent water processed 

through filtration unit

Prepare once per 

week and. archive

Measured 

concentrations < 

MDL

Measure archived samples if review of other laboratory blank 

information suggests source of contamination is sample processing.

Detection Limit Quality Control Check 

Sample (QCCS): (all analyses except 

true color, turbidity, and TSS), prepared 

so concentration is approximately four to 

six times the required MDL 

Once per batch Control limits < ±MDL Confirm achieved MDL by repeated analysis of appropriate standard 

solution. Evaluate affected samples for possible re-analysis.

Calibration quality control check sample 

(CQCCS)1

Before and after 

sample analyses

Control limits < 

precision objective: 

Mean value < bias 

objective

Repeat CQCCS analysis.

Recalibrate and analyze CQCCS.

Reanalyze all routine samples (including performance evaluation and 

field replicate samples) analyzed since the last acceptable CQCCS 

measurement. 

Internal Reference Sample: (suggested 

when available for a particular analyte)

One analysis in a 

minimum of five 

separate batches

Control limits < 

precision objective.

Mean value < bias 

objective

Analyze standard in next batch to confirm suspected imprecision or 

bias.

Evaluate calibration and CQCCS solutions and standards for 

contamination and preparation error. Correct before any further 

analyses of routine samples are conducted. Re-establish control by 

three successive reference standard measurements that are 

acceptable. Qualify all sample batches analyzed since the last 

acceptable reference standard measurement for possible reanalysis.

Laboratory Replicate Sample: (all 

analyses)

For closed headspace syringe samples, 

a replicate sample represents a second 

injection of sample from the sealed 

syringe.

One per batch Control limits < 

precision objective

If results are below MDL:

Prepare and analyze split from different sample (volume permitting).

Review precision of CQCCS measurements for batch.

Check preparation of split sample.

Qualify all samples in batch for possible reanalysis.

Matrix spike samples: (only prepared 

when samples with potential for matrix 

interferences are encountered)

One per batch Control limits for 

recovery cannot 

exceed 100±20%

Select two additional samples and prepare fortified subsamples.

Reanalyze all suspected samples in batch by the method of standard 

additions. Prepare three subsamples (unfortified, fortified with solution 

approximately equal to the endogenous concentration, and fortified with 

solution approximately twice the endogenous concentration).

1 For turbidity, a CQCCS is prepared at one level for routine analyses (U.S. EPA 1987). Additional CQCCSs are prepared as needed for 

samples having estimated turbidities greater than 20 nephelometric tubidity units (NTU). For total suspended solids determinations, CQCCS 

is a standard weight having mass representative of samples.

3.3.1 Filter Blanks, Analytical Blanks, and Field Blanks

Blanks are aliquots of DIW that are processed and analyzed in the same manner as project 

samples. Filter blanks are analyzed only for constituents that require filtration. Filter-blank 

analysis indicates whether detectable contamination has occurred during any step in sample 

handling that occurs in the laboratory, including bottle-washing procedures, filtration, sample 
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preservation, and chemical analysis. Analytical blanks are aliquots of Type I DIW (ASTM 1984) 

that are processed and analyzed as project samples, except that the filtration step is omitted. 

Contamination of analytical blanks may be attributed to any step in sample-handling other than 

filtration, including the quality of DIW. The use of an analytical blank together with a filter blank 

therefore enables contamination from filtration to be isolated from contamination during DIW

preparation or other phases of sample preparation and analysis. The use of both a filter blank and 

an analytical blank is recommended because the filtration process poses the greatest single source 

of potential sample contamination. A filter blank and an analytical blank should be included as a 

QC pair in the sample stream at a frequency of at least 1 per 50 project samples. 

Some programs require a QA sample referred to as a field blank. The field blank is prepared by 

bringing DIW into the field and transporting it back to the analytical laboratory, with or without 

transferring it to a sample bottle. From that point forward, the DIW in the sample bottle is treated 

as any other sample collected in the field. It is not clear what information this procedure provides 

because the action involved does not replicate any aspect of the actual field sampling. We 

therefore do not recommend the collection of field blanks unless some specific project objective 

requires field filtration of water samples, which could introduce the potential for sample 

contamination. 

3.3.2 Replicate Environmental Samples 

An environmental replicate set generally consists of either two duplicate samples or three 

triplicate samples. The replicated samples are collected at the same field site, following the same 

collection procedure and as close as possible to the same time, as the original sample. The 

purpose of replicate samples is to document sampling and analytical precision using samples that 

reflect the chemistry of actual project samples. The results of analysis of sample replicates 

provide useful information regarding the overall ability of the field and laboratory program to 

quantify the constituents of interest. Differences in measured values between or among replicates 

reflects fine-scale temporal and spatial variability in water quality at the sample site location plus 

any variability or error introduced in the sample collection, sample processing, and/or laboratory 

analysis procedures. Ideally, replicates are collected and analyzed as part of the sampling 

protocols of every project. For some programs, replicates are collected (as back-up) from every 

site, but only a subset of those replicates are analyzed. Environmental samples provide a better 

test of precision than artificial samples because they include natural constituents that could alter 

the reproducibility of a given laboratory method. Precision can also be affected by bottle washing, 

sample-collection, sample-processing procedures, and analysis. 

In long-term monitoring studies, project sites should be selected for replicate collection on a 

rotating basis to evaluate precision within the full variability of project samples being analyzed. 

For the analysis, the laboratory should alternate between analyzing a replicate set consecutively 

(within the same analytical run) and separating the replicate samples in analytical runs that occur 

on different days or at the beginning and end of the analytical run. One set of replicate project 

samples should be included in the sample stream at a frequency of at least 1 per 50 project 

samples. More commonly, this frequency should be 1 per 20 project samples if the budget allows. 
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3.3.3 Spiked Project Samples 

Surface water samples tend to contain a wide variety of chemical constituents with concentrations 

that can be highly variable. As a result, there is the potential for one constituent to interfere with 

the analysis of another constituent. For example, a sample with a high concentration of DOC

(which imparts a brown color to the water) would interfere with some analyses that rely on 

colorimetric measurement to determine concentrations. Well-documented methods specify which 

constituents may interfere with a given analysis and at what concentration range. However, these 

specifications should be verified for the samples within a specific project and its testing 

laboratory to ensure the accuracy of the measurements. If sample concentrations are being 

measured in a range that could cause interference with the measurement of another constituent, 

the sample should be run twice: once untreated and once after being spiked with a known amount 

of the constituent of concern. The measured value of the samples, including the spike, should fall 

within the range of the method. If the concentration of the spiked sample equals the concentration 

of the unspiked sample plus the added amount, then recovery is complete and it can be assumed 

that there is no interference. To express this relationship in terms of percent recovery, the 

equation below can be used:

(4)

The full range of an interfering constituent may not be known in the early stages of project 

sampling. Therefore, including spiked project samples in the sample stream is recommended until 

it is determined that interference is not a concern. Also, when a laboratory is starting to use a new 

method, the inclusion of spiked project samples is recommended to verify method specifications. 

Forest Service staff should ask the laboratory to identify requested analyses that might be subject 

to interference with the types of samples that will be analyzed, intended methods, and analytes. It 

may not be possible, however, for laboratory staff to make that determination until after a given 

water body has been sampled and analyzed. If interferences are probable, then FS staff should 

request analyses of spiked samples to determine the extent of interference.

3.3.4 External Quality Assurance Samples 

A variety of programs exist that provide reference samples for laboratory analysis. These 

programs develop bulk samples that can be subsampled and sent to participating laboratories on a 

set frequency. Usually these samples comprise some type of environmental sample that is 

chemically similar to the samples that a laboratory typically analyzes. Results from all the 

participating laboratories are compiled, statistically summarized, and provided to the laboratories 

(usually without specific identification). Participation in a reference sample program provides the 

opportunity for a laboratory to compare their performance with other laboratories. Program 

participation should be included in the QA/QC program of any laboratory that analyzes 

environmental samples. 

In particular, laboratories that analyze low-nutrient, low-ionic strength water samples should 

participate in reference programs that provide these types of samples. The USGS Standard 

Reference Sample (SRS) Project conducts a national interlaboratory analytical evaluation 
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program semi-annually. The program includes three types of samples: low-ionic strength, 

nutrient, and trace constituents. Typically, the reference samples consist of snow, rain, surface 

water, or DIW that is collected, filtered, and possibly spiked with reagent-grade chemicals to 

meet the goals of the program. Reference samples for low-ionic strength constituents are analyzed 

for calcium, chloride, magnesium, pH, potassium, sodium, specific conductance, and sulfate. 

Reference samples for nutrient constituents are analyzed for ammonium and nitrate. Reference 

samples for trace constituents are analyzed for aluminum, calcium, magnesium, potassium, 

silicon, and sodium. Laboratory personnel are aware of the presence of the SRS sample at the 

time of analysis but do not know the constituent concentrations until results are posted on the 

SRS Project website after the conclusion of each study. The most probable value (MPV) for each 

constituent is equal to the median value calculated from the results submitted by participating 

laboratories. Laboratory results are compared with the MPV for each constituent, and a percent 

difference is calculated and reported.

A second standard reference program is operated by Environment Canada’s NWRI. This program 

sends a set of 10 samples to a group of participating laboratories twice a year. The samples are 

obtained from predominantly low-ionic strength waters representing several sources, such as 

precipitation, snow, lake water, and stream water throughout North America. The concentrations 

of the constituents in the NWRI samples are similar to those of the environmental samples 

analyzed by laboratories that specialize in low-nutrient, low-ionic strength samples. Laboratory 

results are compared with a median concentration value (MCV) calculated from results from all 

participants in the NWRI program. (The USGS MPV and NWRI MCV are the same statistic, 

although named differently.) Laboratory personnel are aware of the presence of NWRI samples at 

the time of analysis but do not know the MCV of the constituents until Environment Canada 

publishes a report at the conclusion of each study. 

A drawback to standard reference sample programs is that the analyst knows that this is a “high 

priority” sample and therefore may give extra attention to its analysis. Therefore the results might 

not fully reflect those obtained in the analysis of routine project samples. This type of analyst bias 

can be avoided with blind reference samples. 

Blind reference samples are processed and analyzed as environmental samples and therefore 

appear to the analyst to be project samples. Ideally, these samples would originate from an 

interlaboratory reference program so that known concentration values would have been or would 

be established for the sample. Implementation of a blind reference sample program requires the 

participation of one person who works in the laboratory. The reference samples must be coded 

and prepared by this person so that they cannot be distinguished from routine samples by the 

analyst. This person is also responsible for retrieving the analysis data from the laboratory 

database and recoding it as QA data. One blind reference sample per 50 project samples is 

recommended. 

When evaluating candidate laboratories, their participation and performance in an interlaboratory 

reference program is a very useful decision criterion. We recommend that, where practical, the FS 

avoid using laboratories that do not participate in such a program. FS staff should request and 

review performance results before making arrangements to use a particular laboratory.
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3.4 FIELD QUALITY ASSURANCE

3.4.1 Sample Containers

The required sample containers and cleaning procedures are described in detail in the laboratory 

protocols. At least 2% of the cleaned containers (randomly selected) must be given a specific 

conductance check, which entails measuring the conductance of DIW in the sample container 

after the 48-hour soak period. Conductance should be lower than 1.2 μS/cm. If the conductance is 

greater than 1.2 μS/cm, re-rinse all the containers cleaned since the last acceptable check. If 

contamination is found, then 25% of the sample containers in subsequent batches should be 

monitored until all monitored containers in a batch pass the conductance test.

3.4.2 Field Measurements

Measurements of dissolved oxygen, temperature, conductance, and pH are often made in the 

field. If these measurements are made in the field, they require field QA procedures and the use 

of both performance evaluation (PE) and QC samples as described in Table 3-3. These samples 

confirm that the measuring devices (often field meters) are functioning properly and are within 

control limits over the entire length of the study.

Table 3-3. Field quality control samples. (Source: Paulsen 1997.)

Measurement

QC Sample 

Type Description Frequency Acceptance Criteria Corrective Action

Dissolved 

Oxygen

PE Sample1 Concurrent determination 

of sample by Winkler 

titration

Once per meter 

per field season

Measured O2 within ±1 

mg/L of O2 estimated by 

Winkler titration

Replace meter and/or probe

QC Check 

Sample

Water-saturated air Daily (at base 

station)

Instrument can be 

calibrated to theoretical 

value

Replace meter and/or probe

Temperature PE Sample Concurrent measurement 

of 0 °C and 25 °C 

solutions with NISTtraceable thermometer

Once per meter Within ±1 °C of 

thermometer reading

Replace probe and/or meter

QC Check 

Sample

Concurrent measurement 

of sample with field 

thermometer

Weekly Within ±1 °C of 

thermometer reading

Replace probe and/or meter

Conductance QC Check 

Sample

Solution of known 

conductance

Weekly Within 10 μS/cm of 

theoretical value

Re-calibrate meter using 

NIST-traceable standards; 

replace probe and/or meter

pH QC Check 

Sample

Solution of known pH Daily Within 0.1 pH unit of 

theoretical value

Re-calibrate meter and 

probe or replace probe

1 PE is performance evaluation sample.

Peck and Metcalf (1991) developed a stable and well-quantified (both theoretically and 

analytically) QC check sample for conductance, pH, and ANC measurements in dilute surface 

waters. It is a 1:200 dilution of the National Institute of Standards and Technology (NIST) 0.025 

mol/kg KH2PO4 and 0.025 mol/kg Na2HPO4 standard pH stock solution. It has a pH of 6.89, a 

conductance of 37.6 μS/cm, and an ANC of 125 μeq/L. This solution is recommended as a QC 
SECTION 3. QUALITY ASSURANCE/QUALITY CONTROL PROTOCOLS

85

check for studies doing field pH and/or conductance measurements in relatively well-buffered 

waters.

Unpublished data from EPA’s National Surface Water Survey showed that pH can be measured 

more precisely in the laboratory using water samples that have been collected in sealed 60 mL 

syringes with no headspace than in the field using portable pH meters. In general, measurements 

made under controlled laboratory conditions are more precise and accurate than those made in the 

field where contamination, weather, and fatigue can induce variability. Thus, we recommend that 

pH and conductance measurements be made in the laboratory. 

Analytes that are sensitive to changes in CO2 concentrations (e.g. pH, DIC) should ideally be 

measured in samples collected in the field into glass bottles having septum caps or into syringes 

with no air bubbles and analyzed within 72 hours of collection if the sample CO2 concentration is 

likely to be supersaturated with respect to the atmospheric CO2 concentration. Stream samples 

affected by discharging ground water (springs) and lake samples from the hypolimnion of 

stratified lakes are especially likely to be supersaturated with CO2. Typically, you need one 

syringe for pH and DIC. Collecting an extra syringe is recommended in case additional sample 

volume is needed in the laboratory. 

Temperature measurements must be made in the field. Dissolved oxygen measurements are also 

made in the field using a meter.

3.5 REPORTING QUALITY ASSURANCE DATA

Before selecting a laboratory, their QA results should be evaluated to ensure that the data quality 

delivered by that laboratory will be suitable for the planned project. The most common and 

perhaps most effective method of reporting QA data is through the use of control charts, which 

plot QC or QA data through time. The control charts 1) indicate whether the laboratory DQOs are 

met for individual QC samples, 2) reveal long-term biases or trends within and outside the control 

limits, and 3) provide comparisons with results from other laboratories. Each constituent has 

prescribed control limits that are set by the laboratory (Table 3-1). Ideally, when no bias is 

present, half the data points in a control chart would be above and half below the target value line. 

Although QC samples are used to evaluate data quality and identify samples that need to be rerun 

during the analysis, when plotted on control charts, QC samples also provide useful data to 

evaluate method performance over time, thereby also providing QA information. 

Results from the analysis of QC samples are plotted on control charts in which the central line is 

equal to the target value (known concentration) of the control sample (Figure 3-2). Both a high 

and a low concentration QC sample, relative to the expected concentration distribution of the

project samples, should be analyzed. If the QC sample is a blank, the target value is set to zero. A 

constituent analysis is considered biased if 70% or more of the data points on a chart are either 

above or below the target value line. The upper and lower control-limit lines on each chart 

represent the range of satisfactory data based on the DQOs. The QC-high and QC-low samples 

are plotted on separate graphs by constituent and date of analysis, and the control charts are 

evaluated for trends and/or bias and precision. 
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Figure 3-2 provides 3 years of data for a QC sample used for low concentration measurements of 

nitrate. Virtually all of the data fall within the control limits without any indication of trends or 

bias. 

Figure 3-2. Results from analysis of low-concentration QC samples for nitrate analysis. The target 

value of the control sample is represented by the central line; the upper and lower dotted 

and dashed lines represent the range of satisfactory data based on the DQO.

Results of the QC sample analysis shown in Figure 3-3 also indicate a reliable method; only one 

value fell outside the control range. However, within the control range, an upward trend occurred 

in 2006, followed by a downward trend in 2007. If either of these trends had continued, the data 

would have drifted out of control limits. To ensure early detection of trends in QC, control charts 

should be updated daily to monthly depending on the sample load.

For the analysis of filter blanks and analytical blanks, the control range is defined by zero and the 

DQO threshold. For replicate sample concentrations, the CV of the two or three replicate samples 

is plotted and the control limits are determined by ± the DQO for accuracy and precision. Control 

charts can be used to show results from interlaboratory comparisons by plotting the percent 

difference from the most probable value. Control limits are defined by the acceptable percent 

difference from the most probable value, which might be designated to be 10%, for example, 

unless the concentration of the test solution is low, in which case a higher value should be 

selected. 

Documentation of QA data should be readily available to projects that use or contemplate using a 

specific laboratory. Forest Service staff should request, from the laboratory, in advance of starting 

a project, QA results that would be applicable to the planned project. These results should 

include, at a minimum, QC results plotted on control charts and comparisons with results from 

other laboratories.
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87

Figure 3-3. Low-concentration QC samples for sulfate analysis. The target value of the control 

sample is represented by the central line; the upper and lower dotted and dashed lines 

represent the range of satisfactory data based on the DQO.

3.6 LABORATORY AUDITS AND CERTIFICATION

All laboratories conducting chemical analyses should be periodically audited by a qualified 

external body on a set frequency that does not exceed 3 years. These audits should be 

comprehensive in covering every aspect of laboratory activities. Documentation of audit results, 

recommendations, and actions taken by the laboratory should be maintained and available to 

projects that use the laboratory. Ideally, audits are conducted as a component of a certification 

program. In addition, round-robin programs, in which multiple labs analyze the same set of 

performance evaluation samples and compare results, are an excellent way to evaluate the 

performance of an individual laboratory and to ensure that it provides high-quality data. If a 

laboratory is not able to document that it provides high-quality data, then the FS may wish to find 

an alternate laboratory for analyzing samples considered to be important from a regulatory or 

decision-making perspective. Any laboratory chosen for important project work should be able to 

provide documentation of audits within the last 2 years of their laboratory procedures. 

There is currently one national accreditation program for laboratories that analyze environmental 

samples. This program is administered through the National Environmental Laboratory 

Accreditation Conference (NELAC), formed on the recommendation of the EPA. A cooperative 

association of state and federal agencies, NELAC was formed to establish and promote mutually 

acceptable performance standards for the operation of environmental laboratories. The standards 

cover both analytical testing of environmental samples and the laboratory accreditation process. 

To accomplish these goals, NELAC developed the National Environmental Laboratory 

Accreditation Program (NELAP). This program recognizes state programs as accrediting 

authorities that administer the program. For example, a laboratory headquartered in New York 

State would apply to New York State’s Environmental Laboratory Accreditation Program 
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88

(ELAP). As each laboratory becomes accredited under a NELAP-recognized accrediting 

authority, the laboratory and its accredited scope of testing will be entered into a national 

database. One of the fundamental principles of NELAC is that of reciprocity among NELAP 

accrediting authorities. For example, once a laboratory is accredited in one state for testing under 

a specific EPA program, it can be accredited in another state for that EPA program without 

having to meet additional accreditation requirements. We recommend that the FS use accredited 

laboratories or, at a minimum, laboratories that can demonstrate their ability to produce highquality data, as described above. 

3.7 DATA ENTRY

Field crews will be required to enter the field data directly into a Web-based form for inclusion of 

the data into NRM Air. Information such as SampleID, Location, Date/Time, and other field data 

will be entered into the form. When laboratory analyses are complete, lab results will be merged 

with the field data and brought into the database. National Resource Manager Air will support 

two import formats for lab results: 1) an Access database in a STORET-compatible format and 2) 

an Excel template that can be output by an NRM Air tool. Data that pass the QC checks are 

electronically transferred into the laboratory database, where they can be reviewed for errors that 

could result from mislabeling, data entry mistakes, misidentification of samples, contamination, 

or a number of other potential sources of error. Once the data have been verified, they should be 

placed in a location accessible to project personnel for downloading. All types of data storage 

should be backed up on a daily basis.

3.8 SUMMARY

As discussed in the preceding sections, there are many types of QA/QC data and a variety of 

ways to evaluate the reliability of the data collected for a particular project. In the absence of such 

QA/QC analyses, it is impossible to determine whether or not the collected data can meet project 

objectives. Key elements of the QA/QC program that should be used in evaluating a laboratory 

for possible use in analyzing samples from dilute lakes or streams are summarized as follows:

1. Develop project DQOs. These are project specific goals for data quality for each measured 

variable. Table 3-1 is a good starting point but will not be optimal for all projects. Add or 

subtract any variables that are not pertinent to the project at hand. Revise criteria based on 

specific project needs. 

1a. Detection Limits. The detection limit is the threshold below which measured values are 

not considered different from zero concentration. Detection limits must be evaluated in 

relation to levels of ecological concern. If the detection limit is near or above the level 

of concern, then the usefulness of the data might be limited.

1b. Precision and Accuracy. Precision is measured by repeated analysis of the same sample 

to determine the variability in the analytical data for each variable. Accuracy is 

measured by blind analysis of samples with known concentration. The deviation of 

analytical measurements from the true known concentration is called “bias.” The need 
SECTION 3. QUALITY ASSURANCE/QUALITY CONTROL PROTOCOLS

89

for precision and accuracy for a project is dependent on the magnitude of the effect 

being studied: if you are trying to quantify small differences among groups or small 

changes over time, then you will need higher precision and accuracy of measurement.

1c. Comparability. Data need to be comparable to what is being measured at other 

locations and times. To help ensure comparability, the analytical laboratory should be 

certified and/or participate in sample “round-robin” programs in which the same 

samples are analyzed by multiple laboratories and results compared. 

1d. Completeness. Ideally, all sites intended for sampling and all measurements intended at 

each site will be made, constituting 100% completeness. In reality, completeness is 

typically somewhat less than 100%. Minimum completeness goals should be set 

depending on how much of a problem missing data will be in the final analysis of the 

data.

2. Evaluate laboratory QA data to see how well they meet your DQOs.

2a. Evaluate analytical results for blank samples. 

2b. Determine results for sample replicates. 

2c. Quantify expected versus observed results for spiked samples. 

2d. Examine laboratory performance for external QA audit samples. 

3. Evaluate field sampling QA procedures and data.

3a. Sample bottles may be provided already cleaned by the laboratory or they may need to 

be cleaned and tested as described in the Field Protocol.

3b. We recommend that water sample filtration and pH measurements be made in the 

laboratory, where precision is higher and risk of contamination lower, rather than in the 

field. If field filtration is done, field blank samples can indicate potential sample 

contamination. Water for pH measurements in the lab should preferably be collected in 

60 mL syringes, avoiding the introduction of any air bubbles, and then sealed in the 

syringes with a stopcock. 

3c. Evaluate results of field QC check sample analyses for any field measurements as 

described in Table 3-3.

SECTION 4. DATA ANALYSIS PROTOCOLS

91

SECTION 4. 

DATA ANALYSIS PROTOCOLS T.J. Sullivan, A.T. Herlihy, G.B. Lawrence, and J.R. Webb 

4.1 BACKGROUND AND OBJECTIVES

A data analysis protocol (DAP) provides the basis for translating water quality data generated in 

the analytical laboratory into meaningful guidance for data analysis and interpretation. It connects 

the raw data to the information goals for which the data were collected. There are a number of 

information goals that are relevant to water sampling efforts within the FS ARM program, as 

described in Section 1 of this protocol. These goals stem from specific questions that are 

formulated to inform management decision making. 

The purpose of this DAP is to describe graphical, statistical, and other approaches to be used in 

validating, presenting, analyzing, and understanding water quality data. The DAP can support 

analytical efforts by FS personnel who do not have advanced training in chemistry and statistics. 

However, we recommend that, whenever possible, data analysis should be conducted by staff 

members who have a good grounding in water chemistry and a basic understanding of statistics. 

In particular, staff involved in trends analysis should have had formal training in statistics and/or 

consult with a trained statistician while conducting the trends analysis. 

The DAP is divided into sections, as follows:

1. Develop a statement of the objectives of the data analysis.

2. Evaluate and assure the quality of the dataset.

3. Prepare raw data for graphical and statistical analysis.

4. Conduct exploratory analyses.

5. Conduct, if needed, formal statistical analyses.

6. Report data in standardized formats.

As described in the Surface Water Sampling Protocol, collection of surface water chemistry data 

should always have a purpose. Specific questions need to be formulated, and the nature of these 

questions will guide the design of the study, including what, where, when, and how to sample. To 

some degree, these questions will also help guide data analysis.

Some example approaches for data analysis that we recommend for FS staff consideration are 

outlined in Table 4-1. Each example data analysis approach given in the table is tied to a specific 

purpose.
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Table 4-1. Some example approaches for FS ARM program data analysis, tied to the purpose 

and general approach of the field study.

Purpose General Approach Example Data Analysis1

1. Determine whether lake or 

stream is N-limited

A. Measure N, P, and chlorophyll a during 

snow-free season

a. Plot molar N:P ratio over time relative to published ratios that have 

been shown to be associated with N-limitation. Compare to changes 

in chlorophyll a concentrations.

2. Quantify episodic 

excursions from base flow 

chemistry

A. Measure water chemistry during 

rainstorms and/or snowmelt

a. Plot changes during episodes in discharge and selected water 

chemistry parameters to reveal episodic changes in ANC, pH, and 

Ali, and to illustrate changes in potential drivers of those parameters.

b. Plot changes over time during spring and summer in discharge and 

selected water chemistry parameters.

c. Plot changes over time (including episodes) in the ratio of NO3

-

concentration to [NO3

- + SO42-

] concentration versus ANC to 

illustrate the relative importance of NO3

- as a mineral acid anion in 

driving episodic ANC changes.

3. Determine spatial patterns 

in water chemistry across 

a forest or wilderness

A. Conduct regional survey of water 

chemistry

a. Map various chemical concentrations across the landscape to reveal 

spatial patterns. Correlate those to landscape features (geology, 

elevation, etc.).

b. Examine patterns in the data across spatial gradients. 

c. Construct histograms to identify regional (or forest-level) outliers in 

concentrations of key variables. 

4. Quantify long-term 

changes over time in 

water chemistry

A. Sample over an extended period of time 

(at least 8 to 10 years), preferably 

monthly or seasonally during open-water 

seasons

a. Standardize sampling to account for episodic changes or otherwise 

address variability.

b. Conduct trends analyses.

c. Compare trends in biologically relevant variables (ANC, pH, Ali) with 

trends in potential drivers and buffers (SO4

2-

, NO3

-

, SBC, DOC). 

5. Determine extent to which 

air pollution is affecting 

water resources

A. Characterize index chemistry for multiple 

lakes or streams expected to be sensitive

a. Plot central tendency and variability (i.e., box and whisker plots) in 

key variables across sites. 

b. Compare with common benchmark thresholds for ANC (0, 20, 50, 

100 μeq/L), pH (5.0, 5.5, 6.0), and Ali (2, 7 μM) and calculate the 

percent of sites that exceed thresholds. 

B. Conduct survey of waters in study area a. Plot central tendency and variability in key parameters.

b. Compare to common benchmark thresholds. Map chemical 

concentrations. 

C. Use dynamic model to hindcast past 

changes in water chemistry since about 

1900 or earlier

a. Map simulated changes in chemical concentrations.

b. Compare simulated changes with changes derived by other means 

(i.e., long-term monitoring, paleolimnology, space-for-time 

substitution). 

c. Plot simulated changes in ANC versus aspects of current chemistry 

(i.e., ANC, ANC/SO4

2-

). 

d. Estimate proportional changes in ANC versus SO4

2-

, SBC versus

SO4

2-

, ANC versus (SO4

2- + NO3-

), etc. to reveal interactions among 

variables. 

D. Collect and analyze diatom remains in 

sediment core(s)

a. Analyze as described above for dynamic model hindcasts a through 

c. 

E. Simulate critical load of S and/or N 

deposition

a. Map critical loads and exceedances (amount by which ambient 

deposition exceeds critical load). 

b. Plot critical load as function of water chemistry (i.e., ANC, 

ANC/SO4

2-

, etc.).

c. Develop procedures with which to extrapolate critical load on basis 

of water chemistry and/or landscape characteristics. 

6. Evaluate possible need for 

mitigation

A. Follow approaches similar to 5 a. Follow approaches similar to 5. 

1 The following abbreviations are used here: nitrogen (N); phosphorus (P), acid neutralizing capacity (ANC), inorganic monomeric aluminum 

(Ali), nitrate (NO3

-

), sulfate (SO4

2-

), sum of base cations (SBC), dissolved organic carbon (DOC).
SECTION 4. DATA ANALYSIS PROTOCOLS

93

4.2 EVALUATION OF DATA QUALITY 

The first step in analyzing any raw dataset provided by an analytical laboratory should always be 

to conduct an evaluation of the quality of the data. This should include reviewing the QA/QC data 

provided by the laboratory, determining if DQOs have been met by the laboratory, and 

conducting or reviewing the results of data validation. 

There are a variety of protocols that can be used to evaluate and confirm overall analytical data 

quality. These include comparing measured with calculated variables when both are intended to 

represent the same parameter. If measured and calculated values are similar (within an expected 

range of error), then there is an increased likelihood that the data used in the comparison are of 

high quality. If measured and calculated values of the same parameter differ by more than the 

expected variability, then it can be inferred that one or more of the values in the calculations 

represented in that comparison may be in error. This approach can be helpful in flagging certain 

samples or measurements for re-examination to determine if there were recording errors or some 

other kind of error that might be identified and corrected. 

Data validation protocols also include constructing plots of variables that might be expected to 

correlate with each other. Any sample that deviates substantially from the expected relationship 

might be further examined for potential error or flagged in the dataset. 

Sample contamination, analytical error, and/or reporting error can lead to incorrect data values 

that are not representative of conditions in the field. Many such errors can be identified and, in 

some cases, corrected through data validation. Data validation is the process of checking for 

internal consistency among data values using the ionic relationships among the analytes in the 

dataset. If data validation is done in a timely manner, problematic values can sometimes be 

reanalyzed. If nothing else, values that fail validation checks can be flagged in the dataset and 

considered potentially suspect in various data analyses. Sample analysis results from the 

laboratory are considered preliminary until the internal consistency checks described below are 

performed.

4.2.1 Charge Balance

The sum of positively charged ions (cations) in water must equal the sum of those with negative 

charge (anions). Major discrepancies between the sum of measured anions and cations thus reflect 

analytical errors, failure to measure all ions with significant concentrations, or a combination of 

both. Although charge balance calculation and comparison alone cannot necessarily identify the 

cause of a charge imbalance, they can serve as a QC check on the completeness and accuracy of 

the ion chemistry data. A high-quality dataset will show reasonable agreement between the 

calculated cation and anion sums after accounting for the failure to measure all ions in solution. 

To assess the quality of the data for the ionic species in water, ion charge balances involving a 

comparison of the sum of cations to the sum of anions should be calculated for all water samples. 

If the data are provided in mass units (e.g., mg/L), then they must be first converted into 

equivalence units (i.e., microequivalents per liter; μeq/L) by multiplying the concentration in 

mg/L by the appropriate factor in Table 4-2. Anion and cation sums in units of μeq/L are then 

approximated as defined in Equations 1 and 2 below:
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Sum of cations = Ca2+ + Mg2+ + Na+ + K+ + NH4

+ + H+ (1)

Sum of anions = SO4

2-

+ NO3

-

+ Cl-+ F-+ (ANC + H+) (2)

Table 4-2. Factors for converting mg/L units or pH units to μeq/L units.

Analyte To Convert from mg/L to μeq/L, multiply by:

Ca2+ 49.90

Mg2+ 82.29

Na+ 43.50

K+ 25.58

NH4

+ (mg NH4/L) 55.44

NH4

+ (mg N/L) 71.39

SO4

2- 20.82

SO4

2- (mg S/L) 62.38

ANC (mg CaCO3/L) 19.98

Cl- 28.21

F- 52.63

NO3

-

(mg NO3/L) 16.13

NO3

- (mg N/L) 71.39

Aln+ (inorganic monomeric) 74.131 (assumes a +2 charge)

DOC2 5 to 10 (rough approximation)

To convert pH to H+ in units of μeq/L: H+ (μeq/L) = 10-pH x 1,000,000

1 The factor given for conversion of Al concentration from mass units to equivalence units assumes an average 

charge of +2 on the inorganic Al species present in the water. If the water sample has very low pH (less than 

about 4.8), then use a factor of 111.19 instead of 74.13 (assumes average charge of +3). 2 To convert DOC in mg/L to DOC concentration in μmol/L, multiply by 83.33. To estimate the equivalent 

concentration of organic acid anions from the DOC, multiply DOC in mg/L by a value of 5 to 10 to generate a 

very rough estimate of organic acid anion concentration in μeq/L.

The (ANC + H+) term in the anion sum is determined by laboratory measurements of ANC and 

pH. The hydrogen ion concentration (H+

) is calculated from pH as described in Table 4-1. To 

make this conversion on a calculator: take the pH value; change the sign to negative; hit the 

inverse log10 key; and multiply by 106

. 

A charge balance plot should be made by plotting the sum of anions versus the sum of cations 

(Figure 4-1). Some deviation from the one-to-one line (i.e., y = x) is expected because of 

analytical errors associated with the measurement of the individual anions and cations. Although 

random analytical errors would tend to cancel out in calculating the sum of anions or cations, the

analytical accuracy and precision and their relative contribution to the ion sum differ for each of 

the ions measured. Some charge imbalance may occur because of differences in analytical 

precision and accuracy. Thus, the calculated charge balance is an imprecise measure of data 

quality. It is useful as a tool for determining rather large deviations from the expected relative 

concentrations of anions and cations.
SECTION 4. DATA ANALYSIS PROTOCOLS

95

Figure 4-1. Cation sum versus anion sum. In this figure, two potential outliers warrant further 

investigation to determine if an error was made in analyzing or reporting the 

concentration of one or more ions.

Percent ion balance differences (% IBD; [cation sum - anion sum]/[cation sum + anion sum] x 

100) should be calculated for all samples. As a general guideline, we recommend the following 

criteria for % IBD (Table 4-3):

If the sum of anions + cations ≤ 100 μeq/L, % IBD should be ≤ 25%.

If the sum of anions + cations > 100 μeq/L, % IBD should be ≤ 10%.

For any samples that do not satisfy these criteria, the analytical data should be reviewed to 

determine if the cause of the imbalance is due to data entry error, analysis error, or some other 

identifiable error. If laboratory analysis error is discovered, the sample should be reanalyzed for 

those analytes that do not exceed laboratory holding times. If no error can be determined through 

data review and/or reanalysis, the results can be finalized without change, assuming that the 

imbalance is due to unmeasured ions.

Although the calculated charge balances do not include all ions that could potentially contribute 

to the sum of the cations and anions, those that are included contribute most to the overall anion 

and cation sum in most dilute freshwater environments. Inorganic ions not included, such as P 

and trace metals, are generally present in relatively low concentrations in most low-ionic strength 

waters and are not significant contributors to the total ion balance. Silica is not included in the 

charge balance because in most natural waters it exists predominantly in an uncharged form and 

does not contribute to either the anion or cation charge balance. 
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Table 4-3. Data validation quality control procedures. (Source: Paulsen 1997.)

Activity or Procedure Requirements and Corrective Action

Range checks, summary statistics, frequency distributions, 

and/or other exploratory data analysis (e.g., box and whisker 

plots).

Identify suspect values. Review field notes and laboratory data for 

possible problems or errors. Correct reporting errors or qualify as 

suspect or potentially invalid.

Ion balance. Calculate percent ion balance difference (% 

IBD) using data from cations, anions, pH, and ANC.

If total ionic strength1 ≤ 100 μeq/L, % IBD should be ≤ 25%.

If total ionic strength > 100 μeq/L, % IBD should be ≤ 10%. 

Determine, if possible, which analytes are the largest contributors 

to the ion imbalance. Review suspect analytes for possible 

analytical error and reanalyze any samples for which analytical 

error appears likely.

If analytical error is not indicated, qualify sample to attribute 

imbalance to unmeasured ions. Reanalysis is not required.

Conductivity check. Compare measured conductivity of each 

sample to a calculated conductivity based on the equivalent 

conductances of all major ions in solution.

If measured conductivity ≤ 25 μS/cm,

([measured - calculated] ÷ measured) should be ≤ ±25%.

If measured conductivity > 25 μS/cm,

([measured - calculated] ÷ measured) should be ≤ ±15%.

Determine, if possible, which analytes are the largest contributors 

to the difference between calculated and measured conductivity.

Review suspect analytes for analytical error and reanalyze any 

samples for which analytical error appears likely.

If analytical error is not indicated, qualify sample to attribute 

conductivity difference to unmeasured ions. Reanalysis is not 

required.

Aluminum check. Compare results for organic monomeric 

aluminum and total monomeric aluminum. 

[Organic monomeric] should be < [total monomeric]. 

Review suspect measurement(s) to confirm if analytical error is 

responsible for inconsistency. 

ANC check: 

1) Calculate carbonate alkalinity based on pH and 

dissolved inorganic carbon (DIC). Compare to 

measured ANC.

2) Calculate charge balance ANC and compare with 

laboratory measured (titrated) ANC.

Review suspect measurements for samples with titrated ANC < 

carbonate alkalinity or those with differences > 15% or >15 μeq/L 

for samples with ANC < 150 μeq/L. Determine if data entry error, 

analytical error, or non-carbonate alkalinity is likely to be 

responsible for lack of agreement.

Review samples having ([measured – calculated] ÷ measured) > ± 

15% (for low DOC waters) to 20% (high DOC waters). Determine if 

observed discrepancy can be attributed to organic anions. 

Strong organic acid anions are expected to decrease titrated ANC, 

compared with calculated charge balance ANC, by an amount 

equal to approximately (as a crude approximation) five times the 

DOC concentration in mg/L for acidic waters (ANC < 0 μeq/L) to 10 

times the DOC concentration in mg/L. Determine, if possible, if data 

entry error or analytical error is likely to be responsible for the 

observed inconsistency. 

1 Total ionic strength is calculated as the sum of cations (Equation 1) added to the sum of anions (Equation 2)

Two types of water bodies, however, often have charge imbalance due to ions that are not 

included in Equations 1 and 2. In acidic waters (pH less than about 5.5), aluminum (Al), which 

becomes more soluble with decreasing pH, may be a major contributor to the cation sum. Also, in 

waters with relatively large amounts of dissolved organic carbon (DOC higher than about 3 mg/L 

[250 μmol/L] to 5 mg/L [417 μmol/L]), organic anions can be a major contributor to the anion 

sum. 
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97

In acidic waters, failure to include Al in the charge balance may cause a cation deficit (anion sum 

higher than cation sum). At pH greater than 5.5, Al concentrations are typically so low as to be 

unimportant in the overall ion balance. At lower pH, however, Al should be incorporated into the 

cation sum for charge balance checks. Typically, there are several different forms (species) of 

inorganic Al, and they can have different charges. The concentration of Al in μeq/L can be 

approximated by converting measured values of Ali in mg/L to equivalence units using the 

conversion factor in Table 4-2, which assumes a +2 average charge for Al. For highly acidic 

waters (pH less than about 4.8), an average charge of +3 should be assumed for Al (as given in 

the footnote to Table 4-2). Alternatively, if Al is not measured, a cation deficit (anion sum higher 

than cation sum) in acidic waters (pH less than about 5.2) should not necessarily be interpreted as 

a QA problem as it can be assumed that some or all of the cation deficit results from unmeasured

Al.

Naturally occurring organic anions (derived from organic acids) contribute to the overall anion 

sum. Because there is no direct measure of organic anions, they are typically not included in the 

anion sum as represented in Equation 2. Where they are present in significant concentrations, the 

charge balance will show an anion deficit (cation sum higher than anion sum). Dissolved organic 

carbon concentration may be used as a surrogate variable for organic anions to check whether any 

observed anion deficit could be related to organic acids. In general, when DOC is less than about 

3 mg/L (250 μmol/L) to 5 mg/L (417 μmol/L) organic anion contributions to the ion balance are 

relatively minor and can be ignored. When DOC is greater than about 5 mg/L, there should be an 

appreciable anion deficit, calculated with the following equation:

Anion deficit = (cation sum) – (anion sum) (3)

The anion deficit should be roughly proportional to the DOC, with higher anion deficit in samples 

having higher DOC, and to some extent also higher pH. In general, the slope of the plot of anion 

deficit (in μeq/L; y-axis) versus DOC (in mg/L; x-axis) should be about 5 to 10 μeq of anion 

deficit per mg of DOC.

4.2.2 Calculated Versus Measured Conductivity 

The presence of ions in water increases the electrical conductivity (also called specific 

conductance) of that solution. Conductivity, therefore, provides an indication of total ion 

concentration. Further, since the relationship between ion concentration and conductivity is 

known for most ionic species, the measured conductivity of a water sample can be used as an 

internal check on both the accuracy and the completeness of the measurements of ionic species by 

comparing the measured and expected conductivity. The expected conductivity is calculated as 

the sum of the product of the ionic concentration times the equivalent conductances of each of the 

measured ions in water. For relatively dilute waters (conductivity below 200 μS/cm), Equation 4 

is used. For higher conductivity waters, a more complex equation is used that adjusts for high 

concentration effects. All waters that are sensitive to acidification from acidic deposition, and 

most waters that are sensitive to nutrient enrichment effects from atmospheric N deposition will 

have conductivity less than 200 μS/cm. Thus, we recommend use of Equation 4 and do not 

present the more complex equation. All of the concentrations in the equation need to be expressed 

in units of μeq/L. Conversion factors to convert from mass units to equivalence units are given in 

Table 4-2. For samples having conductivity lower than 200 μS/cm:
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Calculated conductivity = ((Ca2+ x 59.47) + (Mg2+ x 53.0) + (K+ x 73.48) + 

(Na+ x 50.08) + (NH4

+ x 73.5) + (H+ x 349.65) + (SO42-

x 80.0) + 

((ANC+H+

- OH-) x 44.5) + (Clx 76.31) + (NO3

-

x 71.42) + 

(OHx 198)) / 1000 (4)

Calculated conductivity should be plotted against measured conductivity as a first step to look for 

gross outliers (data values that fall well outside the normal range; Figure 4-2). As a more 

quantitative QA check, the % conductivity difference should be calculated as:

% conductivity difference = (calculated-measured)/measured x 100 (5)

Figure 4-2. Calculated versus measured conductivity. Two obvious outliers warrant further 

investigation to determine if an error was made in analyzing or reporting the 

concentration of one or more ions.

As a general guideline, we recommend careful review of samples for which the % conductivity 

difference exceeds 25% for samples in which measured conductivity is less than 25 μS/cm. We 

further recommend careful review of samples for potential data entry or analysis error if the % 

conductivity difference exceeds 15% for samples in which measured conductivity is more than 25 

μS/cm (Table 4-3).

4.2.3 Calculated Versus Measured ANC 

CARBONATE ALKALINITY VERSUS TITRATED ANC

There are two methods for evaluating the internal consistency of the dataset on the basis of 

observed differences between laboratory measurements (titrations) of ANC and calculated ANC 

or carbonate alkalinity using various ion measurements. The first involves comparisons between 

calculated carbonate alkalinity and laboratory measures of ANC made by acid titration. In almost 

all surface waters, the vast majority of ANC is made up of carbonate alkalinity (Figure 4-3). 

Carbonate alkalinity ([Alkc]) is calculated directly from laboratory measurements of pH and DIC 

concentrations (Hillman et al. 1987) and is a measure of just the carbonate ions (HCO3

- and CO32

) 

in the sample that would react with acid during an ANC titration. Alkc in μeq/L is calculated from 

the equation:
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Alkc=((DIC/12011) x ((Hmolar x K1 + 2 x K1 x K2)/(Hmolar x Hmolar + Hmolar x K1 + 

K1 x K2))) (KW/Hmolar) - Hmolar) x 106 (6)

where DIC is in mg/L, Hmolar = 10-pH, K1=4.4463x10-7, K2=4.6881x10-11, and 

KW=1.01x10-14. 

Figure 4-3. Calculated carbonate alkalinity versus laboratory titrated ANC with no obvious outliers. In 

general, calculated carbonate alkalinity is slightly lower than measured ANC (as it should 

be). 

ANC is a measure of all ions that react with acid during the acid titration. It includes all the 

carbonate ions that are represented in Alkc. Thus, ANC must be greater than or equal to Alkc. If 

calculated Alkc is higher than the titrated ANC, the discrepancy must be due to analytical error(s) 

in the measurement of ANC, pH, or DIC; the presence of noncarbonate ions that react with acid 

during the titration; or a combination of both. Comparisons of the Alkc and titrated ANC can 

serve as a QC check on the measured pH, DIC, and ANC. In nonacidic (ANC greater than 0 

μeq/L) waters with low DOC, samples that have ANC less than Alkc or those with (ANC-Alkc) 

differences greater than 15% (greater than 15 μeq/L for samples with ANC less than 150 μeq/L)

should be carefully reviewed for potential QA problems (Table 4-3). Acidic waters (ANC less 

than 0 μeq/L) and higher DOC waters (above about 3 mg/L [250 μmol/L] to 5 mg/L [417 

μmol/L]) often have other ions (Al, weak organic acid anions) that react with acid during the 

titration. Therefore, [ANC-Alkc] differences do not necessarily indicate a QA problem, but they 

do suggest that the data should be reviewed for potential errors.

CALCULATED VERSUS TITRATED ANC

The second QA check of ANC values compares laboratory titrated ANC with a charge balance 

definition of ANC, calculated as:

ANCG = Ca2+ + Mg2+ + K+ + Na+ + NH4+ + Al2+ - SO4

2-

- NO3

-

- Cl- (7)

where ANCG is laboratory Gran titrated ANC, and all parameters are expressed in units of 

μeq/L. 

-100

100

300

500

700

900

-100 100 300 500 700 900

Calculated Carbonate Alkalinity (μeq/L) .

Measured ANC (μeq/L)
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For Al species, assume an average charge of +2 (thus, Al concentration in μeq/L equals Al 

concentrations in μmol/L times 2) for waters having pH above about 4.8. For waters having lower 

pH, assume an average charge on the Al species of +3 (thus, Al concentration in μeq/L equals Al

concentration in μmol/L times three). For low DOC (less than about 3 mg/L [250 μmol/L] to 5 

mg/L [417 μmol/L]) waters, laboratory titrated and calculated charge balance ANC should be 

approximately equal, plus or minus an allowance for analytical errors. In general, the errors on 

the individual ions should cancel each other out and the two estimates of ANC should be within 

about 15% of each other (or within about 15 μeq/L for relatively low ANC [less than 50 μeq/L] 

waters). If they differ by more than this amount, it suggests errors in one or more of the 

measurements that go into the calculations and the comparison. 

For higher DOC waters (greater than about 3 mg/L [250 μmol/L] to 5 mg/L [417 μmol/L]), 

laboratory titrated ANC should be lower than calculated charge balance ANC by an amount equal 

to the concentration of strong organic acid anions in solution. That concentration of strong 

organic acid anions can be roughly approximated by multiplying the DOC concentration, 

expressed in mg/L, by the estimated organic acid charge density (average charge per mg of 

DOC). Thus, for high DOC waters, DOC-adjusted titrated ANC is calculated as:

DOC-adjusted ANCG = Ca2+ + Mg2+ + Na+ + K+ + NH4+ - SO4

2-

- NO3

-

-Cl-- A- (8)

where Ais the estimated strong organic acid anion concentration (defined as those with acid 

dissociation constants giving them an equilibrium pH less than about 4), which is 

very roughly approximated by:

A-(μeq/mg) ≈ DOC (mg/L) x 4 μeq/mg (9)

If the DOC-adjusted laboratory titrated ANC differs from calculated charge balance ANC by 

more than about 20% (or 20 μeq/L for relatively low [≤ 50 μeq/L] ANC samples) in high DOC 

waters, that suggests the possibility of data entry error or analytical error in one or more of the 

parameters that enter into the calculations. In that case, laboratory and data entry records should 

be reviewed for possible errors. 

Note that these methods for estimation of the equivalent concentrations of Al and strong organic 

acid anions are crude approximations for the purpose of evaluating the internal consistency of the 

dataset and for identifying possible incorrect values for further examination. For high-DOC 

waters in particular, lack of agreement between calculated and titrated ANC does not necessarily 

mean that there are errors in the dataset. More rigorous approaches are available for calculating 

the equivalent concentrations of Al and organic anions, but these are not needed for the purpose 

of dataset validation.

4.2.4 Other Validation Procedures

The dataset should be examined in other ways to look for outliers (data values that fall well 

outside the normal range for that water body over multiple samplings or for multiple water bodies 

within a forest or region). The range of values in the dataset and/or a histogram plot (Figure 4-4) 

should be used to look for outliers in all variables. Outliers may also be identified by plotting 

each variable by sample date to look for isolated gross variations over time. Analysis and sample 

collection records should then be reviewed to determine if the cause of any outlier is likely due to 
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101

data entry error, analytical error in the laboratory, or sampling error in the field. If errors are 

discovered, samples can be reanalyzed or rejected. If no error can be determined, the results 

should be assumed to be correct and accepted without change. Outliers should not be rejected 

unless there is a strictly objective basis for rejection. If there is a clearly identified error, the result 

should be rejected and, if possible, corrected; if there is an unexplained anomaly, the data should 

be retained.

Another useful procedure is to plot variables in the dataset against each other for variables that 

are known to be highly correlated. Examples of strongly correlated variables can include Ca2+–

Mg2+, Ca2+–ANC, Na+

–Cl-, DOC–color, and N–P. Data points that fall outside the cloud of data 

points defining the general relationship warrant closer examination.

Figure 4-4. Patterns in DOC concentration in streams within a particular region. The data are not 

normally distributed; rather, they are skewed towards lower concentrations (≤ 3 mg/L).

4.2.5 Final Data Quality Determination

Each of the dataset internal consistency checks outlined above provides an opportunity to identify 

potential problems in the data related to sampling error, data entry error, and/or laboratory error. 

Some of the problems identified through these analyses might be corrected by reanalysis or 

simply by replacing a value that was entered incorrectly into the dataset. In other cases, the cause 

of the anomaly will be unknown and will represent an error of some sort or the presence of one or 

more unmeasured analytes. Such unexplained deviations from expected patterns should generally 

not be altered or deleted from the dataset. As described in the preceding sections, the final 

validated dataset should show:

• Good charge balance agreement;

• Good agreement between measured and calculated conductivity;

• Good agreement between laboratory titrated and defined ANC based on various ion 

measurements;

• Reasonable (readily understandable) distribution of parameter values as reflected in 

frequency distributions across space and/or across time; and
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• Clear patterns between paired variables that are known to be strongly correlated with each 

other. 

Note that there is generally no clear-cut definition of what constitutes “good” or “reasonable” 

agreement, although targets for percent and absolute variation are presented where applicable. 

The purpose of these analyses is not to discard measurements that are not completely 

understandable but rather to identify the samples and/or measurements that appear to have a 

higher likelihood of some kind of error. In the best of cases, the error is identified and corrected. 

In other cases, the error remains unknown or there may not be an error at all but rather an aspect 

of the water chemistry that is not fully understood. 

Unless these internal consistency checks are conducted and unless the dataset is found to be 

generally internally consistent, it is not possible to determine whether analysis of these data will 

yield meaningful and representative results. To the extent that the water chemistry data make 

sense, greater confidence can be placed in conclusions drawn from analysis of those data. 

4.3 APPLY PROCEDURES TO PREPARE RAW DATA 

FOR GRAPHICAL AND STATISTICAL ANALYSIS

4.3.1 Censored Data

Data that have reduced certainty are often censored for reporting or analysis purposes or both. 

Examples can include measured values below the MDL or measured values below the reporting 

limit (e.g., if nitrate concentration is reported as being less than 1 μeq/L). Censored data can 

cause problems in statistical calculations if there is no real number that can be used in the 

calculations. Other problems can arise in deciding what to do with censored data when reporting 

limits are approximately the same as analyte concentrations of ecological concern. If detection 

and reporting limits are well below any real level of concern, there is generally no substantial 

problem with interpretation or treatment of censored data. However, when detection or reporting 

limits are approximately at or below the same level as the level of concern, then interpretation 

based on censored data may be problematic regardless of how censored data are handled.

We recommend that laboratory measurements below the MDL be reported by the laboratory as 

zeros. These measurements are not statistically different from zero. We recommend that 

measured values above the MDL but below the reporting limit be flagged as having potentially 

higher uncertainty than do values above the reporting limit. Nevertheless, these values are entered 

as measured into the database and used as measured in data analyses, with no adjustment. We do 

not recommend reporting any data values in the dataset as being less than a particular value. 

Rather we recommend reporting actual values, including zeros for samples measured as below the 

MDL and including measured values that are less than the reporting limit but above the MDL.

4.3.2 Outliers and Missing Values

Outliers can be difficult to identify and interpret. Their importance is dependent on the type and 

objective of the analysis being conducted. There are a number of statistical outlier tests that one 

could apply but we caution against removing any outlier unless there is a good argument for 

removal based on a clearly identified analytical error or data management issue (e.g.,
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103

typographical error). It is often useful to run the statistical analysis with and without any 

suspected outliers to see if the results are substantially different. If not, then the point is moot and 

it doesn’t matter. If the outlier causes a big change in the results, then it will require some careful 

reexamination of the data before deciding what to do. In general, any conclusion that relies on the 

presence of one or a few extreme values should not be considered a robust conclusion. If it is 

determined that an important conclusion does depend on the inclusion of one or a few data 

outliers, we recommend not drawing that conclusion, but rather going back into the field to 

collect additional data. 

In general, because they are not routinely removed from the dataset, outliers can be subjectively 

identified visually, without a formal statistical test. However, if a measured value appears to be an 

outlier, the analyst may wish to eliminate this deviant value and not include it in various 

calculations, data analyses, or data presentations. This generally should not be done unless it can 

be objectively determined that the questionable value is indeed likely to be erroneous. We 

recommend Dixon’s Q-test as a relatively simple test to determine outlier status. The test is 

conducted as follows:

1. Arrange the values of the observations in ascending order.

2. Calculate the experimental Q-statistic (Qexp) as the ratio of the difference between the 

suspect value and the value of its nearest neighbor (in the ascending series of values that 

comprise the dataset) divided by the range of values in the dataset. For example, to test 

whether the lowest value (x1) is an outlier, calculate Qexp as: 

(10)

where x1 is the lowest value in the series

x2 is the second lowest value in the series

xN is the highest value in the series

3. Similarly, to test whether the highest value is an outlier:

(11)

where xN-1 is the second highest value in the series and the other terms are as defined 

above. 

4. Compare the calculated Qexp to a critical Q-value (Qcrit) that is taken from a table (see, for 

example, Table 4-4, Rorabacher 1991). You must first choose your confidence level (CL). 

We recommend a 95% CL. For example, as shown in the table, at the 95% CL and a total 

number of measured values equal to 9, the Qcrit is 0.493. 

5. If Qexp > Qcrit, the questionable value can be designated as an outlier.

Missing values are a fact of life in most statistical analyses of environmental data. They are more 

problematic in parametric tests such as regression or analysis of variance than in non-parametric 

tests. Parametric tests are used for estimating parameter values and testing hypotheses concerning 

them when the form of the underlying data distribution is known (typically, the data are normally 
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distributed). For tests of data for which we do not know the underlying data distribution, 

including those that are not normally distributed, nonparametric tests must be used. These tests 

compare the distributions, rather than the parameters. 

Missing values can be synthesized from other data, but we would not recommend this approach 

as a general procedure without careful consideration. The EPA’s National Surface Water Surveys 

synthesized a small number of missing values using regional regression models in order to make 

complete regional population estimates. The general approach is to substitute for the missing 

value a synthetic value developed from the remainder of the dataset or published relationships. 

The synthetic value can be calculated as the median of the existing measured values for that 

parameter or using a regression relationship based on one or more other variables. For example, if 

a measured value of Na+ is missing, one can estimate the missing Na+ concentration from the 

measured Cl- concentration using a regression approach based on Na+ and Cl- measurements in 

the dataset. Thus, the regression equation, developed from the existing data, with which to 

estimate the Na+ concentration from the measured Cl- concentration should be used to estimate 

any missing values of Na+ concentration. Similarly, Ali concentration can be estimated from pH 

or H+ concentration using a linear regression approach. 

The median of existing measured values can also provide a reasonable substitute for a missing 

value. However, one should be careful to avoid using the median of data points known to exhibit 

a wide range of values, especially when there is an opportunity to reduce that variability. For 

example, if a Ca2+ concentration measurement is missing from a dataset containing first- through 

fifth-order streams and where the Ca2+ concentration varies strongly with stream order, it would 

be better to take the median of all streams in the dataset that are of the same order as the stream 

having the missing value rather than the median of all streams of all stream orders. 

It can be considered acceptable to create synthetic substitutes for a small number of missing 

values, but these should generally not constitute more than 5% of the data for any variable. In 

general, we recommend not creating synthetic substitutes for missing values unless these missing 

values prevent the use of a particular analysis needed for a project objective. For example, a 

principal components analysis cannot be performed on lake ion chemistry using samples that 

have one or more missing variable values. Thus, any sample that has even one missing value 

cannot enter into the analysis unless the void is first filled with a synthetic value. Furthermore, 

some missing values may not be particularly important to interpretation of the data (for example, 

a missing NH4

+ concentration in a lake that is expected to be very low). It is advisable to avoid, if 

possible, deleting that entire sample from the analysis simply because the NH4

+ measurement is 

missing. If synthetic values are to be constructed, we recommend using the most robust empirical 

approach that can be developed from that particular dataset.
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105

Table 4-4. Critical Q-values for Dixon’s outlier Q-test, at the 0.95 confidence level.

(Source: Rorabacher 1991.)

Number of Measurements Qcrit

(CL: 95%)

Number of 

Measurements

Qcrit

(CL: 95%)

3 0.970 17 0.365

4 0.829 18 0.356

5 0.710 19 0.349

6 0.625 20 0.342

7 0.568 21 0.337

8 0.526 22 0.331

9 0.493 23 0.326

10 0.466 24 0.321

11 0.444 25 0.317

12 0.426 26 0.312

13 0.410 27 0.308

14 0.396 28 0.305

15 0.384 29 0.301

16 0.374 30 0.298

4.3.3 Multiple Observations 

We do not recommend averaging the results of replicate (duplicate or triplicate) samples in the 

dataset. Rather, the first sample collected at a given site and sample occasion is considered to be 

the normal sample. It is used in statistical and other data analyses to represent the chemistry of 

that lake or stream on that sampling occasion. Any second or third sample (replicate) collected on 

that sampling occasion is used only for QA purposes to assist in quantifying the cumulative 

variability and error associated with the collection and analysis of the water in that lake or stream. 

The replicate sample result is not used in routine data analyses. 

If multiple samples are collected within a given rainstorm, season, or year, results of analyses of 

those samples are maintained as separate values in the dataset. Depending on the objectives of a 

particular study or analysis, they might be averaged in the process of analyzing the data. For 

example, if the objective is to compare spring base-flow chemistry across streams in a particular 

forest, one may choose to average all samples collected during the spring season (avoiding 

rainstorm and snowmelt periods) over a finite period of time (perhaps 5 years). Such an approach 

is appropriate if, for example, some streams in the forest were sampled only once and others 

multiple times within that 5-year period. If there is reason to believe that stream chemistry 

changed appreciably during that 5-year period, then it may not be advisable to average the data 

across multiple years. Instead, one may choose to use the spring base-flow sample collected at the 

time closest to April of a particular year, for example. One should be particularly careful about 

averaging multiyear data if part of the sampling window occurred during a wet year(s) and part 

during a dry year(s). 
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If most or all sample sites were sampled during multiple years, an analysis of the spatial 

distribution of water chemistry across a forest will often be conducted using 3- or 5-year averages 

of chemistry to represent each site. Such averages should not combine samples collected during 

different seasons unless it is clear that seasonality is not an important issue. 

4.3.4 Treatment of Zeros and Negative Values

For studies of dilute surface waters potentially impacted by air pollutants, the only major variable 

expected to have negative values is ANC. Some, but not all, lake or stream datasets will have 

some negative ANC values. Because negative ANC values are real measurements, they must be 

left as negative numbers. However, some transformations (e.g., log transforms) required for some 

statistical analyses may only be applicable to non-negative and non-zero numbers. If that 

becomes an issue for an analysis planned for a particular dataset, add a constant whole number 

just larger than the largest negative number in the data (i.e., add 50 μeq/L if the lowest ANC is -

49) to all ANC measurements so that there are no longer any negative numbers in the analysis. 

This should be done only for that particular analysis. Designate the new variable as [ANC + 50 

μeq/L]. This manipulation of the data must be taken into consideration in interpreting the results 

of the analysis. For zero values, we recommend adding one to all values of that variable when 

almost all the data are greater than one, and changing the name of the revised variables to be used 

in the analysis to, for example (sodium + 1 μeq/L). This works well for log10 transforms as when 

x=0, log10(x+1)=0. When many of the data values are less than one, add a constant number that is 

smaller than almost all the data values to each zero value in the dataset. For example, zero values 

for NH4

+ concentration, which may be fairly common in many surface water datasets, may be 

adjusted by adding a constant of 0.001 μeq/L before transformation. 

4.3.5 Treatment of Seasonality

Seasonal variation in water chemistry data can affect data analysis and interpretation in two 

fundamental ways. First, chemical parameters that affect the suitability of the water to support 

various species and biological communities tend to vary with season. This is the case in many 

waters with respect to pH, ANC, Ali, DOC, NO3

-

, SO4

2-

, and base cation concentrations. Thus, the 

chemical conditions that are most stressful to biota may occur to a greater or lesser degree 

depending on season. These seasonal differences are most pronounced in regions that experience 

substantial seasonal changes in rainfall or temperature. Interpretation of chemical parameter 

values above or below known biological stress thresholds will be highly influenced by when the 

samples were collected. 

Second, seasonality in the data can affect certain statistical tests (such as trends analysis, for 

example). A dataset having substantial seasonality may require use of different statistical tests as 

compared with a dataset lacking seasonality. This is further discussed in Section 4.5, Conduct 

Statistical Analysis, of this DAP. 

This DAP does not recommend the need for any particular adjustment of seasonal data. Some 

sampling studies may choose to reduce the effects of seasonality on the data by careful timing of 

field activities. Other studies may strive to quantify the seasonality that occurs. It can also be very 

useful to quantify the relationship between annual average or base-flow chemistry and observed 

extreme values that are influenced by season and/or episodic processes. For example, Sullivan et 
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107

al. (2003) illustrated the relationship between median spring season ANC and the minimum ANC 

reported in the data record for streams in Shenandoah National Park (Figure 4-5). Such an 

analysis could also be conducted to compare spring median or spring minimum ANC to summer 

or fall index ANC. These kinds of relationships can be useful in evaluating the likelihood of 

experiencing extreme values that exceed various response thresholds for expected biological 

effects.

Figure 4-5. Minimum stream ANC sampled at each site during each year versus median spring ANC 

for all samples collected at that site during that spring season. Data are provided for all 

intensively studied streams within Shenandoah National Park during the period 1993-

1999. A 1:1 line is provided for reference. The vertical distance from each sample point 

upwards to the 1:1 line indicates the ANC difference between the median spring value 

and the lowest sample value for each site and year.

4.4 CONDUCT EXPLORATORY ANALYSES

4.4.1 Analysis of Water Quality Status

Various graphical and statistical methods are available for describing ambient water quality and 

assessing differences in water quality across a forest or region. Current status of water quality 

should be compared among sites, with previously obtained data for individual sites, with criteria 

values or standards used in water quality assessments, and with values that represent ecological 

thresholds. 

There is no standard procedure for the statistical analysis of water quality data for the purpose of 

evaluating sensitivity to, or effects from, atmospheric deposition. Rather, there exists a range of 

options for depicting results and/or analyzing differences over space or time. Selection of 

methods will depend on a host of issues, including project objectives; the quantity and quality of 

the data; number of sampling locations; length of the period of record; extent to which samples 

were collected across years, seasons, and hydrological episodes; and specifics of the resulting 

dataset. Important dataset issues include the presence and abundance of extreme outliers, 

censored data, and negative values; normal versus non-normal distribution of the data; seasonality 
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108

and episodicity of the data; and extent to which data values are missing and/or are lower than 

reporting limits. It is generally advisable to consult with a statistician or a staff member who is 

knowledgeable in statistics before conducting trends analyses and other complex statistical 

analyses. Nevertheless, there are some commonly used and accepted data analysis approaches and 

statistical tests that are often applicable to the types of data analyses needed by the FS ARM 

program. These are described in the sections that follow. 

4.4.2 Graphing and Quality Analysis

Graphics used to visualize water quality data include scatter plots of values for single or multiple 

sites by date. Water quality should also be examined relative to continuous variables, such as 

elevation, watershed area, or discharge. The range and distribution of data for different periods of 

time or for different lakes or streams can be depicted with histograms (Figure 4-4) or box-andwhisker plots (Figure 4-6). The box plot graphically represents the central tendency and 

variability in a dataset. The range indicated by the box (top to bottom) represents the middle half 

of the data and is bisected by a line that represents the median value of the data. Because the 

bottom of the box represents the lower-quartile (25th percentile) of the data and the top of the box 

represents the upper-quartile (75th percentile) of the data, the vertical length of the box represents 

the “interquartile range” (IQR) of the data. The end of each whisker represents the last value from 

the dataset that is no more than 1.5 times the IQR. The outliers (values beyond 1.5 times the IQR) 

are all shown on the plot. Data points marked with a star are greater than 1.5 times but less than 

three times the IQR and are considered possible outliers; those that are marked with an open 

circle are greater than three times the IQR and are considered probable outliers.

Figure 4-6. Box plots comparing hypothetical lake ANC values measured in samples collected 

during the spring versus the fall season.

Graphics should be used to examine temporal variation in data for individual sites, including 

patterns associated with season or discharge, as well as gradual or more sudden changes in 

values. Spatial variation among multiple sites can also be represented graphically, including 

variation related to differences in watershed properties, land use, or exposure to pollutants. Even 
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109

when a more quantitative statistical analysis of water quality data is desired, qualitative visual 

data examination is recommended as a preliminary step.

The steps that one should take in analyzing the dataset will depend to a large degree on the 

specifics of the dataset itself and the purpose of the analysis. Common management issues for FS 

ARM program staff that involve analysis of surface water field sampling data are outlined in 

Table 4-1. That table identifies six major approaches (each tied to a purpose):

1. Determine whether one lake or stream, or a group of lakes or streams, is N-limited for algal 

growth.

2. Quantify episodic excursions from base flow conditions in surface water chemistry during 

hydrologic events.

3. Determine the distribution of lake or stream water chemistry across a particular forest or 

wilderness.

4. Quantify long-term changes in lake or stream ANC (or other variable) over time in a 

particular lake or stream.

5. Determine to what extent air pollution is currently affecting the water resources in a 

particular forest or wilderness.

6. Evaluate whether the current condition of acid or nutrient sensitive waters warrants 

mitigation.

The analyses that could or should be conducted will depend in part on which approach is required 

to answer particular management questions.

Every dataset will offer its own challenges and, if sufficiently examined, reveal its own, often 

unique, patterns. Regional differences are important. Furthermore, water quality data analysis is 

exploratory in nature. To properly analyze a water quality dataset, the analyst must experiment 

with different approaches and eventually find some that work both with that dataset and those 

specific analysis objectives. 

Despite these difficulties and the site-specificity of water quality data analysis, it is possible to 

offer recommendations and examples of steps to be considered by FS ARM program staff. A 

successful analysis method will develop through trial and error. The example analyses illustrated 

in this section of the DAP show some of the approaches that we have found to be useful, and FS

staff may find some of these examples to be successful in some cases. Nevertheless, an analyst 

should always explore multiple options to determine what works best for a particular dataset. If 

the data are of high quality, it is likely that they will tell a story. Some creativity may be required 

to reveal that story.

4.4.3 Recommended Data Analyses

We recommend various types of data analysis in this section. These recommendations are specific 

to the purpose of the data analyses outlined in Section 1 of this document. Forest Service staff 

may find alternative approaches to be as or more successful than those provided here. There is no 

one clear choice of how to approach exploratory data analysis. 
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SUBSET THE DATA 

Exploratory data analyses should be conducted using all of the available data. In addition, 

however, it is often very helpful to also create various subsets of the data and analyze them 

individually. This is because inherent variability can obscure the patterns that might exist in the 

particular subset of the data that represents the more sensitive or affected bodies of water, time of 

year, hydrological conditions, geological settings, etc. Therefore, FS staff should explore various 

ways to subset their data before conducting exploratory analyses in order to determine whether

some patterns are only evident or are strongest for one or more subsets when compared with the 

dataset as a whole. 

Data can be subset for exploratory analysis using water chemical criteria (Table 4-5). In addition 

to using chemical criteria, data can be subset using features of the landscape, hydrology, or 

morphology (Table 4-6). This is an opportunity for the analyst to be creative. Try different 

approaches, and see what works. The objective is to improve your understanding of the data and 

the story that they have to tell.

Table 4-5. Example variables with which to subset water quality data, according to measured 

water chemistry for analysis.

Variable Possible Cutoff Values for Designating Lake or Stream Classes

ANC 0, 20, 50, 100 μeq/L

NO3

- 5, 10, 15 μeq/L

DOC 200, 400, 500 μM

pH 5.0, 5.5, 6.0, 6.5

Ali 2, 7 μM

Ca + Mg1 Highly region/forest specific

SO4

2- Highly region/forest specific

1Can be analyzed individually or in combination; in some cases (where they are quantitatively important), 

Na and/or K might also be included.

Table 4-6. Example variables with which to subset water quality data for analysis according to 

features other than measured water chemistry.

Variable Possible Lake or Stream Classes

Geologic class For example: siliciclastic, granitic, argillaceous, etc.

Elevation Can use above or below a specific cutoff, or as discrete elevational bands

Lake type Drainage, seepage, type of seepage lake (perched or flow-through)

Stream Strahler order Can combine into classes (i.e., 1st plus 2nd, 3rd plus 4th, etc.) or analyze as individual orders

DETERMINE WHETHER ONE LAKE OR STREAM, OR A GROUP OF LAKES OR 

STREAMS, IS N-LIMITED FOR ALGAL GROWTH 

Productivity of surface water can be limited by a multitude of factors. For example, small streams 

are commonly limited by available sunlight; if the stream is highly shaded by riparian vegetation, 

then primary productivity may be low even if nutrient concentrations are high enough to support 
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algal growth. Streams can also be limited by substrate type. For example, if suitable substrate is 

not available for attachment, then algal productivity may be low relative to nutrient availability. 

Streams and lakes can also be limited by presence of a nutrient, most commonly P and/or N; if N 

is limiting, then atmospheric contributions of N can enhance productivity, contribute to 

eutrophication, and perhaps alter species composition and abundance. 

The relative importance of N and P as potentially limiting or co-limiting nutrients can be 

evaluated by conducting a rough screening analysis based on the molar ratio of N:P 

concentrations in surface water. This ratio ideally should include all forms of N and P, both 

particulate and dissolved, both organic and inorganic. Thus, total N is the sum of the 

concentrations of NO3

-

-N, NH4

+

-N, and organic N. Units are in μmol/L for both elements. 

Phosphorus is measured as total P. If measurements of total N are not available, one can use an 

estimate of total inorganic N (TIN), calculated as the sum of the molar concentrations of NO3

- and 

NH4

+

. Note that the concentrations of NO3

-

-N in μmol/L are the same as the concentrations in 

μeq/L; no conversion is needed. Total N in μmol/L is equal to the concentration of total N in 

mg/L times 71.38. Total P in μmol/L is equal to the concentration in mg/L times 32.29. 

Based on available experimental data, a large majority of lakes that have total N:total P below 

about 44 have been found to indicate N-limited phytoplankton growth (Elser et al. 2009). This is 

an area of active research, and interpretations may be subject to change in the near future. Lakes 

and streams that are nutrient (N and/or P)-limited, rather than limited by light or some other 

factor, may change in their limitation status over time, perhaps with season. Therefore, temporal 

patterns in the N:P molar ratio could be examined over time. In addition, we recommend 

evaluation of spatial patterns in N:P to determine, for example, if water bodies in a forest tend to 

be N-limited primarily at certain elevations, on certain geological types, or in certain vegetation 

communities. Thus, N:P ratios could be mapped relative to landscape condition to reveal such 

patterns, if they occur. 

Determination that a water body or a group of water bodies exhibits potential N-limitation based 

on the N:P ratio is not sufficient evidence to indicate that the system is, in fact, N-limited. 

Furthermore, such an analysis does not necessarily identify any variability that might occur in 

that nutrient status throughout the growing season. These recommended N:P ratio analyses are 

screening-level analyses that may suggest the possibility of N-limitation. 

The next step in the assessment process is to conduct laboratory studies to determine N versus P 

limitation. These could involve collection of multiple liters of lake water, which is then shipped 

on ice to the laboratory. The water is dispersed into flasks (typically, at least three flasks per 

treatment). Treatments may involve multiple light levels and varying (low and high) nutrient 

additions of N only, P only, N+P, and control (no nutrient addition). Incubation, with continuous 

or periodic mixing, is conducted under approximate ambient lake temperature conditions. Algal 

growth can be tracked daily by measuring the chlorophyll a concentration in an aliquot of water 

from each flask. 

Based on the results of the laboratory incubation studies, and the degree of rigor required for the 

project in the determination of nutrient limitation, it may be desirable to progress to in situ

incubation studies. Such experiments should involve in situ incubation of water over a period of 

time during the growing season in multiple containers suspended in the lake or stream. The 
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containers should include a control (no nutrient addition), multiple N addition containers (at least 

two levels: high and low), multiple P addition containers, and multiple N + P addition containers. 

For example, one may double and triple the ambient nutrient concentrations in the two containers 

for each type of nutrient input. Changes in the concentration of chlorophyll a over time in the 

treatment containers, relative to the control, indicate productivity responses to nutrient addition. 

Such experiments can verify whether or not, and when, a lake or stream (or a group of lakes or 

streams) may be susceptible to eutrophication effects associated with atmospheric N deposition.

QUANTIFY EPISODIC EXCURSIONS FROM BASE FLOW CONDITIONS IN SURFACE 

WATER CHEMISTRY DURING HYDROLOGIC EVENTS 

Changes in the concentrations during episodes of major ions, pH, and ANC should be evaluated 

for a given lake or stream by plotting individual measured values during multiple storms. An 

example for one lake or stream during one storm or snowmelt episode is shown in Figure 4-7. 

Figure 4-7. Changes in the concentration of major water chemistry constituents during a 

hypothetical hydrological episode in one stream. Data for each variable of 

interest should be plotted along the same time axis and compared at the same 

scale relative to the pattern of discharge.

The extent to which ANC and pH decrease and the extent to which Ali increases in response to 

hydrological episodes provides an indication of the chemical extremes aquatic biota are exposed 

to during hydrological episodes. Patterns of episodic responses of the sum of base cations (SBC), 
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DOC, SO4

2-

, and NO3

-

concentration can reveal important information regarding the causes of 

episodic excursions of ANC, pH, and Ali. Both ANC and pH can decrease and Ali can increase, in 

response to base cation dilution (decreased SBC), NO3

- leaching, SO42- leaching, and DOC 

mobilization. The relative importance of these potential drivers varies by watershed, region, 

season, and hydrologic event. Examination of the kinds of plots shown in Figure 4-7 can reveal 

these patterns in the various potential drivers at one site during one event. It may be necessary to 

sample and analyze multiple sites and multiple events. 

Similarly, temporal patterns of changing water chemistry in a given lake or stream should be 

examined across the annual or seasonal cycle. In regions having marked snowpack development, 

such an analysis should include the entire snowmelt period, as is shown in Figure 4-8. 

Figure 4-8. Time series of major ions and discharge in Treasure Lake in the Sierra Nevada during 

snowmelt in 1993. (Source: Stoddard 1995.) Seasonal and episodic changes in surface 

water chemistry can be examined using these simple time series plots.
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Intensive time series data, where available, provide finer resolution of episodic changes in 

chemistry. For example, Figure 4-9 shows dramatic changes in NO3

- concentration in two alpine 

streams in Rocky Mountain National Park in response to seasonal and snowmelt patterns. In this 

example, peak stream water NO3

- concentrations occurred relatively early in the snowmelt 

process in May and June. Subsequently, there was an extended period of declining NO3

-

concentrations as snowmelt proceeded throughout the summer, followed by an increase during 

the fall.

Figure 4-9. Daily discharge (A) and nitrate concentration (B) in Icy Brook and Andrews Creek within 

the Loch Vale watershed, Rocky Mountain National Park, in April-September 1992.

(Source: Campbell et al. 1995.). This graphic shows an approach for displaying data 

from repeated sampling of two streams for the purpose of documenting changes in NO3

-

concentration as snowmelt proceeds within a given year.

An analysis of surface water NO3

- concentration as a fraction of the total combustion-related 

mineral acid anion (SO4

2- plus NO3-

) concentration can reveal the relative roles of NO3

- and SO42-

in influencing surface water chemistry. In many surface waters, under certain hydrological 

conditions, SO4

2- dominates the total mineral acid anion concentration. When this occurs, NO3-
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has relatively little influence on the ANC or pH of the water. There can, however, be times when 

NO3

- is also important to the acid-base status of the water. For example, data collected from four 

Adirondack Mountain streams during hydrological episodes (Figure 4-10) illustrated that, for the 

study streams, NO3

- generally provides less than half of the contribution of mineral acid anions 

from the atmosphere (SO4

2- provides the majority), but the relative importance of NO3-

, compared 

to SO4

2-

, increases at lower ANC values (which occur during high flow periods).

Figure 4-10. Ratio of NO3

-

:(SO4

2-

+ NO3

-

) concentration versus ANC in stream water samples 

collected during hydrological episodes in four streams included in the Adirondack region 

of EPA’s Episodic Response Program. The different symbols on the graph represent

different streams. (Source: Sullivan et al. 1997.)

DETERMINE THE DISTRIBUTION OF LAKE OR STREAM WATER CHEMISTRY 

ACROSS A PARTICULAR FOREST OR WILDERNESS

Patterns in water chemistry should be mapped to illustrate spatial patterns in the data. Figure 4-11 

shows one way to do that (in this case, for lake water NO3

- concentration in Adirondack lakes). 

Each bar represents one lake; the base of the bar reflects the lake location, and the height of each 

bar is proportional to the NO3

- concentration. In this example, concentrations of NO3- are highest 

in the southwestern portion of the Adirondack Park and in the central High Peaks region, the 

general locations where N deposition and precipitation amounts are highest. Thus, spatial patterns 

in surface water chemistry should be compared with various factors that are known or suspected 

to be associated with water chemistry. These might include geology, soils types, elevation, 

atmospheric deposition, precipitation amounts, vegetation types, etc.
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Figure 4-11. Map of summer NO3

-

concentrations in drainage lakes sampled by the 

Adirondack Lakes Survey Corporation in the Adirondack region of New York.

(Source: Sullivan et al. 1997.) Maps such as this can reveal spatial patterns in 

the concentration of any surface water variable across a study area.

Spatial patterns can also be analyzed across a gradient of deposition or across a gradient of 

expected resource sensitivity using space-for-time substitution analysis. In this approach, it is

assumed that the lakes or streams across the study area were initially relatively homogeneous in 

their chemistry, and furthermore, that differences observed across space at the present time

correspond with changes that occurred in the past. Figure 4-12 shows an example space-for-time 

substitution analysis across a gradient in atmospheric deposition across the Upper Midwest. Such

an analysis could be conducted across a gradient in elevation, deposition, slope steepness, etc. 

rather than, or in addition to, across a gradient in deposition. In this case, the data show decreases

in [HCO3

- - H+

], DOC, and [Ca + Mg] over a gradient of increasing acidic deposition. Similarly, 

the ratio of SO4

2- to SBC concentrations increases with increasing acidic deposition.

Furthermore, the slopes of the data on these graphs provide estimates of the quantitative 

importance of the various changes. Such quantitative estimates can be combined with model 

estimates of changes at selected locations in a weight-of evidence assessment.
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Figure 4-12. Example space-for-time substitution analysis for four variables across a gradient of expected 

response. This example is based on subsetting the available data, across a gradient of acidic 

deposition in the Midwest, to include only seepage lakes having low Cl- concentration (to 

remove road salt influence) and those having ANC below 50 μeq/L. Note that the variable 

[HCO3

-

- H+] is one representation of ANC. (Source: Sullivan 1990.)

Portions of a region or a forest within which most of the acid-sensitive waters are expected to 

occur can sometimes be identified and mapped. In an example from the Southern Appalachian 

Mountains Initiative study, the area delimited by an acidification sensitivity classification scheme 

is shown in Figure 4-13. The darkly shaded area includes the siliceous geologic sensitivity class 

surrounded by a 750 m buffer. In addition, all areas lower than 400 m elevation have been deleted 
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and areas higher than 1,000 m elevation have been added. The area thus circumscribed includes 

95% of the known acidic streams and 88% of the known streams having ANC ≤ 20 μeq/L (of 

more than 900 streams surveyed) within the region. Furthermore, all known streams having ANC

≤ 20 μeq/L are in close proximity to the final mapped area. FS staff can use an approach such as 

this to circumscribe portions of a forest thought to contain most of the sensitive or impacted water 

bodies. 

Figure 4-13. Results of a classification system devised to indicate expected low-ANC streams (in this 

example, based on geology and elevation) compared with actual low-ANC streams, 

which are represented as dots on the map. (Source: Sullivan et al. 2007.)

The distribution of data values across a given study region can reveal important information about 

the source of a constituent. For example, it can be helpful to plot the frequency distribution of 

surface water SO4

2- concentrations within a relatively small study area. Atmospheric S 

contributions to watersheds are expected to yield a reasonably well-defined bell-shaped or half 

bell-shaped curve in surface water SO4

2- concentrations. Differences from one study watershed to 

another in such features as elevation, aspect, vegetation type, and topography contribute to 

variability, but the overall patterns should be relatively homogeneous if the study area is 

relatively homogeneous. The observed outlier lakes or streams having much higher 

concentrations of SO4

2- than the population of lakes or streams at large can be presumed to 
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119

receive contributions of geological S unless there is a good reason why atmospheric deposition 

should be markedly higher at those outlier locations. For example, histograms showing the 

frequency of occurrence of SO4

2- concentration in lakes in four regions of EPA’s Western Lakes 

Survey show clear outlier values in the Sierra Nevada, NW Wyoming, and the Colorado Rocky 

Mountains (Figure 4-14). These lakes likely receive geological S from their watersheds. Lakes or 

streams that contain appreciable geological S are not good candidates for monitoring or study to 

quantify effects from atmospheric sources of S.

Figure 4-14. A histogram of the frequency of occurrence within a region (or forest) of surface water 

SO4

2-

concentration can reveal the typical distribution of the range of values that may be 

attributable to broad patterns of regional atmospheric inputs versus the more sporadic 

occurrence of high values that more likely derive from geological sources of S. In this 

example, the concentrations of SO4

2- (μeq/L) in lakes in (A) Sierra Nevada, (B) Cascade 

Mountains, (C) Idaho Batholith, (D) NW Wyoming, and (E) Colorado Rocky Mountains 

are examined (Western Lake Survey). If a given study area receives relatively 

homogenous levels of atmospheric S deposition, then it can be assumed that the 

observed outlier high concentrations in lake water represent largely non-atmospheric 

input. (Source: Sullivan 2000.)
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Spatial patterns in water chemistry and/or landscape characteristics can also be used to aid in 

extrapolating results from a relatively few intensively studied sites to the larger region. In many 

cases, model simulations of future chemistry or critical deposition load may be available for only 

a small subset of the lakes or streams in a given forest. Such results can sometimes be 

extrapolated to the wider population of waters using relationships with water chemistry (such as, 

for example, ANC in the example shown in Figure 4-15) and/or landscape features that correlate 

with sensitivity.

Figure 4-15. Critical load simulated by the MAGIC model to protect streams in Shenandoah National 

Park against acidification to ANC below 0 (top panel) and 20 μeq/L (bottom panel) by the 

year 2040 is plotted as a function of 1990 ANC. Stream sites are coded to show

differences in geology. (Source: Sullivan et al. 2008.) This approach yields a predictive 

equation for estimating the model projected value (in this example of critical load) for a 

specific stream based on the measured value of ANC in that stream.
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QUANTIFY LONG-TERM CHANGES IN LAKE OR STREAM CHEMISTRY OVER TIME

Detection of trends in water quality over time can be complicated by analytical error and 

measurement uncertainty that contribute to “scatter” in time series data. In addition, inter- and 

intra-annual variability contribute additional scatter. In particular, seasonal and episodic 

variability (which are largely driven by climate and hydrology) often contribute to short-term 

changes in water chemistry by acidification (Figure 4-16). Therefore, many years (we recommend 

at least eight) of monitoring data may be needed to reveal a probable trend. In general, it is 

helpful to collect data multiple times each year to measure the seasonal variability that exists and 

to discern the trend that exists within the noise of that variability. In other cases, it can be helpful 

to standardize time series data to minimize the influence of hydrological differences in 

constituent concentrations; the purpose in this is to minimize or eliminate the variability 

associated with seasonality in the data. This can be accomplished in multiple ways, including:

• Focusing on summer or fall index chemistry, with collection of samples under conditions 

having minimal influence of snowmelt or rainstorm events;

• Representing the available data as discharge-weighted average values if discharge values are 

available for the site in question or can be estimated or indexed from a nearby site that is 

gaged; or 

• Focusing only on the minimum (i.e., ANC) or maximum (i.e., Ali) concentration measured 

during each year; in this case, the plot would be of the lowest of multiple measured ANC 

values for each year over a period of 8 or more years. 
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Figure 4-16. Example trends analyses of ANC in lake and stream waters, based on data from EPA’s 

Long Term Monitoring program. (Source: Sullivan 2000.) These examples illustrate that 

inter annual and intra annual variability can sometimes be larger than the change over 

time in the variable of interest.
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DETERMINE TO WHAT EXTENT AIR POLLUTION IS CURRENTLY AFFECTING THE 

WATER RESOURCES IN A FOREST OR WILDERNESS

There are many ways to analyze data to shed light on this question. In general, no single approach 

should be considered definitive. When multiple approaches converge to provide similar 

conclusions, there is greater confidence in the validity of that conclusion. Some of the figures 

presented previously (e.g., Figures 4-7 through 4-12) can provide useful information. Another 

approach entails plotting the relationships between the ratio of SO4

2- (or SO42- + NO3-

) to the SBC

concentrations versus ANC (Figure 4-17). A clear pattern across sites of decreasing ANC as the 

ratio of mineral acid anion (presumed to have been derived from acidic deposition) to base cation 

increases (reflective of ecosystem acid buffering) suggests that ANC has decreased in response to 

acidic deposition. The concentration of DOC can alter the relationship by decreasing ANC below 

what would otherwise be expected at a given SO4

2-

/SBC ratio. This is clearly evident in the top 

panel (Adirondack mountain lakes). Streams in the Catskill Mountains (middle panel) tend to 

have uniformly low DOC, and so subsetting on DOC concentration is not necessary. Because 

NO3

-

concentration is relatively high at some Catskill stream sites, the ratio includes NO3

-

in the 

middle panel and is presented for that region as [SO4

2-

+ NO3

-

]:SBC. In the Cascade mountain 

lakes (bottom panel) there is little evidence of a regional acidification signal.

Assessments of acidic deposition effects and recovery generally rely on ANC and pH as the 

primary chemical indicators. However, both of these measurements can be influenced by 

naturally produced organic acidity associated with DOC, which can be abundant in streams and 

lakes draining wetlands and coniferous forests. In waters with significant concentrations of DOC, 

acidity from acidic deposition can be distinguished from natural organic acidity using the basecation surplus (BCS). The BCS is an index that is based on the mobilization of toxic inorganic 

aluminum within the soil. In the absence of acidic deposition or geological S, inorganic Al 

remains in the soil in a non-harmful form. However, acidic deposition dissolves soil Al in a form 

that moves from soils into surface waters and harms both terrestrial (Minocha et al. 1997, Long et 

al. 2009) and aquatic life (Baldigo et al. 2007 and 2009 and Lawrence et al. 2008a). A BCS value 

less than 0 μeq/L in surface water generally indicates that the soil has become sufficiently 

acidified by acidic deposition to enable toxic forms of aluminum to be mobilized (Lawrence et al. 

2007, Lawrence et al. 2008b). A negative BCS value could also occur from acid mine drainage or 

where drainage waters pass through geologic deposits rich in sulfide-bearing minerals. A BCS 

value between 0 and 50 μeq/L indicates a watershed with low calcium availability, which is at 

risk of future acidification from continued acidic deposition and can limit the productivity of 

aquatic (Jeziorski et al. 2009) and terrestrial (Long et al. 2009) ecosystems. 

The BCS can be calculated using variables typically measured in low-ionic strength waters at risk 

from acidic deposition: 

BCS = (Ca2+ + Mg2+ + Na+ + K+) – (SO4

2− + NO3− + Cl− + RCOO−

s) (12)

RCOO−

s = 0.071(DOC) – 2.1 (13)

where all concentrations used in Equation 12 are in μeq/L and the concentration of DOC in 

Equation 13 is expressed in μmol/L.
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Figure 4-17. Plot of ANC versus ion ratio of SO4

2- concentration divided by the sum of base cation 

concentrations (SBC) for low ANC (≤ 200 μeq/L) surface waters in (a) Adirondacks, (b) 

Catskill Mountains, and (c) Cascade Mountains. The base cation sum includes Ca2+ + 

Mg2+ + Na+ + K+. This plot reveals that surface water ANC is strongly related to the ratio 

SO4

2-

/SBC in the Adirondack region, and the ratio (SO4

2-

+ NO3

-

)/SBC in the Catskill 

region, but weakly related to the ratio SO4

2-

/SBC in the Cascade Mountains region. Such 

an analysis sheds light on factors controlling surface water ANC. (Source: Sullivan 

1990.)

Perhaps the most straight-forward way to determine whether or not and to what extent a given 

lake or stream has acidified is to construct a model hindcast of past water chemistry. The two 

models most commonly used for such purpose are MAGIC (Cosby et al. 1985) and PnET-BGC 

(Gbondo-Tugbawa et al. 2001). Each of these models has been widely used across the United 

States to model watershed acid-base chemistry, including hindcasts of past chemistry, forecasts of 
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125

future chemistry under differing future deposition rates, and to estimate critical loads of 

deposition to provide resource protection or to allow damaged resources to recover. 

In addition, paleolimnological reconstructions of past lakewater chemistry can be constructed 

from the fossil remains of algal diatoms or chrysophytes (cf. Charles et al. 1990a and 1990b) in 

dated lake sediment cores. If both process model hindcast simulations and paleolimnological 

reconstructions suggest past acidification (especially if the estimates of change are quantitatively 

similar), then there is increased confidence in the validity of that conclusion. 

It can be helpful to discern what types of lakes and/or streams within a forest have the lowest 

ANC, highest NO3

- concentration, highest Ali concentration, etc. Such kinds of waters become 

potentially important sites for further study or enhanced protection. For example, it may be that 

the lakes or streams in a forest having particularly low ANC may be generally, or entirely, small. 

In the Adirondack Mountain region, lake ANC is related to lake area (Figure 4-18). In this 

example, small lakes are more likely to be both low and high in ANC; intermediate-sized lakes 

tend to have more intermediate chemistry. Few lakes larger than 20 ha are acidic in this region. 

Similarly, it can be helpful to examine relationships between lake or stream chemistry and other 

morphometric features of the landscape, such as watershed area, stream order, lake depth, 

watershed slope, etc. 

Figure 4-18. Relationship between lake size and lake ANC in the Adirondack Mountains. (Source: 

Sullivan et al. 1990.)

Across a given region, the leaching loss of NO3

- in drainage water, which can be expressed as 

NO3

- (sometimes including also NH4+

) outputs in units of mass of N per unit watershed area per 

year, is related to N inputs in deposition (expressed in the same units). For example, this was 

shown for research sites across northern Europe (Figure 4-19). In this example, N output was 
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126

very low at N deposition levels less than about 9 kg N/ha/yr. Here, N leaching became 

pronounced for some, but not all, sites at N deposition greater than 9 kg/ha/yr. In addition, N 

leaching became consistently high at N deposition greater than 25 kg/ha/yr. Within a given forest 

or FS region, it might be possible to use this analysis approach to identify at what level of N 

deposition leaching of N to stream or lake water becomes pronounced.

Figure 4-19. Nitrogen outputs in soil water or stream water versus N deposition inputs throughout 

Europe. (Source: Dise and Wright 1995.)

Nitrogen leaching is not always governed entirely, or even mainly, by N deposition. Other 

factors, especially climatic factors, can also be important. For example, Moldan and Wright 

(1998) showed a strong relationship between N leaching and air temperature at a research site in 

Sweden (Figure 4-20). This analysis suggests that N dynamics at this research site might be 

strongly controlled by climatic condition, in this case air temperature. Precipitation and/or 

snowpack condition could similarly be important.
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127

Figure 4-20. Observed relationship between NO3

- leaching loss in runoff and mean air temperature at 

an experimental watershed site at Gårdsjön, Sweden. Each point represents an average 

of data collected over a period of 14 to 90 days. (Source: Moldan and Wright 1998.)

Nitrogen saturation of aquatic ecosystems has been described in stages, from Stage 0, which 

reflects relatively pristine, unimpacted conditions, to Stage 3, which reflects advanced N 

saturation (Stoddard 1994). Seasonal surface water nitrate concentration peaks at Stage 0 are 

generally rather low (less than about 25 μeq/L) and of relatively short duration (Figure 4-21). At 

Stage 1, the peaks in surface water NO3

- concentration are higher and the period of elevated NO3-

concentration is more extensive. Under Stage 2 N saturation, NO3

- concentrations remain elevated 

throughout the annual cycle but some seasonality is still evident. At Stage 3, the N output may 

actually be greater than the N input (when expressed as mass per unit area per year). The 

temporal pattern of NO3

- concentration in a given lake or stream can indicate the probable stage 

of N saturation of the watershed. For a given stream or lake, this type of analysis can illustrate the 

stage of N saturation of that water body as well as its drainage area.
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Figure 4-21. Example patterns of NO3

-

concentration in surface water at four sites at various stages of 

watershed N-saturation. (Source: Stoddard 1994.)

EVALUATE WHETHER THE CURRENT CONDITION OF ACID- OR NUTRIENTSENSITIVE WATERS WARRANTS MITIGATION 

There is no standard analysis approach that will answer this question. This is a management 

judgment that should be based on a variety of analyses, as outlined above. Potential mitigation 

strategies can include imposing tighter controls on atmospheric emissions of S and/or N and 

adding base cations to waters or watersheds by liming. Virtually all of the approaches suggested 

above can contribute to such decision-making.

4.5 CONDUCT STATISTICAL ANALYSES 

4.5.1 Statistical Tests for Difference

It should not be automatically assumed that formal statistical tests are needed in analyzing a 

dataset. Much can be gained by conducting routine exploratory data analyses, such as those 

outlined in the previous section, without adding the complexity of formal statistical tests. In many 

cases, the assistance of a statistician or other person who is quite knowledgeable about statistics 

will be needed for conducting such tests. 

Statistical tests are often used to determine the existence of significant differences between 

groups of sites or samples. The most common tests are parametric, and include t-tests and 

analysis of variance (ANOVA) that use means and variances to determine significant differences 

among group means. These parametric tests, however, make assumptions about data normality 

and independence that need to be examined before using them. Data are normally distributed if 

the various concentrations measured at different times for the same site or at different sites are 

bell-shaped (Figure 4-22). If the data are not normally distributed, they must be transformed 
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before parametric analysis or they must be analyzed using a non-parametric test. For water 

chemistry, variables are often log-transformed to achieve a normal distribution. There are a 

number of statistics that can test for normality (e.g., the Kolmogorov-Smirnov test), but we 

recommend plotting histograms or some other type of frequency plot and visually inspecting the 

graph to see if the distribution deviates grossly from a bell-shaped normal distribution.

Figure 4-22. Schematic representation of data normality.

Non-parametric or distribution-free statistics are used to test for group differences in skewed (notnormally-distributed) data or when the analyst is not comfortable with assumptions about 

normality. Water chemistry data are commonly not normally distributed. The non-parametric 

tests are based on sample ranks (rank order number from low to high) and not actual data values. 

Because the test is based on rank, no assumption is made about the underlying data distribution. 

The best known non-parametric test for group difference is the Mann-Whitney U test, or as it is 

also known, the Wilcoxon rank-sum test. 

Non-parametric tests, including the rank-sum test, tell you nothing about the magnitude of the 

difference between groups: just whether the group differences are significant. Analysts should be 

cautioned, however, that with large enough sample sizes, groups can be statistically different but 

such differences may have little ecological significance. The magnitude of any differences needs

to be examined; running a statistical test to determine if differences are statistically significant is 

not sufficient. 

There are multiple forms of the rank-sum test, and which form to use is a complex choice (Helsel 

and Hirsch 1992). We recommend that this test not be applied by persons lacking formal training 

in statistics. The rank-sum test determines whether one group of measurements tends to produce 

higher values than another group of measurements. In other words, the test determines if both 

groups of data are from the same population. The groups might represent different lake types, 

different periods of time, different seasons, etc. 

For comparing more than two independent groups of data points, the Kruskal-Wallis test is often 

used. It can be computed by an exact method used for small sample sizes (typically five or fewer 

samples per group), a large-sample (chi-square) approximation, or by ranking the data and 

performing a parametric test on the resulting ranks (Helsel and Hirsch 1992). The latter two 
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methods only produce valid p-values when sample sizes are large. The null hypothesis for all 

variations of this test specifies that all of the groups have identical data distributions (or have the 

same median value); the alternate hypothesis specifies that at least one group differs from the 

others with respect to its data distribution or its median value. 

As in a rank-sum test, all observations are combined and ranked from lowest (1) to highest (N). 

The average group rank ( ) is compared to the overall average rank to calculate the test statistic. 

And like the-rank sum test, we recommend that this test should be conducted by someone who 

has had formal training in statistics.

4.5.2 Trend Detection

Trends in water chemistry over time can be evaluated using simple linear regression (SLR) or 

using a more sophisticated statistical approach such as the non-parametric seasonal Kendall tau 

test (SKT; Hirsch and Slack 1984) for determining monotonic trends in seasonally varying water 

quality. The SLR approach is simpler to apply and will sometimes yield nearly identical estimates 

of slope as the SKT (Sullivan et al. 2003). An example SLR analysis for SO4

2- concentration in 

Deep Run, a small acid-sensitive stream in Shenandoah National Park, is shown in Figure 4-23. 

The slope of the regression (-0.57 μeq/yr) indicates the rate of change (in this case a decrease) in 

the variable (SO4

2-

) over time. Whether or not the relationship between SO4

2- concentration and 

time is statistically significant can be determined using the p-statistic. We recommend a p-value 

of less than or equal to 0.05 as the benchmark for determining statistical significance. The r

2

-

statistic can be used to determine the percent of the variation in SO4

2- concentration that is 

explained by the variable time.

Figure 4-23. Plot, with regression line, of SO4

2- concentration in Deep Run, Shenandoah National 

Park, over the period of monitoring record. (Source: Sullivan et al. 2003.)
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If the regression relationship is statistically significant at p ≤ 0.05, the next step is to conduct a 

test to determine whether the slope of that relationship is significantly different from zero. If it is 

determined that the slope is statistically either greater or less than zero, then it can be concluded 

that the parameter in question is truly increasing or decreasing over time. 

The test statistic for determine if the slope is different from zero is expressed as:

(14)

where r is the correlation coefficient of the regression and n is the number of data points. 

The null hypothesis that the slope of the regression equals zero is rejected if the absolute value of 

t is higher than tcrit, where tcrit is the point on the Student’s t-distribution with (n-2) degrees of 

freedom and with a probability of exceedance of α/2. The Student’s t-distribution for that 

confidence level is given in Table 4-7. 

Table 4-7. Values of t. (Source: Steel and Torrie 1980.)

df Probability (α=0.05) of a 

numerically larger value of t

df Probability (α=0.05) of a 

numerically larger value of t

1 12.706 18 2.101

2 4.303 19 2.093

3 3.182 20 2.086

4 2776 21 2.080

5 2.571 22 2.074

6 2.447 23 20.69

7 2.365 24 2.064

8 2.306 25 2.060

9 2.262 26 2.056

10 2.228 27 2.052

11 2.201 28 2.048

12 2.179 29 2.045

13 2.160 30 2.042

14 2.145 40 2.021

15 2.131 60 2.000

16 2.120 120 1.980

17 2.110 ∞ 1.960

Probability (α/2=0.025) of a larger positive value of t

In general, we recommend that FS staff conduct SLR analyses as the routine approach to 

determine changes in water chemistry over time. If it is deemed necessary to obtain a more 

rigorous trends estimate, then a more complex statistical analysis can be conducted. 

In some cases, there may be a visually obvious change in the slope of the data points in the midst 

of the time series. For example, some water bodies in the United States experienced acidification 
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during the 1980s and 1990s, but recovery (decreasing ANC) is evident after about the late 1990s. 

In such a situation, it can be helpful to visually split the data into two time periods (acidification 

and recovery) and perform a SLR separately on each time period to determine if there has been a 

change over time. 

Simple linear regression analysis is a good first approach for analyzing temporal data and can be 

performed using common spreadsheet and statistical software. However, regression analysis is 

sensitive to data normality and other assumptions and is often not the most robust method to 

quantify the statistical significance of temporal trends. Loftis et al. (1989) evaluated a number of 

different trend detection methods under a number of different conditions and found that there is 

no one method that outperforms the others under all conditions. However, they found that the 

most powerful methods under most conditions were non-parametric tests that looked at the 

correlation between rank order and time. For annual data, the Kendall-tau (also called MannKendall test for trend) test was generally the most powerful. For seasonal data, the SKT or 

seasonal analysis of covariance (ANCOVA) on ranks were the most powerful tests. These nonparametric tests performed about as well as parametric tests with normal data and outperformed 

them when the data were non-normal. We recommend their use for testing the statistical 

significance of temporal trends when a more rigorous statistical application than SLR is desired. 

Note that these tests are the most powerful for testing whether or not water quality is changing 

over time; they do not, however, quantify the magnitude of change.

Trend detection in surface waters can be conducted using complex statistical tests that attempt to 

adjust for natural variation related to seasonality and variations in discharge. Different methods 

have been developed for assessing trends or the effect of time, with proper method selection 

dependent on assumptions related to the distribution and independence of the data and on whether 

change occurs monotonically or as a step change. It is also possible to construct complicated 

models that incorporate flow, temperature, and/or other environmental factors in addition to the 

time variable to quantify trends. These regression-based approaches, however, are sensitive to 

issues of data normality and independence. In addition, different methods are available for 

assessing trends for individual streams or lakes, as well as for assessing regional trends associated 

with classes or populations of streams and lakes. These methods are described by Helsel and 

Hirsch (1992), Stoddard et al. (2003), and Irwin (2008). In general, we do not recommend 

application of such tests for routine water chemistry assessment in the FS ARM program. 

If a more complicated test is to be conducted, trends in time-series data collected at quarterly, 

monthly, or weekly intervals for individual surface waters are most commonly assessed using the 

SKT developed and described by Helsel and Hirsch (1992). The non-parametric SKT is based on 

the correlation between the ranks of the dependent variable (concentration) and an evenly spaced 

time interval. The SKT is popular because of its relative simplicity compared to other approaches 

and minimal data assumptions. It is appropriate for data showing seasonal cycles, and it is robust 

with respect to issues of normality, missing or censored data, and serial correlation. It can be 

applied on unadjusted chemical concentration data or on residuals from ordinary least-squares 

regression of concentration on estimated discharge, thus accounting for the effects of changes in 

discharge. An alternative approach is to remove the seasonality from the dataset (for example by 

subsetting to only include samples collected during summer base-flow) and then analyzing the 

subsetted data as an annual, rather than seasonal, dataset. The SKT provides the significance and 
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direction of any trend. A different test based on the median of the set of slopes calculated for all 

possible pairs of points in the time series is commonly applied to calculate the slope or rate of 

change associated with the overall trend (Sen 1968).

Regional trends associated with classes or populations of surface waters can be assessed using a 

median trend test (SAS 1988, Altman et al. 2000, Stoddard et al. 2003). This test, a meta-analysis, 

is based on the median of slopes obtained for linear regression of concentrations with time for the 

individual surface waters in the class or population of concern. Regional trend significance is 

tested by estimating confidence intervals around the median values, with median slopes 

significantly different from zero taken to indicate regional increasing trends (positive median 

slope) or regional decreasing trends (negative median slope). This test allows determination of

trends for a resource management or geographic unit as a whole. It can be applied to include 

multiple predictor variables in the regression models, thus accounting for other factors, such as 

discharge, in addition to time.

4.5.3 Statistical Power

For purposes of inferring differences or change, data analysts and resource managers will have to 

make decisions concerning acceptable error levels, and such decisions should be based on the 

allocation of risk, given the relative importance of ensuring resource protection compared to the 

cost of potentially unnecessary responses. These decisions are typically made in terms of 

statistical power and significance. Statistical power refers to the avoidance of false negatives, or 

wrongly concluding that a change or impact has not occurred when, in fact, a change or impact 

has occurred (type II error). Statistical significance refers to the avoidance of false positives, or 

wrongly concluding that a change or impact has occurred when, in fact, no change or impact has 

occurred (type I error). 

Statistical power and significance levels are stated as percentages. A statistical power level of 

90%, for example, would mean that, 90% of the time, an effect of a specified size (whatever it is) 

will be correctly identified. A statistical significance level of 5%, for example, would mean that 

only 5% of the time would an identified effect be incorrectly identified (or that there is a 95% 

probability that the identified effect is, in fact, correctly identified). Ideally, statistical power and 

significance objectives are established as part of the initial study or monitoring program design 

and not during data analysis. They will be stated in terms of the project’s ability to detect a 

specific effect or a specified trend (a magnitude of change within a specified time period). 

The ability of a monitoring program to detect temporal trends is a function of a number of 

different factors. For detecting a trend at a single site, for a specified type I and type II error rate 

(for example, a 95% confidence level), trend detection is determined by the magnitude of the 

actual trend you wish to detect, how long you have to detect the trend, and the variability of the 

water quality parameter being assessed. For water chemistry assessment, trend magnitude is 

usually expressed as a percent change in the variable of interest per year. Evaluation of the 

magnitude of the trend one wishes to be able to document is usually based on program objectives 

related to ecologically significant changes in water quality values within a time period of policy 

relevance. There is no standard procedure for this, but in very general terms, one typically wishes 

to be able to document, for acid-base chemistry monitoring, a change in ANC of at least 1-2 

μeq/L/yr over a period of about 10 years. 
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The ability to detect a trend is also related to how long a monitoring program is continued. Small 

trends that are not detectable with 5 years of data can be obvious in 100 years. It requires a very 

large trend magnitude for a trend to be detectable in a short amount of time. Trend detection is 

dependent on water-quality parameter variability both in terms of analytical precision and natural 

(e.g., climatic) temporal variability. For a given trend magnitude of interest, it will take longer to 

detect a trend in a noisy, highly varying indicator than in a more precise and temporally stable 

indicator. Similarly, to detect a trend over a fixed amount of time, smaller trend magnitudes can 

be detected in stable indicators more readily than in noisy indicators. 

For detecting regional trends (average trend across a number of sites), the number of regional 

sample sites is also an important factor. Regional trend detection ability increases with the 

number of sample sites. Thus, you can enhance your ability to document a trend by: 1) 

monitoring over a longer time period, 2) reducing short-term variability due to seasonality, 

episodes, or data quality, and/or 3) monitoring more sites. Larger trends will be easier to 

document than smaller trends. 

4.6 REPORT DATA IN STANDARDIZED FORMATS

Data is managed in NRM Air. Specific requirements for incorporation into NRM Air are detailed 

in the NRM Air User Guide available at 

<http://fsweb.nris.fs.fed.us/products/air/documentation.shtml>.

4.7 SUMMARY AND CONCLUSIONS

It is not possible to specify a routine set of data analyses that should be conducted for every water 

chemistry data set assembled by the FS ARM program. Decisions regarding analysis of the data 

will be influenced by the distribution of the collected data and the objectives of the particular 

investigation. Nevertheless, we do suggest many of the kinds of analyses that should be 

considered for the datasets typically collected within the FS ARM program.

The first step in analyzing a water chemistry dataset is to evaluate the overall quality of the data. 

In the process of evaluating data quality, it is often possible to identify data that are incorrect and 

sometimes to reanalyze or otherwise correct the identified errors. Next is a series of steps to 

prepare the raw data for graphical and statistical analysis. This involves applying procedures to 

deal with such potentially confounding issues as censored data, outliers, missing values, multiple 

observations, and treatment of zeros and negative values. Recommendations are provided. 

A range of kinds of exploratory data analyses are illustrated with examples from the published 

literature. These include suggestions regarding how to subset the data in order to increase data 

analysis efficiency. Specific analyses suited to various study objectives are provided as examples. 

Finally, the role of formal statistical analysis is considered. Frequently, such analyses will require 

the assistance of someone with formal training in statistics. In many cases, however, formal 

statistical analyses are not required. Much can be gained via routine exploratory analyses and 

application of simple graphical and analytical procedures.
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SECTION 5. 

FIELD SAMPLING PROTOCOLS 

FOR AQUATIC BIOTA A.T. Herlihy and T.J. Sullivan

5.1 BACKGROUND

Aquatic invertebrates can be good indicators of water quality and can provide documentation for

ecological effects of changing water quality. Bottom dwelling (benthic) invertebrates have been 

used extensively to assess biological conditions in streams. Benthic macroinvertebrates can also 

be used in assessing lake biology, but their use for this purpose has not been common in the 

United States. More commonly in this country, biological conditions in the epilimnion of 

thermally stratified lakes are evaluated using zooplankton. Both stream macroinvertebrate and 

lake zooplankton data can provide useful information to reveal some of the ecological effects that 

result from atmospheric deposition, and subsequent alterations of surface water chemistry. 

5.1.1 Lake Zooplankton

Zooplankton are important components of the biological community of lakes. There may be as 

many as 200 species or more that occur within lakes in a given region. They constitute key 

portions of the aquatic food web, and play a major role in transferring energy from the primary 

producers (mainly phytoplankton) to predatory invertebrates and to fish and other vertebrates. 

Individual zooplankton species and the zooplankton community as a whole respond to a number 

of environmental stressors. These include acidification, nutrient enrichment, sedimentation, fish 

stocking, and habitat manipulation. Effects of these environmental stressors can sometimes be 

revealed by evaluating changes in the presence/absence of known regional indicator species, 

overall species composition, biomass, body size distribution, and/or the structure of the food web. 

Lake zooplankton include crustaceans, rotifers, pelagic insect larvae, and aquatic mites. Many 

species, especially crustaceans and rotifers, are known to be sensitive to changes in chemistry (cf. 

Gerritsen et al. 1998, Melack et al. 1989, Sullivan et al. 2006). Nevertheless, the species 

composition and trophic structure of zooplankton communities are controlled by multiple factors, 

of which aquatic acid-base chemistry is only one. Populations of zooplankton can be strongly 

influenced by changes at both lower and higher trophic levels because zooplankton are sensitive 

to changes in the distribution and abundance of both algae and predators. Predation occurs by 

planktonic predators and by fish. Thus, the presence or absence of plankton-feeding fish can have 
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a large influence on the presence, abundance, and body size of various species of zooplankton. It 

can therefore be difficult to infer the cause(s) of observed changes in the zooplankton community 

over time unless data are also available regarding the status of the fish populations in the lake(s) 

under study. Therefore, if fish population data are available for lakes within a given forest or 

wilderness, it can be helpful to select lakes for zooplankton sampling that also have data on fish. 

In addition, both intra- and inter-annual variability in zooplankton species distributions can be 

high. In general, the greatest development of zooplankton occurs from June to mid-October, and 

mid-summer is considered to be a relatively stable period for zooplankton monitoring. 

5.1.2 Stream Macroinvertebrates

Benthic macroinvertebrates inhabit the bottom substrates of streams and provide a good 

indication of overall biological condition (Kerans and Karr 1994, Barbour et al. 1999, 

Reynoldson et al. 2001, Klemm et al. 2002 and 2003, Clarke et al. 2003, Bailey et al. 2004, 

Griffith et al. 2005). Monitoring these assemblages is useful in assessing the status of the water 

body and investigating the possibility of trends over time in ecological condition. Benthic 

macroinvertebrate species respond to a variety of stressors in different ways, and it is often 

possible to determine the type of stress that has affected the macroinvertebrate assemblage (e.g., 

Klemm et al. 1990). 

Because many stream macroinvertebrates have life cycles of a year or more and are relatively 

immobile, macroinvertebrate assemblage structure is a function of present and past conditions 

and provides an integration of the variability that typically occurs in stream condition with season 

and with changing hydrology (Barbour et al. 1999, Peck et al. 2006). For general bioassessment 

purposes, stream macroinvertebrates are typically sampled in summer. However, for assessing 

acidic deposition impacts, we generally recommend sampling during spring base-flow as that is 

the season of maximum impact (lowest ANC/pH). At high elevation in regions that experience 

substantial snowpack development, spring sampling may not be feasible; at such locations, 

summer sampling is recommended. 

The insect order Ephemeroptera (mayflies) is an excellent indicator taxa for acidification effects. 

However, it comprises just one order of benthic invertebrates, and there are many other 

taxonomic groups that make up the benthic stream community that should be considered for a full 

biological assessment. These other taxa also contribute a great deal of information about stream 

condition. In addition to acidification impacts, macroinvertebrates are excellent indicators of 

substrate alteration (e.g., sedimentation), nutrient enrichment, metal pollution, and habitat 

alteration. In the EPA National Wadeable Stream Assessment, the greatest risk of poor stream 

macroinvertebrate condition was found in streams with excess sediment (enhanced erosion) 

and/or with nutrient enrichment (Van Sickle and Paulsen 2008). 

Some studies have found that acidified streams host fewer invertebrate taxa than streams with 

higher acid neutralizing capacity (ANC) and pH (e.g., Feldman and Connor 1992, Kaufmann et 

al. 1999, Sullivan et al. 2003). This is especially true for mayflies (order Ephemeroptera) and, to a 

lesser extent, caddisflies (order Tricoptera) and stoneflies (order Plecoptera). Aquatic insect status 

is sometimes evaluated on the basis of these three orders using what is known as the EPT 

(Ephemeroptera-Plecoptera-Tricoptera) Index (EPT taxa richness). 
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5.2 STUDY DESIGN

Aquatic invertebrates can be collected and analyzed as part of lake or stream characterization 

studies, synoptic surveys, long-term monitoring, or used to augment model projections of future 

chemical conditions. As is described in detail in the Water Chemistry Field Sampling Protocols 

(Section 1), the design of an aquatic biota study should be a function of the study purpose and 

questions being asked. Some example approaches for biological characterization or monitoring 

are outlined in Table 5-1. Each example approach is tied to a specific purpose. The reader is also 

referred to the discussion with examples, provided in Section 1.1.2, Study Purpose and 

Objectives.

Table 5-1. Example approaches for FS ARM program biological sampling, tied to the purpose of 

the field study.

Purpose Approach

1. Determine spatial patterns in biological 

assemblages relative to chemical 

and/or deposition gradients.

a. Conduct survey of lake zooplankton or stream benthic macroinvertebrate 

communities in waters that exhibit varying water chemistry and/or that receive 

varying atmospheric deposition levels.

2. Quantify long-term changes over time 

in biology.

a. Conduct trends analysis in species richness.

b. Compare trends in biota with trends in water chemistry.

3. Determine extent to which air pollution 

is affecting water resources.

a. Characterize biology of multiple lakes or streams expected to be sensitive.

b. Plot changes in species richness versus changes in ANC or NO3

-

concentration.

Evaluation of the status of the aquatic invertebrate biota can be used, along with assessment of 

chemical status and/or change, to estimate the impacts of nutrient enrichment, acidification, and 

various kinds of habitat disturbances. In general, changes in aquatic chemistry are more easily 

documented than are changes in aquatic biology. Nevertheless, the concerns on the part of land 

managers and the public regarding changes in chemistry are fundamentally rooted in widespread 

concerns about protecting resources against biological damage. The ultimate purpose of studying 

or monitoring aquatic chemistry on federal lands is often mainly to aid in the protection of 

biological resources. Also, biota reflect conditions over longer time periods than the single point 

in time represented by a water sample. Therefore, there is additional power in the inclusion of a 

biological component in the investigation or monitoring of chemical conditions. 

For evaluation of biological responses to acidification, study designs most commonly include 1) 

documentation of relationships between water chemistry (usually ANC; also pH, Ali, or NO3

-

concentration) and macroinvertebrate taxonomic composition across sites (lakes or stream 

reaches) within a reasonably small area (i.e., National Forest or wilderness), or 2) evaluation of 

changes over time in water chemistry and macroinvertebrate taxonomic composition. Such 

studies often focus on stream aquatic insects in the orders Ephemeroptera (mayflies), Tricoptera 

(caddisflies), and Plecoptera (stoneflies). For a more rapid and less expensive stream assessment, 

the analysis can be restricted to only the order Ephemeroptera, which is the order most 

susceptible to acidity. However there may be no mayfly species present if the ANC in a given 

study stream is especially low (near or below zero). For lakes, acidification studies typically focus 

on zooplankton, mainly crustaceans and rotifers. We recommend, if available funding allows, that 
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138

such studies be conducted on a suite of acid-sensitive lakes or streams within a given National 

Forest or region, if lake or stream acidification is believed to be an important issue. If, after a 

prolonged period of monitoring, a trend is indeed documented, a decision will have to be made 

regarding whether or not to continue monitoring. In general, we recommend continued 

monitoring. Data collected will help to identify long-term and future interactions between climate 

change and air pollution effects. 

For a more complete assessment of biological condition, we recommend use of an Index of Biotic 

Integrity (IBI). Such an index provides a more complete assessment of biological condition than 

the rapid single order or EPT assessments discussed above. An IBI can provide a more rigorous 

assessment of biological conditions and response to multiple stressors where in-depth study is 

warranted as it requires compiling available information about the feeding groups, pollution 

tolerance, and habits of the taxa in the study waters. It is somewhat more expensive, however, and 

requires more specialized taxonomic and autoecological (individual species ecology) expertise.

For this reason, we do not recommend implementation of an IBI as a routine procedure on FS 

lands.

5.3 SITE SELECTION

In general, criteria for site selection for the purpose of conducting a biological assessment are the 

same as criteria for site selection for the purpose of investigating aquatic chemistry: random 

versus non-random site selection, and the establishment and documentation of the stream 

reach(es) and/or lake(s) to sample. See the discussion in Section 1.3 in the Water Chemistry Field 

Sampling Protocols for a more complete discussion of these issues. 

By necessity in many cases, the sites included in a biological study will be only a subset of the 

sites included in the chemical investigation. It is important to choose this subset wisely. In 

general, one may wish to avoid sites having substantial disturbance other than atmospheric 

deposition that may influence the acid-base chemistry or nutrient status of drainage waters (i.e., 

geological sulfur, forest fire, insect infestation, tree disease, large windthrow, or other substantial 

disturbance). In general, one should include biological characterization for monitoring sites that 

are expected to be sensitive to the stresses of interest. For acidification studies, these are usually 

the lakes or streams having ANC less than about 50 to 100 μeq/L. For nutrient enrichment 

studies, these are usually N-limited water bodies. Short of direct experimentation, it is difficult to 

predict which water bodies might be N-limited. The Redfield ratio, based on the molar N:P ratio 

in phytoplankton, suggested that water bodies might be N-limited if the molar N:P ratio was less 

than 16. Subsequently, experimental studies suggested higher cutoffs. Recent research suggests 

that N-limited lakes generally include lakes having the molar ratio of N:P less than about 44 

(Guildford and Hecky 2000, Schindler et al. 2008, Elser et al. 2009). 

5.3.1 Laying Out the Support Reach for Stream 

Macroinvertebrate Sampling 

Unlike chemistry, which can be measured at one point, characterizing stream biota requires 

sampling a length of a stream to capture the range of available habitat. There are a large number 

of field protocols for sampling stream macroinvertebrates for bioassessment, and they all specify 
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139

collecting a number of different net samples (kick, Hess, or Surber) from different places along 

the stream sample reach and compositing them into either a single composite sample or habitat 

type (e.g., riffle) composite sample. The procedures that we recommend (summarized here) are 

based on the procedures developed by the EPA for the Environmental Monitoring and 

Assessment Program Surface Water (EMAP-SW) sampling program (Peck et al. 2006). These 

protocols have been used in studies of streams across the entire U.S., and they work in a wide 

variety of stream types. They were also designed to be implemented by different types of field 

crews and require a minimum of field decision-making. 

At each selected stream sampling location (called the X-site) the support reach (the length of 

stream to be sampled at the sampling location) must be laid out. The support reach must be 

sufficiently long to represent the biological community being sampled. Based on several studies 

(Robison 1998, Li et al. 2001, Reynolds et al. 2003), a support reach with a length of 40 times the 

average wetted channel width measured near the X-site is sufficient for almost all sampling. The 

support reach is established about the X-site using the procedures described below. Field staff 

should reconnoiter the support reach to make sure it is clear of obstacles that would prohibit 

sampling and data collection activities. Record the channel width used to determine the support 

reach length and identify the support reach length upstream and downstream of the sample-site. 

Figure 5-1 illustrates the principal features of a hypothetical support reach, including the location 

of 11 cross-section transects from which the samples will be collected.

To lay out the support reach (from Peck et al. 2006):

1. Use a surveyor's rod or tape measure to determine the wetted width of the channel at three

to five places considered to be of typical wetted width within approximately five channel 

widths upstream and downstream from the X-site. Average the readings together and round 

to the nearest 0.5 m. If the average width is less than 3.5 m, use 150 m as a minimum 

support reach length. Record this width on the Stream Verification Form.

For channels with interrupted flow, estimate the width based on the unvegetated width of 

the channel (again, with a 150 m minimum).

2. Check the condition of the stream upstream and downstream of the X-site by having one 

team member go upstream and one downstream. Each person goes until he/she has visited 

the candidate sample reach to a distance of 20 times the average channel width in each 

direction (equal to one-half the support reach length, but a minimum of 75 m) determined 

in step 1 from the X-site.

For example, if the support reach length is determined to be 150 m, each person would go 

75 m from the X-site to lay out the reach boundaries.

3. Determine if the support reach needs to be adjusted about the X-site due to confluences 

with higher order streams (downstream), lower order streams (upstream), impoundments 

(lakes, reservoirs, ponds), physical barriers (e.g., falls, cliffs), or because of access 

restrictions to a portion of the initially-determined support reach.

If such a confluence, barrier, or access restriction is present, note the distance and flag the 

confluence, barrier, or the limit of access as the endpoint of the reach. Move the other 

endpoint of the support reach an equivalent distance away from the X-site. The X-site must 
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still be within the support reach after adjustment. The total support reach length does not 

change, but the support reach is no longer centered on the X-site.

Note: If the sampling sites are statistically (randomly) selected, do not slide the support 

reach to avoid man-made obstacles such as bridges, culverts, rip-rap, or channelization, or 

in streams with interrupted flow to obtain more inundated areas to sample. If the sites are 

not statistically selected, it is recommended to avoid sites that are influenced by substantial 

human-caused channel disturbance. 

4. Starting back at the X-site (or the new midpoint of the reach if it had to be adjusted as 

described in step 3), measure a distance of 20 channel widths downstream on one side of 

the stream using a tape measure. Be careful not to “cut corners.” Enter the channel to make 

measurements only when necessary to avoid disturbing the stream channel before sampling 

activities. This endpoint is the downstream end of the support reach, and is flagged as the 

location of transect A.

5. Using the tape measure, measure 1/10 (4 channel widths in big streams or 15 m in small 

streams) of the required stream length upstream from the start point (transect A). Flag this 

spot as the next cross-section or transect (transect B). For transect A, roll one die to 

determine if it is a left (L), center (C), or right (R) sampling point (facing downstream) for 

collecting benthic macroinvertebrate samples. A die roll of 1 or 2 indicates L, 3 or 4 

indicates C, and 5 or 6 indicates R—or use a digital wristwatch and glance at the last digit 

to determine the sampling point (1-3=L, 4-6=C, 7-9=R). Mark L, C, or R on the transect

flagging.

6. Proceed upstream with the tape measure and flag the positions of nine additional transects 

(sequentially labeled “C” through “K”) as you move upstream at intervals equal to 1/10 of 

the reach length. Assign sampling spots to each transect in order as L, C, or R from the first 

random selection.

For example, if the sampling spot assigned to transect A was C, transect B is assigned R, 

transect C is L, transect D is C, etc.

There are some conditions that may require adjusting the end-points of the support reach about 

the X-site (i.e., the support reach will be shifted either upstream or downstream so that the X-site 

will no longer be located at the midpoint of the support reach) to avoid features we do not wish to 

(or physically cannot) sample across. The full length of the support reach should be of the same 

stream order as the X-site. Do not extend the support reach upstream if the stream order decreases 

or downstream if the stream order increases. If you encounter an impoundment such as a lake, 

reservoir, or pond or an impassible barrier (e.g., a waterfall or a cliff) while laying out the support 

reach, adjust the reach such that the barrier is at one end. Adjusting or sliding the support reach 

involves noting the distance of the confluence, barrier, or other restriction from the X-site, 

flagging the confluence, impoundment, or barrier as the endpoint of the reach, and adding the 

distance to the other end of the reach such that the total support reach length remains the same 

though it is no longer centered about the X-site. In cases where you are denied access permission 

to a portion of the support reach, you can adjust the reach to make it entirely accessible; use the 

point of access restriction as the endpoint of the reach.
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Figure 5-1. Equipment, supplies, and sequence for collecting benthic macroinvertebrate samples.

(Source: Peck et al. 2006.)

5.3.2 Lake Selection for Zooplankton Sampling 

Acidification effects on individual species of zooplankton and on the zooplankton community in 

general may occur across a rather wide spectrum of lake pH, ANC, and Ali concentrations. 

Effects are usually observable at ANC values below about 50 to 100 μeq/L and pH below about 

6.0 to 6.5. Such ANC and pH cutoff values generally correspond with Ali concentrations near 2 
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μM. Lakes having pH below 6.0 and/or ANC below about 50 to 100 μeq/L have an increased 

likelihood of having Ali above this general response threshold. Nevertheless, it is possible that 

effects on zooplankton also occur at somewhat higher pH and ANC (and lower Ali) values. 

A complicating factor relates to the influence of lake size and watershed area on lake biology. In 

general, smaller lakes in smaller watersheds are more likely to be lower in pH and ANC and to 

have less diverse zooplankton communities than larger lakes in larger watersheds. To some 

degree (often to a large degree) this relationship is controlled by the effects of lake chemistry on 

biota; but additionally, smaller lakes in smaller watersheds might be expected to have less diverse 

biotic assemblages than larger lakes in larger watersheds as a consequence of their physical 

simplicity and reduced number of available niches. Thus, the often-observed patterns of changing 

zooplankton species composition and taxonomic richness among lakes are likely only partly due 

to water acid-base chemistry. This makes it difficult to tease out the effects of changing water 

chemistry when evaluating changes over time in the biological community. For these and other 

reasons, it is important to give careful consideration to site selection for zooplankton monitoring 

or characterization. It can be helpful to include multiple lakes in the study, selected to cover a 

range of acid-base chemistry and perhaps within rather narrow windows regarding lake and 

watershed areas. In addition, it can be helpful to study the zooplankton communities of lakes that 

are also being studied with regard to their fish and algal communities; this may allow an 

improved opportunity to sort out what may be a multitude of factors that simultaneously influence 

the lake zooplankton community. 

Biological effects are more likely to be observable in lakes that have relatively low ANC (< 50 

μeq/L) and pH (< 6.0). Nevertheless, having lakes in the study with somewhat higher ANC and 

pH is also important, especially for evaluating effects on taxonomic richness or the 

presence/absence of particular indicator species. This will help to make sure that a sufficient 

range of response occurs in the dataset so as to increase the likelihood of being able to document 

what may be a very “noisy” relationship. Further discussion of the interpretation of zooplankton 

data is provided in Section 5.10, Interpretation. It is likely that study of the zooplankton 

community will be less helpful for evaluation of effects related to nutrient N enrichment: such 

effects on zooplankton communities have not been as well documented as the effects from 

acidification.

Zooplankton samples should generally be collected at the lake index location for water chemistry 

sampling. This should be the deepest part of the lake. The index location is described in Section

1.3.1, Where to Sample. It is important to collect the zooplankton tows in the deepest part of the 

lake because, especially in mid- to late-summer, the size of the cold-water hypolimnion can be 

reduced substantially. Missing the deep spot can cause exclusion of individuals occupying the 

cold water stratum, thereby confounding interpretation of the true zooplankton assemblage in the 

lake. 

Zooplankton tows can be compromised by high algal production and/or high DOC because of 

algal or organic particle fouling of the net. This problem can be partially ameliorated by using a 

reducing collar. Alternatively, if fouling is a major problem, zooplankton can be collected in

integrated water column samples using a hose and pump system. 
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5.4 PRE-TRIP PREPARATION

5.4.1 Equipment and Supplies

Table 5-2 shows the checklist of equipment and supplies required to complete the collection of 

benthic macroinvertebrates from streams. Use this checklist to ensure that equipment and supplies 

are organized and available at the stream site in order to conduct the activities efficiently. 

Similarly, Table 5-3 provides the checklist for zooplankton sampling. These checklists can also 

be found in Appendix H.

Table 5-2. Equipment and supplies for benthic macroinvertebrates. (Source: Peck et al. 2006.)

Quantity Item 

1 Modified kick net (D-frame with 500 μm mesh) and 4-ft handle

Spare net(s) and/or spare bucket assembly for end of net

1 Watch with timer or a stopwatch

2 Buckets, plastic, 8- to 10-qt capacity (collapsible for back country)

1 Sieve with 500 μm mesh openings or sieve-bottomed bucket, 500 μm mesh openings

2 pr. Watchmakers’ forceps (straight and curved)

1 Wash bottle, 1-L capacity, labeled STREAM WATER

1 Small spatula, spoon, or scoop to transfer sample

1 Funnel with large bore spout

4 to 6

Each

Sample jars, HDPE plastic, with leakproof screw caps, 500-ml and/or 1-L capacity, suitable for use 

with ethanol

2 gal 95% ethanol in a proper container (smaller amounts can be carried in for back country work or 

ethanol can be added to sample containers at the vehicle after returning from the field)

2 pr. Rubber gloves suitable for use with ethanol

1 Cooler (with suitable absorbent material) for transporting ethanol and samples in the vehicle

2 Preprinted benthic sample labels with sample ID numbers

4 Preprinted benthic sample labels without sample ID numbers

6 Blank labels on waterproof paper for placing inside of jars

1 Sample Collection Form for the site(s)

Soft (#2) lead pencils

Fine-tip indelible markers

1 pkg. Clear tape strips

4 rolls Plastic electrical tape

1 Knife, pocket, with at least two blades

1 Scissors

1 Pocket-sized field notebook (optional)

1 pkg. Kim wipes in small resealable plastic bag

1 copy Field operations and methods manual

1 set Laminated sheets of procedure tables and/or quick reference guides for collecting benthic 

macroinvertebrates
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Table 5-3. Equipment and supplies for collecting zooplankton samples. (Source: U.S. EPA 2007.)

Quatity1 Item 

2 Wisconsin fine mesh (80 μm2) net with attached collection bucket

2 Wisconsin coarse mesh (243 μm2) net with attached collection bucket

2 Sample line, marked at 0.5 m increments

2 Secchi disk with cable

2/site+ 125 ml wide-mouth polyethelene sample jars (two per site, plus additional jars for replicates and 

other back-up sampling)

1 Squirt bottle with DIW

95% ethanol

2/site+ CO2 tablets

1/site+ 500 ml wide-mouth container

2 Two lids converted to form strainers (one with 80 μm, one with 243 μm mesh), made by drilling 

two holes in each lid and gluing a piece of the netting to the inside of the lid using silicone glue

2/site+ Zipper-lock plastic bag

Clear tape for covering labels

Electrical tape

1/site+ Zooplankton Sample Data Form

Pencils and permanent markers

Mild (10%) bleach solution for cleaning net and strainer lids between lakes and backwashing net 

(with a garden hose) after use

1 It is advisable to include some extras beyond what is needed for the number of sites to be sampled.

2These two mesh sizes (80 and 243 μm) are general guidelines. Other sizes could be used.

5.4.2 Equipment Cleaning Protocols 

Field survey personnel and/or their equipment can serve to transport pathogens and invasive 

species among water bodies. Field personnel should take appropriate precautions to minimize or 

eliminate this risk. General equipment cleaning guidelines are provided below. In addition, field 

staff should consult with local experts to determine if local conditions require any additional 

specific precautions. Between sample sites and at the duty station subsequent to field sampling, 

all gear that was exposed to stream or lake water should be thoroughly cleaned. Clothing, skin, 

and fingernails should also be cleaned. Gear should be disinfected using a 10% bleach solution or 

a solution of an alternative disinfectant and then thoroughly rinsed. Use of a high-pressure hose 

can be helpful. Gear should then be completely dried or allowed to dry before re-use at another 

lake or stream.

It is important to follow appropriate safety precautions when working with disinfectant products, 

especially as concentrated solutions. Such precautions include appropriate ventilation, use of 

impervious gloves and splash goggles, and access to eye wash stations.
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5.5 COLLECTION PROCEDURES 

5.5.1 Stream Benthic Macroinvertebrates 

The procedures recommended in this protocol for the collection and preservation of stream 

macroinvertebrates are based largely on the protocol designed by EPA for the EMAP-SW surface 

water sampling efforts. EMAP-SW protocols for invertebrate sampling are described in detail by 

Peck et al. (2006). 

The EMAP-SW benthic macroinvertebrate protocol was designed to evaluate the biological 

condition of wadeable streams in the United States for the purpose of detecting stresses on 

assemblage structure and assessing the relative severity of these stresses (Peck et al. 2006). It is 

based on the Level III procedure for benthic macroinvertebrates of the EPA Rapid Bioassessment 

Protocol (Plafkin et al. 1989, Barbour et al. 1999), which has been adopted for use by many 

states. 

Benthic macroinvertebrates are collected at each of 11 equidistant transects spaced throughout the 

support reach to ensure distribution of individuals among available major habitat types, eliminate 

individual sampler bias, and provide a comparable and consistent sample from every reach. All 

11 transect samples are combined into a single composite sample to characterize the support 

reach and reduce the cost and effort in processing and analysis (Patil et al. 1994, Barbour et al. 

1999, Roth et al. 2002). The number of individual field collections is expected to provide a 

composite sample having a sufficient number of individuals to characterize the taxonomic 

composition and relative abundance of the stream assemblage (e.g., Larsen and Herlihy 1998).

Samples are collected from each support reach with a D-frame kick net that can generally be used 

in the stream by one person (Figure 5-2). Typically, a field crew of two people collects kick net 

samples for benthic macroinvertebrates. One person typically collects the samples while the

second person times the collection of samples and records information on the field data form. 

However, in swift waters, two people may be needed to collect the samples.

Each kick net sample is collected at each of the 11 cross-section transects (transects A through K) 

at an assigned sampling point (Left, Center, or Right) as illustrated in Figure 5-1. Assign the Left, 

Center, or Right sampling point at transect A at random. Once the first sampling point is 

determined, assign points at successive transects in order (Left, Center, Right). At transects 

assigned a Center sampling point where the stream width is between one and two net widths 

wide, pick either the Left or Right sampling point instead. If the stream is only one net width wide 

at a transect, place the net across the entire stream width and consider the sampling point to be 

Center. If a sampling point is located in water that is too deep or otherwise unsafe to wade, select 

an alternate sampling point nearby. Never sample at an unsafe location. 

Collect a kick net sample at each transect as described below, beginning at the transect that is 

furthest downstream. Never collect benthic macroinvertebrates from a streambed location 

which you have recently disturbed (e.g., walked in). If a replicate composite sample is to be 

collected, do so at each transect within that support reach before moving upstream to the next 

transect. 
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Figure 5-2. Modified D-frame kick net. A): schematic drawing (not drawn to scale). (Source: Peck 

et al. 2006.) B): photograph showing EMAP crew member sampling a 

macroinvertebrate transect with modified D-frame net in Utah. (Photo: A. Herlihy.)

At each sampling point, determine if the flowing water or the slack water procedure is to be used 

based on whether or not there is enough current to extend the net. For each kick net sample, 

record the dominant substrate type: fine/sand, gravel, coarse substrate (coarse gravel or larger), or 

other (e.g., bedrock, hardpan, wood, aquatic vegetation, etc.), and the habitat type (pool, glide, 

riffle, or rapid) on the Sample Collection Form. Note that these substrate types and habitats are 

A)

B)
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defined in the table. Collect only from the upper 4 to 5 cm (1.5 to 2 in) of the substrate. As you go 

upstream from transect to transect, combine all the kick net samples into a single container, 

whether they were collected using the flowing water or slack water procedure.

To collect kick net samples for the reach-wide composite sample (taken from Peck et al. 2006):

1. At each cross-section transect, beginning at the downstream end of the reach with transect 

A (see Figure 5-1), locate the assigned sampling point (Left, Center, or Right as you face 

downstream) as 25%, 50%, and 75% of the wetted width, respectively. If you cannot 

collect a sample at the designated point because of deep water or unsafe conditions, 

relocate the point nearby on the same transect.

2. Attach the handle to the kick net. Make sure that the handle is on tight or the net may twist 

in a strong current, causing the loss of part of the sample.

3. Determine if there is sufficient current in the area at the sampling point to extend the net 

fully. If so, use the flowing water procedure (go to step 4). If not, use the slack water

procedure (go to step 10). 

For vegetation-choked sampling points where neither procedure can be used, sweep the net 

through the vegetation within a 0.09 m2 (1 ft2

) quadrat for 30 seconds. Place the contents of 

this hand-swept sample into the sampling container. Go to step 14.

Flowing Water Procedure:

4. With the net opening facing upstream, position the net quickly and securely on the stream 

bottom to eliminate gaps under the frame. Avoid large rocks that prevent the sampler from 

seating the net properly on the stream bottom.

Note: If there is too little water to collect the sample with the kick net, randomly pick up 10 

rocks from the riffle and pick and wash the organisms off them into a bucket which is halffull of water.

5. Holding the net in position on the substrate, visually define a rectangular quadrat that is one 

net width wide and one net width long upstream of the net opening. The area within this 

quadrat is ~0.09 m2 (1 ft2

). Alternatively, place a wire frame of the correct dimensions in 

front of the net to help delineate the quadrat to be sampled.

6. Hold the net in place with your knees. Check the quadrat for heavy organisms, such as 

mussels and snails. Threatened, endangered, or sensitive (TES) species must be left in place 

and not removed but they need to be recorded on the field form. Remove any heavy nonTES species from the substrate by hand and place them into the net. Pick up any loose 

rocks or other larger substrate particles in the quadrat. Use your hands or a small scrub 

brush to dislodge organisms so that they are washed into the net. Scrub all rocks that are 

golf ball-sized or larger and which are situated over halfway into the quadrat. Large rocks 

that are less than halfway into the sampling area are pushed aside. After scrubbing, place 

the substrate particles outside of the quadrat.

7. Keep holding the sampler securely in position. Start at the upstream end of the quadrat, use 

your foot and toes to vigorously kick the upper 4 to 5 cm (1.5 to 2 in) of the remaining 
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finer substrate within the quadrat for 30 seconds (use a stopwatch). Avoid going too deep 

into the substrate with your kicking.

Note: For samples located within dense beds of long, filamentous aquatic vegetation (e.g., 

algae or moss), kicking within the quadrat may not be sufficient to dislodge organisms in 

the vegetation. Usually, these types of vegetation lie flat against the substrate due to 

current. Use a knife or scissors to remove only the vegetation that lies within the quadrat 

(i.e., not entire strands that are rooted within the quadrat but extend beyond it) and place it 

into the net.

8. Pull the net up out of the water. Immerse the net in the stream several times to remove fine 

sediments and to concentrate organisms at the end of the net. Avoid having any water or 

material enter the mouth of the net during this operation.

9. Go to step 14.

Slack Water Procedure: 

10. Visually define a rectangular quadrat that is one net width wide and one net width long at 

the sampling point. The area within this quadrat is ~0.09 m2 (1 ft2

). Alternatively, lay a wire 

frame of the correct dimensions in front of the net at the sampling point to help delineate 

the quadrat.

Note: If there is not enough water present to use the net, spend 30 seconds collecting and 

examining pieces of substrate from about 0.09 m2 (1 ft2

) of substrate at the sampling point. 

11. Inspect the stream bottom within the quadrat for any heavy organisms, such as mussels and 

snails. Remove these organisms by hand and place them into the net or into a bucket. Pick 

up any loose rocks or other larger substrate particles within the quadrat and hold them in 

front of the net. Use your hands (or a scrub brush) to rub any clinging organisms off of 

rocks or other pieces of larger substrate (especially those covered with algae or other 

debris) into the net. After scrubbing, place the larger substrate particles outside of the 

quadrat.

12. Use your foot and toes to vigorously kick the upper 4 to 5 cm (1.5 to 2 in) of the 

remaining finer substrate within the quadrat while dragging the net repeatedly through the 

disturbed area just above the bottom. Continuously move the net forward so that the 

organisms trapped in the net do not escape. Continue kicking the substrate and moving the 

net for 30 seconds. 

Note: If there is too little water to use the kick net, vigorously stir up the substrate with 

your gloved hands and use a sieve with 500 μm mesh size to collect the organisms from the 

water in the same way the net is used in larger pools.

13. After 30 seconds, remove the net from the water with a quick upstream motion to wash the 

organisms to the bottom of the net.

All samples:

14. Invert the net and transfer the contents into a bucket or wide-mouthed container with a lid 

labeled REACHWIDE. Inspect the net for any residual organisms clinging to the net and 
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149

deposit them into the REACHWIDE container. Use a squirt bottle with stream water and 

watchmakers’ forceps if necessary to remove organisms from the net. Carefully inspect any 

large objects (such as rocks, sticks, and leaves) in the bucket and wash any organisms 

found off of the object and into the bucket before discarding the object. Remove as much 

detritus as possible without losing any organisms. Replace the lid on the bucket or 

container.

15. Determine the predominant substrate size/type you observed within the sampling quadrat. 

Place an X in the appropriate substrate type box for the transect on the Benthic 

Macroinvertebrate Sample Collection Form. 

Note: If there are co-dominant substrate type(s), you may check more than one box; note 

the co-dominants in the comments section of the form.

Fine/sand: not gritty (silt/clay/muck < 0.06 mm diam.) to gritty, up to ladybug sized 

(2 mm diam.).

Gravel: fine to coarse gravel (ladybug to tennis ball sized; 2 mm to 64 mm diam.).

Coarse: cobble to boulder (tennis ball to car sized; 64 mm to 4000 mm).

Other: bedrock (larger than car sized; > 4000 mm), hardpan (firm, consolidated fine 

substrate), wood of any size, aquatic vegetation, etc. Note type of “other” substrate in 

comments on field form.

16. Identify the habitat type where the sampling quadrat was located. Place an “X” in the 

appropriate channel habitat type box for the transect on the Sample Collection Form. 

Pool: still water; low velocity; with smooth, glassy surface; usually deep compared to 

other parts of the channel.

GLide: water moving slowly, with smooth, unbroken surface; low turbulence.

RIffle: water moving with small ripples, waves, and eddies; waves not breaking, and 

surface tension is not broken; “babbling” or “gurgling” sound. 

RApid: water movement is rapid and turbulent; surface with intermittent “white water” 

with breaking waves; continuous rushing sound.

17. Proceed upstream to the next transect (including all transects in sequence through transect 

K, the upstream end of the support reach) and repeat steps 1 through 16. Combine all kick 

net samples within the sample reach into the REACHWIDE container.

18. Thoroughly rinse the net with stream water before proceeding to the next sampling 

location. It is also extremely important that all equipment, including waders, be cleaned 

between sites to avoid transmission of non-native invasive species.

If the kick net cannot be used to collect a sample at a flowing water sampling point, select the 

number of rocks necessary to cover approximately 0.09 m2 (1 ft2

) of the streambed from the area 

near the sampling point within the area of flowing water. Inspect and remove any organisms 

found on each rock and place them into the sampling container. If the kick net cannot be used at a 

slack water habitat due to insufficient depth of water, spend about 30 seconds picking up pieces 

of substrate from a 0.09 m2 (1 ft2

) area at the sampling point. Inspect and remove any organisms 

found on each piece of substrate and place them into the sampling container. At vegetation-
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choked sampling points where neither procedure can be used, sweep the net through the 

vegetation for 30 seconds and place the contents into the sampling container.

5.5.2 Lake Zooplankton

The procedures recommended in this protocol for collection and preservation of lake zooplankton 

are based largely on the protocols designed by EPA for the National Lakes Survey conducted in 

2007 (U.S. EPA 2007) using a “Wisconsin” plankton net. General and detailed procedures are as 

follows.

Two vertical plankton tow samples are collected at the lake sampling index site location. One 

sample is typically collected using a fine mesh (typically ~ 50 to 80 μm) and one sample is

collected using a coarse mesh (typically ~ 200 to 250 μm) plankton net (Figure 5-3). We 

recommend a 80 μm Wisconsin net for the fine mesh and a 243 μm Wisconsin net for the coarse 

mesh. Each net is attached to a collection bucket. Some of the larger species of zooplankton can 

swim fast enough to avoid being caught in the net. The coarse mesh net optimizes capture of the 

faster-swimming macrozooplankton because the pressure wave above the net is minimized. The 

fine mesh net optimizes capture of the microzooplankton. The two samples are collected and 

analyzed separately. 

Figure 5-3. Wisconsin net and collection bucket diagram. Some microzooplankton nets have a 

reducing collar attached. (Source: U.S. EPA 2007.)

Each tow is collected by pulling the sampling apparatus from a depth of about 1 m above the lake 

bottom to the lake surface. It is important to avoid touching lake sediments with the sampling 

apparatus, as that can clog the net pores and compromise the integrity of the sample. The net 

should be raised steadily, but rather slowly (~ 1 ft/sec), to reduce the pressure wave that can build 

up at the top of the net during retrieval. Some species can detect this wave and swim out of the 

path of the net. Use of a wide net aperture (30 to 50 cm) can be helpful to avoid missing fastswimming taxa such as Chaoborus, Leptodora, and Mysis.
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If the lake depth at the sampling site is less than 2.0 m and the Secchi disk is visible at the 

bottom, a second vertical tow is made with each net (fine and coarse mesh) and the original (first) 

and the second samples are combined. Note that the samples collected using the fine and coarse 

mesh nets are not combined. If the net or attached collection bucket touches the lake sediment, 

field personnel should retrieve and rinse the apparatus and repeat the process. Description of the 

procedures to be followed for sample collection is provided above. If other mesh sizes are used 

instead of those that we recommend, it is important to standardize these mesh sizes such that 

there is consistency among sites and over time in the sampling program. 

To collect zooplankton sample (taken from U.S. EPA 2007):

1. Fill out sample label.

2. Measure lake depth at sample location.

3. Clean and thoroughly rinse the inside surfaces of the nets and collection buckets with DIW.

4. Inspect nets and buckets for possible holes or tears.

5. Attach collection bucket to small end of each net.

6. Attach marked (every 0.5 m) lines to large end of coarse net.

7. Lower coarse net in constant upright position over side of boat until the mouth of the net is 

about 1.0 m above the lake bottom. If the lake depth is less than 2 m and the Secchi disk 

can be seen at the bottom, collect a second tow with the coarse net and combine the 

replicated samples (make note of this on data form).

8. Retrieve net to surface at constant rate (about 1 ft per second) without stopping.

9. At the surface, slowly move the net up and down in the water column, without submersing 

the net mouth, in order to flush zooplankton from the net into the collection bucket. 

10. Further rinse contents from net into collection bucket by spraying net from outside to inside 

with squirt bottle containing DIW.

11. Holding collection bucket in vertical position, detach it from net.

12. Swirl the bucket without spilling the contents in order to filter excess water out of the 

bucket through the screened sides.

13. Repeat steps 6 through 12 with the fine mesh net on the opposite side of the boat. 

5.6 SAMPLE PROCESSING, PRESERVATION, AND 

HANDLING 

5.6.1 Stream Benthic Macroinvertebrates 

After collecting kick net samples for the reachwide samples, prepare a composite index sample 

from the contents of the container as described below. You will need to record tracking 

information for each composite sample on the Sample Collection Form. Check to be sure that the 

completed label on each jar is covered with clear tape and that a waterproof label is placed in 

each jar and filled in properly. Confirm that the inside and outside labels describe the same 

sample. Replace the lid on each jar and seal with plastic electrical tape. It is helpful to mark the 

lid of each jar with the site number; use a permanent marker or write on a piece of light-colored 
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tape (or a small blank address label) and attach it to the lid. Place the sample jars in a cooler or 

other secure container for transporting and/or shipping it to the laboratory. The container and 

absorbent material placed between the jars both should be suitable for transporting ethanol. Check 

to see that all equipment is returned to the vehicle. Samples do not need to be kept on ice or 

cooled after they are preserved with ethanol.

Procedure for preparing composite samples for benthic macroinvertebrates (taken from Peck et al. 

2006) are:

1. Pour off the water from the reachwide bucket through a sieve (or sieve bucket) with 500 

μm mesh size. Remove any large objects such as sticks, rocks, or large plant material from 

the bucket or container. Inspect these objects carefully and dislodge any clinging organisms 

back into the sample bucket or container before discarding.

2. Estimate the total volume of the sample in the sieve and determine the size (500-ml or 1-L) 

and number of jars that will be needed for the sample. Avoid using more than one jar for 

each of the composite samples if possible, but don’t fill the jar more than 1⁄4 full with each 

composite sample.

3. Fill in a sample label with the stream ID, date of collection, and other required information. 

Attach the completed label to the jar and cover it with a strip of clear tape. Record the 

sample ID number for the composite sample on the Sample Collection Form. For each 

composite sample, make sure the number on the form matches the number on the label.

4. Wash the contents of the bucket or container to one side. Transfer the sample from the 

bucket or container into a jar, using a large-bore funnel if necessary. Use as little water 

from the wash bottle as possible to help transfer material. If the jar becomes too full of 

liquid, carefully pour off the water through the sieve. Continue to transfer sample material 

to the jar until it is not more than 1⁄4 full of solid material. Use additional jars for the 

remaining sample. Carefully examine the bucket or container for any remaining organisms 

and use watchmakers’ forceps to place them into the sample jar.

If a second jar is needed, fill in a sample label that does not have a pre-printed ID number 

on it. Record the ID number from the pre-printed label prepared in step 4 in the SAMPLE 

ID field of the label. Attach the label to the second jar and cover it with a strip of clear tape. 

Record the number of jars required for the sample on the Sample Collection Form. Make 

sure the number you record matches the actual number of jars used. If possible, write Jar N 

of X (N being the sequential jar number and X being the total number of jars for the 

sample) on each sample label using a waterproof marker.

5. Place a waterproof label with the following information inside each jar:

• Stream ID number

• Name of stream

• Date of collection

• Collector’s initials
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6. Remove as much water as you can from each sample jar without removing any sample 

material by pouring it through the sieve. If possible, completely fill each jar with 95% 

ethanol (no headspace) so that the final concentration of ethanol is between 75 and 90%. It 

is very important that sufficient ethanol be used or the organisms will not be properly 

preserved. Do not freeze samples to preserve them.

Note: For backcountry work, prepared composite samples should be transported back to the 

vehicle before adding ethanol. In that case, fill each jar with stream water and a minimal 

amount of ethanol to cushion the sample from the grinding action of non-biological 

material in the sample during transport. Replace the water with ethanol at the vehicle as 

soon as possible.

7. Replace the lid on each jar. Slowly tip the jar to a horizontal position, then gently rotate the 

jar to mix the preservative. Do not invert or shake the jar. After mixing, seal each jar with 

plastic tape.

8. Store the labeled sample jars in a container with absorbent material that is suitable for use 

with 95% ethanol until transport or shipment to the laboratory.

5.6.2 Lake Zooplankton 

After rinsing the outside of the plankton net using the squirt bottle with DIW, transfer the sample 

to one (or more, if needed) sample jars. The collected zooplankton are doped by adding CO2

tablets or alka-seltzer to stop their movement and then preserved. Detailed procedures for sample 

preservation are described below. Zooplankton samples, once preserved, do not need to be stored 

on ice and can be shipped via ground transport to the laboratory. Field personnel should include 

one copy of the data form along with the samples (each in its own plastic bag) when they are 

shipped to the laboratory. Take one copy of the data form back to the office in the lake-data

folder. The development and use of a folder for each study lake is described in Section 1.3.1 of 

the Water Chemistry Field Sampling Protocols, Establishing and Locating Sampling Sites. The 

sample jars should be surrounded with packing material before shipping.

Detailed procedures for processing zooplankton samples (taken from U.S. EPA 2007) are:

1. Place bucket that had been attached to the coarse net into a 500 ml container filled threefourths full with lake water, to which a CO2 tablet has been added (alternatively, AlkaSeltzer or club soda can be used). Wait until zooplankton have been narcotized and stop 

moving (about 1 minute). 

2. Transfer contents of bucket into a 125 ml polyethylene jar using DIW from the squirt 

bottle. Continue to rinse the bucket until the majority of the collected zooplankton are 

transferred to the jar. 

3. Drain much of the excess water out of the jar by attaching a lid modified to create a strainer 

that has been prepared in advance by cutting two holes in the lid and gluing small pieces of 

the appropriate (large or small) mesh material to the inside of the lid to cover the holes. 

Carefully decant the excess water out of the jar while retaining the zooplankton inside the 

jar. 
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4. Fill the jar a little more than half full with 95% ethanol12. If the volume of zooplankton 

collected fills the jar more than half full, use a second (and third, if necessary) jar to 

preserve the additional sample volume. Record the number of jars used on the Zooplankton 

Sample Data Form. Label each jar identically and then add to the labels, as appropriate, “1 

of x”, “2 of x”, etc., with x being the total number of jars used for the sample. 

5. Record the length of tows collected on the Zooplankton Sample Data Form. Verify that all 

required information is provided on sample labels and data form. Cover each label with 

clear tape. 

6. Seal jar lids by wrapping electrical tape around the juncture of the lid in a clockwise 

direction so the lid is pulled tight as tape is stretched around it. 

7. Place each jar in a zippered plastic bag.

8. Repeat steps 1 through 7 for the second (fine mesh) sample collected. 

9. Thoroughly clean and rinse all equipment and the strainer lids before transporting them to 

another lake.

5.7 DOCUMENTATION AND TRACKING

Data collection forms for stream macroinvertebrate sampling and lake zooplankton sampling can 

be found on the Utah State University BugLab website (<http://www.usu.edu/buglab/>). Example 

labels for stream benthic macroinvertebrate samples and lake zooplankton samples are shown in 

Appendix F. Data collection forms and labels should be carefully filled out and double-checked 

for completeness and accuracy before leaving the field site. 

5.8 LABORATORY ANALYSIS 

Biological samples will need to be sent to a contract or agency laboratory, where experts will 

enumerate and identify the individual organisms in the composite sample. Taxonomic richness 

results are very sensitive to both counting effort (e.g., how many individuals are counted) and 

taxonomic resolution. Thus, it is imperative that a consistent laboratory counting protocol be used 

when multiple labs are involved or samples are analyzed over a period of time for trends 

determination. For the stream benthic IBI assessment, we recommend a 500 fixed organism count 

protocol using a gridded sorting tray (typically, a 5x6 grid of 30 cells). Individual grid cells from 

the tray are selected at random and completely processed until more than 500 organisms are 

enumerated. The percent of the sample processed is calculated as number of grids processed/total 

number of grids and this number used to infer the total number and density of individuals in the 

composite sample. For counts of the EPT taxa only, grid cells should be processed either until 

100 individuals of the EPT taxa have been enumerated or until 500 total organisms have been 

enumerated. 

 12 For backcountry sampling, add only a small amount of ethanol in the field and then fill jars to near the top with 

water. Upon returning to the vehicle, discard the water and replace with ethanol.
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Benthic organisms should be identified, if possible, to the genus level except for the following 

non-insect taxa: oligochaetes, polychaetes, and arachnids to family; nematodes and 

platyhelminthes to phylum. In most cases, identification to family should be considered the 

minimal requirement. For a basic EPT taxa richness assessment, the EPT orders should be 

identified to genus. For lake zooplankton, individuals should be identified to the species level, 

where possible. Each net sample is counted independently and at least half the sample volume 

examined. Subsamples are examined and counted until no new species are found or until a total 

of 300 to 500 individuals have been counted. Either approach is acceptable; the choice should be 

based on the standard protocol of the laboratory. 

5.9 QUALITY ASSURANCE

5.9.1 Sample Replication

It is always advisable to replicate a portion of the samples, regardless of whether they are 

chemical or biological. This offers an opportunity to evaluate variability that may be introduced 

in the course of sampling, preserving, and analyzing the samples. Although we do not 

recommend that replicate zooplankton or benthic macroinvertebrate sampling should necessarily 

be required, we do think it is a good idea. Our recommendation is that about 5% to 10% of the 

sampled lakes or streams be replicated. Sample information provided on the data form for the 

replicate in this case will be identical to that of the original sample except for the sample ID and 

the time of sampling, which will differ slightly between the first and second samples at a given 

site. 

The replicate zooplankton sample should be collected at the same general location as the primary 

(first) sample, on the opposite side of the boat. If a stream site is to be replicated, the additional 

(replicate) sample is collected at each transect location to yield a pooled composite replicate, 

comparable to the composite normal sample. The replicated stream benthic macroinvertebrate 

sample at each transect location along the sample stream reach should be collected at a different 

stream location from the normal sample. For the replicate, move the transect sample location 

from L to C; from C to R; and from R to L. Check the box on the data form indicating whether or 

not the sampling was replicated at this site. If the stream is not wide enough to accommodate 

collection of a second (replicate) sample, slide the replicate site upstream about 10 m and collect 

the replicate sample there. With replicate sample, extra caution must be taken to not disturb any 

of the actual sampling sites by walking in them before sampling. 

5.9.2 Taxonomy

A major potential pitfall in any aquatic invertebrate study is the inherent variability and 

uncertainty in taxonomy among aquatic entomologists and among laboratories. This can be 

especially problematic in a long-term monitoring study if different laboratories or laboratory staff 

are involved in the identification of collected organisms over the course of the study. We 

recommend choosing a highly experienced taxonomic laboratory and trying to maintain 

consistency throughout the project. When multiple labs are involved, interlaboratory QA is 

essential. In EPA’s National Stream Survey, which used eight different laboratories, 10% of the 

samples were randomly selected for QC re-identification and sent to an independent taxonomist 
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in a separate laboratory for comparison (Stribling et al. 2008). The results of the sample-based 

comparisons were summarized as percent taxonomic disagreement (PTD) and percent difference 

in enumeration (PDE). Percent difference in enumeration differences among labs were minor 

(<3%) but PTD were on the order of 20%. Having lab taxonomists intensively interact, resolve 

differences, and update the data after the first round of identifications was important, and 

improved PTD substantially in the EPA survey. We also recommend that at least 5% to 10% of 

samples be sent to an alternate laboratory or alternate entomologist to evaluate any differences 

that might arise in taxonomic identification. Such differences should be resolved, if possible, 

before finalizing the dataset. 

5.10 INTERPRETATION

Analysis of lake or stream water quality data can provide critical information regarding the status, 

or change over time, in biologically relevant water chemistry. Thus, it may be known or 

suspected based on measured water chemistry that in-lake or in-stream biota respond to a given 

concentration (or change in concentration) of ANC, pH, Ali, etc. Nevertheless, there is always 

some degree of uncertainty regarding the biological effects that actually occur under a given suite 

of water chemistry. Federal land managers can draw stronger inferences about biological effects 

if the biological resource itself is characterized or monitored. This can be important in setting 

target deposition loads, pursuing litigation, and evaluating damage or recovery scenarios. 

Substantial value can be gained by sampling biota in addition to water chemistry. 

5.10.1 Biological Metrics 

Biological assemblage data are typically analyzed by calculating metrics from the list of the 

species, genera, or families identified and their abundances. For example, the number of different 

mayfly genera in the sample can be tallied and this number becomes the mayfly genus richness 

metric. Richness metrics can be calculated for any defined taxonomic group (e.g., mayflies, 

rotifers, or insects) as well as total sample richness. Similarly, richness can be calculated for any 

other autecological attributes such as functional feeding groups (shredder richness), habitat 

preference (swimmer richness), or tolerance to various pollutants. In addition, the same type of 

metrics can be calculated based on percent of individuals in the sample (e.g., percent mayfly 

individuals or percent shredder individuals). There are also metrics to reflect overall sample 

diversity that are based on equations that aim to mathematically express diversity as a 

combination of overall sample richness (number of different taxa) and evenness (equal number of 

individuals across taxa).

A simple assessment can be made based on a single metric that is responsive to specific pollutants 

of interest. For example, mayflies are very sensitive to pH, and mayfly taxonomic richness is 

therefore a good metric to use for acidic deposition studies. Total sample richness or diversity 

may also be used as a single overall measure of biological condition. The most robust measures 

of biological condition, however, require modeling or combining multiple metrics into one 

overall multi-metric index. Application of a multi-metric index or model requires some expertise 

in the biotic assemblage being assessed. In particular, gathering the necessary autecological 
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information can be time-consuming and require detailed knowledge of the different species that 

occur within the study region. 

5.10.2 Multimetric Indices and Predictive Modeling

There are two major assessment approaches for quantifying whole community biological 

condition: multimetric indices (e.g., IBI) or predictive modeling (e.g., the observed/expected or 

“O/E” approach). A multi-metric index, such as the IBI, is developed by selecting the best 5 to 15 

metrics that quantify condition over a suite of different aspects of biotic integrity and then 

summing individual metric scores into a single index of condition. Metric selection and 

interpretation of metric values at the sampling site are usually based on values observed at leastdisturbed reference sites in similar settings. 

The predictive-modeling approach uses reference sites to assemble lists of taxa that appear to be 

indicative of a least-disturbed reference condition (the expected or “E” list). Taxa lists from a 

specific study site comprise the observed, or “O,” list. The proportion of the expected taxa found 

in the observed list (O/E ratio) is a measure of the proportion of the taxa expected to be at an 

undisturbed site that are actually present at the study site. An O/E ratio of one indicates a highquality site (all expected taxa present). An O/E ratio of <0.5 means that less than half the 

expected reference taxa are present at the site. In practice, the E list is developed for each study 

site by statistical modeling (cluster analysis and discriminate function analysis) of reference site 

data to take into account natural differences in expected taxa distributions. 

This modeling approach was pioneered in Great Britain (Moss et al. 1987) and has been applied 

in many different locations throughout the world. These predictive models require statistical 

expertise to develop in new regions and a large number of reference sites as the basis for the 

modeling. Study-site and reference-site data must be collected with comparable field protocols, 

lab protocols, and taxonomic resolution. Due to its complexity and data requirements, we do not 

recommend the O/E approach as a routine tool for biological assessment on FS lands potentially 

influenced by atmospheric S and N deposition. Nevertheless, this can be a powerful tool for 

stream biological assessment if one is willing to develop the modeling approach and referencesite database for a given region or study area. 

5.10.3 Taxonomic Resolution

In conducting any biological assessment, the level of taxonomic resolution (e.g., order, family, 

genus, or species) is an important consideration. In general, identifications to lower taxonomic 

levels cost more but provide more information. For stream macroinvertebrates, identification is 

usually taken to either the family or genus level. Some organisms can be identified to species,

but, in a given sample, most of the organisms can only be identified to genus due to the lack of 

sample keys for many taxa and the small size of early life history stages of many of the 

individuals. With stream macroinvertebrates, the major laboratory effort involves picking the 

organisms out of the sample matrix rather than identifying them. Therefore, the laboratory cost 

difference between family- and genus-level analyses may not be substantial. In terms of 

information content, Waite et al. (2004) found that family- and genus-level stream 

macroinvertebrate data were similar in their ability to distinguish among the coarse impacts (e.g.,

most severe versus least severe impact classes). 
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Genus data, however, often distinguished the subtler differences in mid-Atlantic streams (e.g., 

mixed/moderate impacts versus high or low impacts) better than family level data. In their 

analysis, acidic deposition impacts were considered a moderate impact and not a severe impact. 

Ordination analysis showed that both family and genus levels of analysis responded to similar 

suites of environmental variables. 

We suggest that identification to the family level can be sufficient for many bioassessment 

purposes. However, identifications to genus do provide more information, especially in generarich families like Chironomidae. Genus or finer levels of identification are important for 

investigating natural history, stream ecology, biodiversity, and indicator species. Decisions about 

the taxonomic level of identification need to be study specific and depend on available resources 

(cost) and study objectives.

5.10.4 Index of Biotic Integrity

The IBI is a multi-metric index that has been used extensively in streams to characterize fish, 

macroinvertebrate, and periphyton condition. Note that we do not recommend application of an 

IBI as part of the routine process of evaluating biological response to atmospheric deposition 

stressors on FS lands. 

In general, individual analyses for one or more of the EPT orders or application of an EPT Index 

is often sufficient. If, however, there is a need to more fully characterize biological conditions in 

a particular stream reach and if appropriate invertebrate taxonomic and autecological expertise is 

available to the project team, then a stream benthos IBI is an appropriate way to proceed. Once 

the samples have been collected, there is not usually a dramatic difference in cost to enumerate all 

taxa (for implementation of an IBI) as opposed to just the insect orders included in the EPT. 

Nevertheless, application and interpretation of the IBI does require that more specialized 

taxonomic and autoecological expertise be available to the project.

There are a number of different approaches to calculating IBIs, but they all follow a similar 

process. First, the metrics that best reflect condition are selected from the set of candidate metrics. 

IBIs typically are comprised of 5 to 15 different metrics. Metric values are then scored to a 

consistent scale (e.g., 0-10 points) and summed to calculate the one overall IBI value. 

A wide variety of IBIs have been developed for different stream types and regions around the 

world. For assessing stream benthos, we recommend as a starting point the macroinvertebrate IBI 

developed by EPA for the National Wadeable Streams Assessment, in part because it was 

designed to be applied nationwide (Stoddard et al. 2008). The Wadeable Streams Assessment IBI 

is formulated differently for each of nine different ecoregions in the United States (Figure 5-4). 

Candidate metrics were divided into six different categories and the best performing metric in 

each category was selected for inclusion in the regional IBI. The six metrics in each regional IBI 

are listed in Table 5-4. The six metrics values were then each scored on a 0-10 scale and summed 

into a final IBI score (see Stoddard et al. 2008 for calculation details). There has been much less 

work done on developing IBIs for lake systems. We do not recommend calculating an IBI for 

lake zooplankton at this time as we are not aware of any existing IBIs that are ready for use.
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Figure 5-4. Location of the nine ecoregions used in Wadeable Streams Assessment IBI development (see Table 5-4 for ecoregion abbreviations).

(Source: USEPA 2006.)
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Table 5-4. List of metrics in each category used in the EPA National Wadeable Stream Assessment. (Source: Stoddard et al. 2008). Metrics were 

selected and scored separately for each of nine aggregate ecological regions: NAP (Northern Appalachians); SAP (Southern Appalachians); 

CPL (Coastal Plain); UMW( Upper Midwest); TPL (Temperate Plains); NPL (Northern Plains); SPL (Southern Plains); WMT (Western 

Mountains); and XER (Xeric).

Metric Category Individual Metrics

Aggregate Ecological Regions

NAP SAP CPL UMW TPL NPL SPL WMT XER

Composition % EPT Taxa X X X

% EPT Individuals X X

% Non-Insect Taxa X

% Non-Insect Individuals X

% Ephemeroptera Taxa X

% Chironomid Taxa X

Diversity Shannon Diversity X X X X X

% Individuals in top 5 taxa X X X

% Individuals in top 3 taxa X

Feeding Scraper Richness X X X X X X X

Shredder Richness X X

Habitb % Burrower Taxa X X X X

% Clinger Taxa X X X X

Clinger Taxa Richness X

Richness EPT Taxa Richness X X X X X X X

Ephemeroptera Taxa Richness X

Total Taxa Richness X

Tolerance Intolerant Richness X X

% Tolerant Individuals X X X X

% PTVa 0-5.9 Taxa X

% PTV 8-10 Taxa X X

a PTV = Pollution Tolerance Value b Habit reflects the life strategy of the various taxa with respect to maintaining position in the stream (i.e., burrowing, clinging)
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5.10.5 Biotic Effects Analysis

If an atmospheric deposition effects study design calls for biological characterization—for 

example, in conjunction with a chemical characterization or monitoring effort—we recommend 

analysis of taxonomic richness of stream benthic macroinvertebrates and/or lake zooplankton for 

a group of streams or lakes across a gradient of acid-base chemistry. Such an analysis should be 

based on at least 10 water bodies and preferably more. The preferred chemical metric is usually 

ANC; the analysis should also be conducted for pH, Ali, and/or NO3

- concentration. The preferred 

biological metric is species richness or genus richness; in some cases family richness is the best 

that can be done because of taxonomic uncertainties. The taxonomic groups to be considered can 

include crustaceans and/or rotifers for lakes; mayflies, caddisflies, and/or stoneflies for streams; 

and/or some combination of the above. 

The basic data analysis for studying the effects of stressors on biological condition involves 

plotting biological metric scores or IBI scores versus water chemistry, as shown schematically in 

Figure 5-5A for mayfly genera richness. The strength of the relationship can be evaluated using 

the r

2 statistic. This analysis provides useful information on the extent to which invertebrate 

biological assemblages are associated with water acid-base chemistry. Trends analysis cannot be 

used to interpret biological change in response to improved or declining acid-base chemistry 

unless this basic analysis is performed and yields a meaningful relationship. 

For water bodies that are included in long-term chemical monitoring, one should also consider 

subjecting at least a subset of those water bodies to biological monitoring. The biological 

monitoring candidates should preferably be relatively low in ANC and pH, exhibit chemistry that 

is not excessively variable within and among years, and exhibit reasonably rich biological 

assemblages. Selection of two to four waters, spread across the ANC gradient (to the extent that 

such a gradient occurs) between about -50 μeq/L and 50 or 100 μeq/L, would be appropriate. 

Resulting monitoring data should be analyzed as shown schematically in Figure 5-5B (or some 

variation thereof). This analysis allows determination of the extent to which chemistry and biotic 

richness are deteriorating or improving over time, and the degree to which those trends are linked. 

Variability in any of the figures used to examine relationships between stream or lake chemistry 

and biological community metrics can be caused by changes in environmental conditions, 

especially hydrological conditions. For that reason, it is always advisable to examine the 

influence of weather/hydrology on the observed relationships. This can easily be done by coding 

the points on any of these figures according to hydrological conditions (in discrete classes). This 

can be based on inlet or outlet stream discharge (i.e., cumulative seasonal or annual stream flow), 

seasonal or annual precipitation, date of snowmelt, or other variable constructed to represent the 

differences between wet years or seasons and dry years or seasons. This allows the analyst to 

determine to a first approximation the extent to which the observed relationships between 

chemistry and biology are influenced by hydrological differences.
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Figure 5-5. Schematic depicting mayfly richness over time in response to changes in stream 

chemistry. Plot A shows richness plotted against chemistry; Plot B shows both richness 

and chemistry plotted against time.

R2 = 0.7942

Average Numer of Mayfly Genera

Recorded in a Given Year

Average Stream ANC Measured in a Given Year

Low High

High

Low

A

0 1 2 3 4 5 6 7 8 9

Average Number of Mayfly Genera

Recorded in a Given Year

Year

Average Measured Stream ANC

Number of Mayfly Genera ANC

High

Low

High

Low

B
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STREAMS

For documenting biological effects in streams in response to changes in atmospheric deposition, 

we recommend analyzing the quantitative relationships between invertebrate community metrics 

and stream ANC in multiple streams selected across an ANC gradient within a given forest or 

wilderness. The same analyses could also be done using the variables pH, NO3

-

, and Ali. For an 

initial analysis, we further suggest that, for studies of response to acidic deposition, the analysis 

can be limited to insects (class Insecta of the phylum Arthropoda) of the orders Ephemeroptera, 

Plecoptera and Trichoptera (EPT, or mayflies, stoneflies, and caddisflies) because of their general 

importance to stream ecology and their demonstrated responsiveness to changes in acid-base 

chemistry. 

We recommend examination of the number of genera (or if that is not possible, families) present 

within each of these three orders, both individually and combined, in relation to differences 

among streams in stream chemistry. Figure 5-6 shows an example of this analysis for mayflies in 

streams in Shenandoah National Park (Sullivan et al. 2003). The analysis was based on both the 

minimum ANC and the average ANC of multiple measurements in a given stream. The same type 

of analysis can be conducted for a single chemistry measurement from each stream, if that is what 

is available. The analysis shown in Figure 5-5 should be conducted for all three of the principal 

insect orders, plus for the EPT Index. 

The EPT Index is calculated either as the total number of genera or the total number of families 

present in a given stream from the orders Ephemeroptera, Plecoptera, and Tricoptera. It represents 

the number of genera or families among all three orders enumerated in a single sample or in the 

average of multiple samples. In general, we recommend basing an EPT Index on the number of 

genera present. If that is not possible, the calculation can be based on the number of families

present. The data shown in Figure 5-6 illustrate, as is often the case, that relationships between 

mayfly richness and stream chemistry are typically stronger than relationships for caddisflies. 

Stoneflies alone are often not very sensitive to changes in ANC and pH. 

For trends analysis of change in benthic insect diversity over time, we recommend plotting the 

number of genera or families (within each of the three orders individually, and combined as an 

EPT Index) recorded for one or multiple (averaged) samplings from a given stream each year 

over a period of at least 8 years. This will provide an assessment of possible changes in benthic 

insect richness over time that can then be related to possible changes in stream ANC or some 

other variable. For example, the average number of genera or families of mayfly recorded during 

various samplings in a given year (y-axis) should be plotted against the average ANC (x-axis) 

determined for those same sampling occasions over the period of study. In addition, both the 

average number of genera or families of mayfly and the average stream ANC should be plotted 

over time (across the years of record) using the same time scale (cf. Figure 5-5). Such analyses 

allow evaluation of the extent to which changes in biota are associated with change in chemistry 

and the degree to which either or both are changing over time. 

If the result of application of an EPT index is not clear, that result may be attributable to a lack of 

invertebrate response or it may be that the index is not sufficiently sensitive to illustrate the 

biological response that has occurred. In such a situation, FS staff should consider the possibility 

of applying an IBI, which may be a more powerful approach.
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Figure 5-6. Average number of families of aquatic insects for each of 14 streams in Shenandoah 

National Park versus the mean (left) or minimum (right) ANC of each stream. The stream 

ANC values are based on quarterly samples from 1988 to 2001. The invertebrate 

samples are contemporaneous. Results are presented for the orders Ephemeroptera 

(top), Plecoptera (center), and Tricoptera (bottom). The regression relationship and 

correlation are given on each diagram. (Source: Sullivan et al. 2003.)

LAKES

We recommend, for lake characterization studies focused on acidification, analyzing zooplankton 

data for more than 1 and preferably 10 or more lakes across an ANC gradient to determine 

whether any relationships exist between zooplankton richness and lake ANC. Parallel analyses 

can also be conducted for pH and Ali in addition to ANC. These analyses should be conducted for 

all zooplankton groups combined (total zooplankton) and for discrete groups of zooplankton. The 

discrete groups should include crustaceans and rotifers at a minimum and could also include large 

cladocerans. An example for Adirondack lakes in New York, showing the number of zooplankton 

species versus ANC at the time of zooplankton survey, is shown in Figure 5-7. 

It is generally expected that variation (or scatter) in the relationships between lake chemistry and 

taxonomic richness may increase as the size of the study area increases. An analysis such as that 

shown in Figure 5-7 for a large region may yield so much variability that patterns are not clear. It 
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may be necessary to restrict analyses such as this to a specific wilderness or to a designated 

subset of a region or forest, such as a certain geological type, ecoregion, or elevational band. It is 

advisable to examine differences in the relationships between zooplankton and lake chemistry 

under varying schemes for subsetting the data into groupings of lakes that are generally more 

similar to each other than the group of all lakes across a given region. 

If there are clear relationships between zooplankton species richness and lake chemistry across a 

wilderness, forest, or designated subset of lakes within a wilderness, forest, or region, then it can 

be useful to develop a time-series database for one or more presumed acid-sensitive lakes (having 

ANC between about -20 and +50 μeq/L). Such a database would entail contemporaneous 

zooplankton species richness and lake ANC measurements over a period of time of at least 8 

years. Plots can then be constructed to determine, for a given lake, the relationship between ANC 

and zooplankton richness and changes in both of these variables over time using plots such as 

those depicted in Figure 5-5. We do not recommend this as a standard component of long-term 

chemical monitoring efforts. However, if it is important to document changes in biological effects 

in response to anticipated changes in lake chemistry, then a time series of zooplankton richness 

(for crustaceans, rotifers, total zooplankton, or other taxonomic grouping) may be the most 

straight-forward and cost-effective strategy.

Biotic assemblages in lakes vary at both temporal and spatial scales influenced by such factors as 

climate, vegetative cover, and disturbance. Therefore, environmental indicators exhibit variability 

that has a great influence on our ability to estimate biological status or trends over time. 

Stemberger et al. (2001) attempted to quantify the various contributions to the variance in 

zooplankton status as part of EPA’s EMAP sampling program in the northeastern United States. 

Variance in zooplankton indicators was attributed primarily to four components of variance:

1. Lake variance: the lake-to-lake variability in zooplankton indicators in the study 

population. This depends largely on such factors as lake size, depth, fish presence/absence, 

pH, thermal characteristics, and productivity (Dodson et al. 2000). 

2. Year variance: coherent variation from year-to-year across all lakes, due, for example, to an 

unusually warm or wet weather pattern.

3. Lake-by-year interaction variance: independent year-to-year variation at each lake due to 

site-specific forcing factors, such as variation in nutrient inflows or mixing regime. 

4. Index variance: local spatial and temporal variance due, for example, to within-index 

period temporal changes, measurement error, or differences among crews or laboratories in 

application of the protocols (Stemberger et al. 2001).

In general, Stemberger et al. (2001) found lake variance to be the largest component of variance 

for zooplankton in the northeastern United States, followed by index variance. Efforts to reduce 

the magnitude of the factors that contribute to zooplankton variance can maximize one’s ability to 

detect differences or trends in the data.
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Figure 5-7. Zooplankton taxonomic richness versus ANC for a combined Adirondack dataset, based 

on 111 lake visits to 97 lakes in the EMAP, ELS, and STAR zooplankton surveys.

(Source: Sullivan et al. 2006.)
SECTION 6. TRANSITION PLAN

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SECTION 6. 

TRANSITION PLAN T.J. Sullivan and G.B. Lawrence

6.1 BACKGROUND

Under contract to the Forest Service, E&S Environmental Chemistry, Inc., and its research 

partners at Oregon State University, U.S. Geological Survey, and the University of Virginia have 

prepared new national FS protocols for the sampling, analysis, and quality assurance of the 

chemistry of lakes and streams on federal lands that are potentially sensitive to adverse impacts 

from atmospheric deposition of air pollutants. The protocols are also accompanied by a Training 

Plan to train field sampling staff in the field components of the new protocols (Section 7). The 

intention is to shift, where practical, FS survey and monitoring program efforts to the new 

protocols. It is hoped that this standardization of sampling and analysis protocols will improve 

comparability of the resulting data within and among regions and will improve the overall quality 

of the water chemistry data collected and analyzed for lakes and streams on National Forest 

System lands. 

It must be recognized, however, that there are risks associated with changing the protocols of an 

ongoing sampling program. In some cases, there may be a substantial period of record established 

for a particular lake or stream that is based on pre-existing protocols, and multiple waters within a 

particular wilderness may already have been surveyed and characterized with a particular set of 

protocols which may differ in important ways from the newly developed protocols. A change in 

approach may introduce bias into future efforts to examine patterns in water chemistry across 

time or across space. Therefore, changes must be carefully considered, and the likely results of 

those changes (if any) must be evaluated before making a wholesale change in sampling and/or 

analysis methods. This Transition Plan provides a framework for considering such protocol 

changes and their likely effects on the resulting data before full implementation of the new 

protocols. 

6.2 TRANSITION STEPS

This Transition Plan is divided into sequential steps to be followed in order to ascertain the 

likelihood that protocol changes might introduce bias into ongoing monitoring or characterization 

efforts. In some cases, the preferred approach might be to continue to monitor surface waters 

using existing protocols or to augment these protocols with additional elements from the new 
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168

protocols while retaining the basics of the existing protocols. In other cases, it might be best to 

transition to the new protocols after first evaluating the ramifications of methods changes. 

1. Read and become familiar with the new national protocols. Each FS field staff should 

read the field sampling protocol. Each FS staff person involved with the analysis, quality 

assurance, and/or interpretation of lake and stream water data should read all relevant 

sections from the National Protocol (field sampling, laboratory, QA/QC, data analysis, and 

biology). Each staff person should become thoroughly familiar with the portions of the new 

protocols that are relevant to his or her work duties and review existing SOPs in local FS 

survey and monitoring programs to determine all significant ways in which the new 

procedures differ from current practices. 

2. Attend a field sampling protocol training session. Anyone who is involved with the 

collection of water samples in the field should attend a training session to receive hands-on 

classroom and field training in the new field sampling protocols. 

3. Determine if any existing lake or stream monitoring sites will be dropped from the 

sampling program or if any new sites will be added. The new protocol emphasizes the 

need for matching sampling sites (and sampling schedules) with research questions and 

needs. Project managers should review existing monitoring programs to determine whether 

and to what extent the sites being sampled provide information required to achieve program 

goals. In some cases, a lake or stream may be included in a long-term monitoring effort, but 

available data might indicate that the water body is not very sensitive to the stressor(s) of 

concern, receives substantial inputs of geological S (which confound evaluation of effects 

of atmospheric S deposition), or is impacted by some form of disturbance to such an extent 

that it is not possible to ascertain the influence of air pollutants. Thus, based on 

considerations and priorities outlined in the National Protocols, the decision could be made 

to drop one or more sites from the monitoring program and/or to add others that might 

better meet program needs. However, before dropping a site from a long-term monitoring 

effort, one must carefully weigh the value of data that have been collected from that site to 

date versus the benefit of replacing that site with a new site that may have little or no data 

associated with it but will provide more useful information for the program in the future. 

4. Determine if sample collection protocols need to be changed. The new protocols could 

involve changes in any or all of the elements in where, what, when, and how to collect 

water samples. A change in collection location (Where) could affect the data even if this 

location is still considered the same site. Furthermore, a sample collected from a lake outlet 

could provide different data than a sample from the upper water column at the deepest 

portion of that same lake. If new field measurements or types of sample collection are 

added (What), the new procedures should be evaluated to ensure that they will not interfere 

with the existing field measurement and sample collection procedures. For example, if the 

existing sample collection location in a stream is downstream of the cross-section for new

flow velocity measurements, the water sample should be collected before the velocity 

measurements are taken to ensure that the water sample is not contaminated with suspended 

solids caused by stream wading. Changing the time of year or frequency (When) that the 

sample is collected could change the data record. Switching from spring sampling to 

summer sampling could bias the data towards lower flow conditions that might prevail 
SECTION 6. TRANSITION PLAN

169

during summer and that are typically less acidic than higher spring flow conditions. 

Switching from weekly sampling to monthly sampling would lower the sensitivity for 

detecting long-term trends. The procedures used to collect the samples (How) could also 

change the data. Water sampling devices that integrate flow or depth can produce different 

results than dipping a bottle. Collection and transport via syringe may yield different values 

for some parameters (i.e., pH and DIC) than collection and transport via bottle. 

Influences such as those described above should be evaluated in conjunction with changing 

methods in the middle of a monitoring program to determine if a sampling bias will be 

introduced. This should be done by performing the collection procedures with both the 

existing and new procedures for a length of time to account for the full variation in 

sampling conditions. In many cases, this could require a year or more of duplicating 

procedures. Results obtained using the original protocol should be compared to results 

obtained using the new protocol using a scatterplot with a 1:1 line added. If one approach 

yields results that are consistently either higher or lower than the other approach, the data 

points will plot consistently either above or below the 1:1 line. If there is no bias introduced 

by the change in protocol, the data points will be approximately evenly distributed above 

and below the 1:1 line. If it is determined that there is a bias introduced by the method 

change, then a decision will need to be made regarding whether to:

a. Stick with the original protocol;

b. Shift to the new protocol and ignore the difference if it is judged that the 

difference is too small to be of consequence to the intended use of the data; or.

c. Develop a regression approach to “correct” the data points obtained using the 

original method to more closely approximate the results obtained with the new 

(and presumably improved) method. 

In general, we recommend the first option unless there is a compelling reason to change. 

This is a judgment call, however, and any of these options can be reasonable depending 

upon circumstances. 

Changes in the manner in which the sample is collected that improve precision without 

adding bias, such as additional steps to prevent possible sample contamination during 

sampling, should not influence the decision as to whether or when to shift to the new 

protocols. An example of this type of change is in instituting the use of latex gloves during 

the sample collection. Any sampling procedure change that is expected to reduce the 

likelihood of sample contamination should be viewed as a positive step that should be 

implemented as soon as is practical. 

5. Evaluate the need for change in chemical measurements done in the laboratory or a 

change in the data quality objectives for the methods being used. If a new measurement 

is needed, the method should be fully evaluated to ensure that the desired results will be 

obtained. Also, if it is determined that a laboratory is unable to meet the data quality 

objectives needed for a FS characterization or monitoring program, an alternative 

laboratory will need to be found. If the laboratory needs to upgrade or replace existing 

instrumentation or to modify existing SOPs, they must provide duplicate results using 
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170

samples that are representative of the relevant study sites for a minimum of 100 samples to 

document that the changes have not introduced a bias. Because of potential methods 

interferences due to the mix of chemical constituents in a water sample, a laboratory 

methods change might alter results for one type of surface water but not another. 

6. Conduct side-by-side sampling and/or analysis to compare results obtained using 

initial protocols with results obtained using new protocols. Such side-by-side 

comparisons should be conducted when potentially significant changes are made in either 

field or laboratory protocols that could affect long-term continuous records. In this 

situation, the data measured with previous protocols will need to be married with the new 

data without bias, which could be misinterpreted as a real change over time. The timeframe 

over which the side-by-side comparisons should be conducted should include the full range 

of variability in the parameters of interest. For marrying long-term records for trends 

analysis, the minimum recommended length of time is 1 year. However, the length of time 

required for duplicate analyses is affected by the sampling frequency. Running duplicate 

sampling and analysis for 1 year would be adequate for a weekly sampling program but 

certainly not for a seasonal sampling program that collects only four times per year. Our 

overall recommendation is a minimum of 1 year and a minimum of 100 samples distributed 

across the various sampling sites. The side-by-side comparisons should be evaluated after 

sufficient data have been collected. The original protocol should not be dropped until the 

analysis of the side-by-side comparison is completed and the results indicate that the 

datasets based on the new and the old protocols can be married without creating artifacts in 

the record. 

7. Provide proper documentation to eliminate ambiguity in protocol applications. Clearly 

label samples collected with the new protocols and samples collected with the old protocols 

in such a way that the differences are documented and unambiguous. For example, if the 

decision is made to replace bottle sampling with syringe sampling for the measurement of 

water pH, procedures must be in place to document this change in the database; in this 

instance, the documentation for that sample in the collection method field of the database 

would indicate “lab pH - bottle” or “lab pH - syringe.” Details of each sampling method 

should also be documented in the field sampling protocols.

6.3 ANTICIPATED PROTOCOL CHANGES

We anticipate that a number of protocol changes will occur in the sampling that is conducted 

within the various FS regions as a result of adoption of the new National Protocols by the FS 

ARM program. The anticipated changes expected to be the most significant in terms of 

potentially introducing bias into the data and/or altering conclusions drawn from the data include 

the following:

FIELD SAMPLING

• Sample at index site near deepest portion of lake.

• Sample for pH and DIC using syringe or glass bottle with septum cap.

• Include discharge measurement.
SECTION 6. TRANSITION PLAN

171

LABORATORY

• Shift from field pH to lab pH measurements.

• Perform filtration in the laboratory.

• Change laboratory instrument(s) or analysis procedure(s).

• Provide documentation of detection and reporting limits.

• Provide full suite of QA and QC.

BIOLOGICAL MEASUREMENTS

• Change net mesh size.

• Change lab counting protocol.

• Change taxonomic resolution.

6.4 DECISION OF WHETHER OR NOT TO CHANGE 

PROTOCOLS 

FS staff should be very careful about methods changes that potentially could influence chemical 

results while in the midst of an inventory or monitoring program. Especially if trends analyses are 

planned for the resulting data, it is always important to “compare apples with apples.” In many 

cases, existing protocols, while not necessarily the preferred way of doing things, might best be 

left in place throughout the duration of a multi-year survey or of a long-term monitoring effort. 

Staff must recognize that a methods change with unquantified impacts on sampling results will 

compromise their ability to make comparisons across space and across time. Especially when a 

long period of monitoring record already exists, potential methods changes must be very carefully 

considered and thoroughly documented with side-by-side sampling and analysis in order to 

preserve the integrity of future data comparisons. If it is determined that a change in protocols 

will require an adjustment of data values obtained using the original protocol, then FS staff 

should consult with a statistician or scientist well-versed in this kind of data adjustment.

SECTION 7. TRAINING PLAN

173

SECTION 7. 

TRAINING PLAN T.J. Sullivan

7.1 BACKGROUND

Training people to collect water samples properly and to record all needed ancillary data is one of 

the most important aspects of a water survey, water quality characterization, or monitoring effort. 

This Training Plan provides guidance on how to conduct an effective training session for water 

sampling. It must be recognized that considerable personnel and financial resources go into field 

sample collection, laboratory analysis, QA/QC, and data analysis using the resulting data. If field 

staff are not attentive to detail or have not been properly trained, all this effort and expense can be 

wasted. All activities down the chain are dependent on collection of a good representative sample. 

This Training Plan is based on National Water Chemistry Field Sampling Protocols for Air 

Pollution Sensitive Waters, developed for the FS ARM program by E&S Environmental 

Chemistry, Inc., and collaborating scientists from Oregon State University, the U.S. Geological 

Survey, and the University of Virginia. Key elements of the Protocols are summarized in this 

Training Plan. The Trainer should understand all elements of the Protocols before planning or 

conducting a training session. The materials in the Protocols are detailed and complex. The 

Trainer should, therefore, plan to devote considerable time to becoming thoroughly familiar with 

the materials in the Protocols before conducting a training session. 

This Training Plan consists of an outline of the training materials and approach (this document), 

to be used in combination with a series of PowerPoint slides, handout materials, and three 

training videos. The slides and training videos serve as supporting materials. The slides will assist 

the trainer in covering all major issues in the training session. They will help to organize the 

presentation and facilitate discussion. The training videos are a more visual component and 

intended to augment the materials covered in the slides. 

The protocols address both lake and stream sampling. Consequently, this Training Plan also 

addresses both types of surface water. A trainer can choose to cover either one or both in the 

training session. Recognize that, if it is not possible to access the deep water index site at a given 

lake, lakes are sometimes sampled at their outlet stream location; outlet stream sampling follows 

stream (not lake) sampling protocols. Therefore, it may be desirable to train all sampling 

personnel in the stream sampling protocols. National forests in some regions of the country 

contain few or no lakes; in such regions, it may not be necessary to conduct any training with 

respect to lake sampling protocols. 
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Seasonal or temporary field staff and volunteers should attend the full training session before

beginning field work. It is recommended that all permanent or long-term field sampling personnel 

attend at least two full training sessions. Field personnel who sample year-after-year should 

attend the full session before each of the first two years of sampling. After that, attending annual 

training sessions could be optional for experienced samplers but they should annually review the 

PowerPoint slides, the stream sampling video, appropriate SOPs, and the lake sampling and 

discharge measurement videos (if applicable) at the beginning of each sampling field season. 

The training session outlined here consists of two parts: classroom training and field training. The 

classroom training support materials include a set of PowerPoint slides, series of handouts, a 

stream sampling video, a lake sampling video, and a discharge measurement video. Classroom 

training should take about 4 hours, allowing limited time for discussion. The basic field training 

can also be conducted in about 4 hours if the lake sampling component and the stream discharge 

aspects are omitted. Thus, it should be possible to complete the full training for stream sampling 

(omitting discharge measurement), including both classroom and field components, in 1 day. If 

the training includes field training in lake sampling and/or discharge measurement, a second day 

may be needed. 

7.2 PLANNING THE TRAINING SESSIONS

There are many things to consider before conducting a field training session for water sampling.

These include:

• How close to the actual sampling period do you conduct the training?

• Where will the training session be held? (Field training will require a representative sample 

site for demonstration purposes and practice). Select a training site that is as similar as 

possible to actual study sites. 

• What is an effective trainer/trainee ratio?

• Are the trainees new to water sampling or are they experienced? In other words: is this initial 

or refresher training?

• All sample teams should have equipment for the training. Will samplers bring their own

equipment or will it be distributed at the session?

• How will sample bottles, syringes, and coolers be distributed?

• Is all of the necessary equipment available at the training site?

• Which portions of the training will be included?

• Will the training session last one day or two? If two days of training are anticipated, what 

arrangements are needed for overnight accommodations?

• How many vehicles will be needed to transport personnel and gear to the field training 

locations? 
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The following major steps should be taken to prepare for the training session:

1. Set the date and inform the trainees. 

2. Make arrangements for overnight accommodations, if needed.

3. Order bottles and syringes from the lab. 

4. Make arrangements for obtaining classroom materials:

• Stream sampling video;

• Lake sampling video (optional);

• Discharge measurement video (optional);

• PowerPoint presentation;

• Projection equipment or viewing monitors for videos and PowerPoint;

• Sample binders with forms and form instructions (one per trainee plus extras);

• Backpack with sampling equipment, bottles, syringes, zipper lock bags, snap-on lid 

plastic containers for transporting syringes; and 

• Sample labels.

5. Cull the PowerPoint slides and handouts (if desired) to retain subsets of slides and handouts 

that are appropriate to your region, sampling needs, available training time, and 

expectations. Some important issues to consider in culling the slides and handouts include 

the following:

• Will you cover stream sampling, lake sampling, or both? Some regions and districts 

have few or no lakes, and, therefore, lake protocols are probably not relevant. If lakes 

are to be sampled, the preferred approach is to sample in deep water from a boat or 

float tube. However, an acceptable backup approach (if deep water sampling is not 

possible) is to sample from the outlet stream using stream sampling protocols. 

Therefore, all staff should be trained in stream sampling protocols; not all staff 

necessarily require training in lake sampling protocols. If you will conduct field 

training in both stream and lake methods, you will probably need more than a half-day 

for field training. 

• Will you conduct field training in discharge measurement? If so, you will need to spend 

several additional hours to do so. 

6. Make arrangements for field session material and logistics:

• Vehicles;

• Bottles and syringes; bags and plastic boxes for transport;

• Maps to locate training demonstration site(s);

• Sample labels;

• Forms and form instructions;

• Transport and access to training demonstration site(s);
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• Coolers;

• Miscellaneous equipment and supplies; and

• Ice.

7.3 CLASSROOM TRAINING

The classroom training addresses major considerations in why, where, what, when, and how to 

collect water samples. There are many issues to consider and multiple options for how to proceed. 

In general, sampling decisions reflect the purpose of the data collection effort. The intended use 

of the data has a major influence on how one should proceed in designing and carrying out a 

sampling program. This should be emphasized to trainees. There is a series of optional 

PowerPoint slides that covers various aspects of study design. You may choose to include all, 

some, or none of these slides. 

The first step in the classroom training is to emphasize to the trainees why it is important that they 

understand the protocols and why they need to take great care in their sampling and 

documentation efforts. The following questions should be addressed:

• Why do you have to be careful about site selection?

• Why do you need to be careful about when to sample?

• Why do you have to be careful about how you collect your sample?

• Why is attention to detail important?

After briefly reviewing the issues outlined above, the classroom training should focus on the 

PowerPoint presentation and/or training videos that can be found on the FS ARM program

website (<http://www.fs.fed.us/air>). The major additional items covered in the PowerPoint 

presentation and videos include: 

• Pre-trip activities;

• Site documentation; 

• Collection of water samples;

• Post sampling actions;

• Logistics of stream sampling and documentation;

• Stream discharge; and

• Logistics of lake sampling and documentation.

All trainees should watch the stream sampling, lake sampling, and/or discharge measurement 

videos. 
SECTION 7. TRAINING PLAN

177

7.3.1 Handouts

Handouts are an important part of training. The recommended handout materials for training are 

the forms and instructions (Appendix E), sample container labeling instructions (Appendix F), 

and the equipment checklists (Appendix H) at the end of this report.

Go over every line of the checklists, label, and forms with the trainees to ensure that they fully 

understand what to do before sampling, what to take with them during sampling, and what is 

required in the way of documentation. Trainees should be given a sample label and each of the 

water chemistry forms (with detailed instructions) for lake and/or stream sampling, as 

appropriate. The trainer should carefully go through the label and each of the forms to make sure 

that trainees fully understand what is required for completing each. 

7.3.2 Questions

Throughout the session and at the end of the classroom training session, ask questions of your 

trainees. Re-emphasize the importance of being careful to follow protocols and collect a good 

representative water sample. 

7.3.3 Prepare for Field Training

Finally, pack up the equipment for the field training session, check to make sure that you have all 

of the needed equipment, supplies, and forms, and pre-fill out paperwork, as needed, for field 

sampling. Be sure that each of the following issues is highlighted in both the classroom and field 

training:

• Importance of attention to details;

• Selection of sampling location;

• How to avoid contaminating your sample;

• How to fill out the forms and labels; and

• Importance of avoiding transmission of invasive species between sites.

7.4 FIELD TRAINING

Field training offers an opportunity to put into practice the things that you learned in the 

classroom training. The field training program should involve having each trainee conduct all 

activities that would normally be conducted during an actual sampling event. The field training 

should include each of the following major elements:

• Pack up gear that will be needed;

• Locate site;

• Collect stream sample;

• Collect lake sample (optional);

• Measure discharge (optional);
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• Clean up site, equipment, clothing, and boots;

• Labeling and documentation;

• Sample transport to vehicle; and

• Sample transport to lab.

Note that it may not be necessary to train field personnel in lake sampling protocols in some 

regions. Training in discharge measurement should also be considered optional. The field training 

session (without either lake sampling or discharge measurement) should take about four hours, 

assuming a very short hike to access the site. If you choose to include lake sampling and/or 

discharge measurement protocols, it might take a full day or two half-days for the field 

component of the training.
SECTION 8. REFERENCES

179

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Appendices

APPENDIX A. ACRONYMS AND ABBREVIATIONS ...................................................191

APPENDIX B. GLOSSARY ......................................................................................195

APPENDIX C. SAMPLING PROCEDURE FOR SURFACE WATER CHEMISTRY ............203

APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING 

ACTIVITIES ......................................................................................211

APPENDIX E. DATA ENTRY FORMS AND INSTRUCTIONS FOR FIELD SAMPLING 

ACTIVITIES ......................................................................................247

APPENDIX F. LABELING INSTRUCTIONS FOR FIELD SAMPLE CONTAINERS ...........271

APPENDIX G. EXAMPLE STANDARD OPERATING PROCEDURES FOR LAB ANALYSIS 

PROTOCOLS .....................................................................................275

APPENDIX H. TRAINING CHECKLISTS ...................................................................313

APPENDIX I. JOB HAZARD ANALYSIS (JHA) .......................................................319

APPENDIX A. ACRONYMS AND ABBREVIATIONS

191

APPENDIX A. 

ACRONYMS AND ABBREVIATIONS 

Al Aluminum

Al3+ Free aluminum (uncomplexed, trivalent)

Ali Inorganic monomeric aluminum

Alm Total monomeric aluminum

Alo Non-labile monomeric (presumed organically complexed) aluminum

Al(F)2

+ An aluminum fluoride molecule

AlF2+ An aluminum fluoride molecule

Al(OH)2+ An aluminum hydroxide molecule

Al(OH)2

+ An aluminum hydroxide molecule

ANC Acid neutralizing capacity

ANCOVA Analysis of covariance

ANOVA Analysis of variance

AQRV Air quality related values

ARML Air Resource Management Laboratory

BCS Base cation surplus

BMP Best management practices

BS Base saturation of the soil

Ca2+ Calcium

CEC Cation exchange capacity 

Cl- Chloride

CL

CO2

Confidence level

Carbon dioxide

CPR Cardio-pulmonary resuscitation

CQCCS Calibration quality control check sample
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CV Coefficient of variation

DAP Data analysis protocols

DIC Dissolved inorganic carbon

DIW Deionized water 

DO Dissolved oxygen

DOC Dissolved organic carbon

DON Dissolved organic nitrogen

DQO Data quality objective

ELAP Environmental Laboratory Accreditation Program

EMAP-SW Environmental Monitoring and Assessment Program-Surface Waters

EPA U.S. Environmental Protection Agency

EPT Ephemeroptera-Plecoptera-Tricoptera Index

F Fluorine

Fe Iron

FS U.S. Forest Service

GIS Geographical information system

GPS Global positioning system

H+ Hydrogen ion

HCl Hydrochloric acid

HCO3

- Bicarbonate

HDPE High density polyethylene 

Hg Mercury

IBD Ion balance difference

IBI Index of Biotic Integrity

IQR Interquartile range

JHA Job Hazard Analysis 

K+ Potassium

LDPE Low density polyethylene

LIMS Laboratory Information Management System

m Meter
APPENDIX A. ACRONYMS AND ABBREVIATIONS

193

MAGIC Model of Acidification of Groundwater in Catchments – a watershed ion 

balance model

MCV Median concentration value

MDL Method detection limit

Mg2+ Magnesium

ml Milliliters

MPV Most probable value

N Nitrogen

Na+ Sodium

NELAC National Environmental Laboratory Accreditation Conference

NELAP National Environmental Laboratory Accreditation Program

NH4

+ Ammonium

NIST National Institute of Standards and Technology

NO3

- Nitrate

NRM National resource manager

NTU Nephelometric turbidity units

NWRI Environment Canada’s National Water Research Institute

P Phosphorus

PDE Percent difference in enumeration

PE Performance evaluation

PFD Personal floatation device

PnET-BGC Photosynthesis and Evapotranspiration – Biogeochemistry Model; A model of 

water, carbon, and nitrogen balance, coupled with a biogeochemistry model

ppm Parts per million

PTD Percent taxonomic disagreement

QA/QC Quality assurance/quality control

QCCS Quality control check sample

S Sulfur

SBC Sum of base cations

Si Silicon

SKT Kendal tau test

SLR Simple linear regression
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SO4

2- Sulfate

SOP Standard operating procedures

SRP Soluble reactive phosphorus

SRS Standard reference sample

SSN Sample serial number

TIME Temporally Integrated Monitoring of Ecosystems program

TOC Total organic carbon 

TSS Total suspended solids

USFS U.S. Forest Service

USGS U.S. Geological Survey

WTRS Surface water sample depth zone: lake/stream water surface and water 

subsurface

μeq/L Microequivalents per liter

μg/L Micrograms per liter

μM Micromoles per liter

μS/cm Microsiemens per cm
195

APPENDIX B. 

GLOSSARY 

Acid anion Negatively charged ion that does not react with hydrogen 

ion in the pH range of most natural waters. 

Acid neutralizing capacity (ANC) The equivalent capacity of a solution to neutralize strong 

acids. The components of ANC include weak bases 

(carbonate species, dissociated organic acids, aluminohydroxides, borates, and silicates) and strong bases 

(primarily OH-

). ANC can be measured in the laboratory 

by the Gran titration procedure or defined as the 

difference in the equivalent concentrations of the base 

cations and the mineral acid anions. It is a key indicator 

of the ability of water to neutralize the acid or acidifying 

inputs it receives. This ability depends largely on 

associated biogeophysical characteristics. 

Acid-base chemistry The reaction of acids (proton donors) with bases (proton 

acceptors). In the context of this report, this refers to the 

reactions of natural and anthropogenic acids and bases, 

the result of which is described in terms of pH and acid 

neutralizing capacity of the system. 

Acidic deposition Transfer of acids and acidifying compounds from the 

atmosphere to terrestrial and aquatic environments via 

rain, snow, sleet, hail, cloud droplets, particles, and gas 

exchange. 

Acidic lake or stream A lake or stream in which the acid neutralizing capacity is 

less than or equal to 0. 

Acidification The decrease of acid neutralizing capacity in water or 

base saturation in soil caused by natural or anthropogenic 

processes. 

Acidified Pertaining to a natural water that has experienced a 

decrease in acid neutralizing capacity or a soil that has 

experienced a reduction in base saturation. 
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Algae Photosynthetic, often microscopic and planktonic, 

organisms occurring in marine and freshwater 

ecosystems. 

Algal bloom A reproductive explosion of algae in a lake, river, or 

ocean. 

Alpine The biogeographic zone made up of slopes above the tree 

line and characterized by the presence of rosette-forming 

herbaceous plants and low, shrubby, slow-growing 

woody plants. 

Analyte A chemical species that is measured in a water sample. 

Anion A negatively charged ion. 

Anthropogenic Of, relating to, derived from, or caused by humans or 

related to human activities or actions. 

Atmosphere The gaseous envelope surrounding the Earth. The dry 

atmosphere consists almost entirely of nitrogen and 

oxygen, together with trace gases, including carbon 

dioxide and ozone. 

Autoecology Study of the ecology of individual species, as opposed to 

the entire community of species. 

Base cation An alkali or alkaline earth metal cation (Ca2+, Mg2+

, K+, 

Na+). 

Base saturation The proportion of total soil cation exchange capacity that 

is occupied by exchangeable base cations; i.e., by Ca2+, 

Mg2+, K+

, and Na+. 

Benthic macroinvertebrates Animals without backbones that inhabit the bottom 

substrates of streams

Bioaccumulation The phenomenon wherein toxic elements are 

progressively amassed in greater qualities as individuals 

farther up the food chain ingest matter containing those 

elements. 

Biological effects Changes in biological (organismal, populational, and 

community-level) structure and/or function in response to 

some causal agent; also referred to as biological response. 
APPENDIX B. GLOSSARY

197

Calibration Process of checking, adjusting, or standardizing operating 

characteristics of instruments or coefficients in a 

mathematical model with empirical data of known 

quality. The process of evaluating the scale readings of an 

instrument with a known standard in terms of the physical 

quantity to be measured. 

Catchment An area that collects and drains rainwater (also called 

“watershed”). 

Cation A positively charged ion. 

Cation exchange capacity The sum total of exchangeable cations that a soil can 

adsorb. 

Chronic acidification The decrease of acid neutralizing capacity in a lake or 

stream over a period of decades or longer, generally in 

response to gradual leaching of ionic constituents. 

Circumneutral Close to neutrality with respect to pH (neutral pH = 7); in 

natural waters, pH 6-8. 

Climate Climate in a narrow sense is usually defined as the 

‘average weather’ or more rigorously as the statistical 

description in terms of the mean and variability of 

relevant quantities over a period of time ranging from 

months to thousands or millions of years. These quantities 

are most often surface variables such as temperature, 

precipitation, and wind. Climate in a wider sense is the 

state, including a statistical description, of the climate 

system. The classical period of time is 30 years, as 

defined by the World Meteorological Organization. 

Coarse stream substrate Cobble to boulder (tennis ball to car sized; 64 mm to 

4000 mm) size substrate. 

Critical load A quantitative estimate of an exposure to one or more 

pollutants below which significant harmful effects on 

specified sensitive elements of the environment do not 

occur according to present knowledge. 

Decomposition The microbially mediated reaction that converts solid or 

dissolved organic matter into its constituents (also called 

decay or mineralization). 

Dissolved inorganic carbon The sum of dissolved (measured after filtration) carbonic 

acid, bicarbonate, and carbonate in a water sample. 
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198

Dissolved organic carbon Organic (derived from the breakdown of plant or animal 

material) carbon that is dissolved or unfilterable (0.45-μm 

pore size) in a water sample. 

Drainage lake A lake that has a permanent surface water inlet and outlet. 

Ecosystem The interactive system formed from all living organisms 

and their abiotic (physical and chemical) environment 

within a given area. Ecosystems cover a hierarchy of 

spatial scales and can comprise the entire globe, biomes 

at the continental scale, or small, well-circumscribed 

systems, such as a small pond. 

Epilimnion The layer of water in a thermally stratified lake that lies 

above the thermocline, is circulating, and remains 

perpetually warm.

Episodic acidification The short-term decrease of acid neutralizing capacity 

from a lake or stream. This process has a time scale of 

hours to weeks and is usually associated with 

hydrological events. 

EPT Index Index of taxonomic richness of three insect orders 

(Ephemeroptera, Plecoptera, and Tricoptera)

Eutrophication The process whereby a body of water becomes overenriched in nutrients, resulting in increased productivity 

(of algae or aquatic plants) and sometimes also decreased 

dissolved oxygen levels. 

Evapotranspiration The process by which water is returned to the air through 

direct evaporation or transpiration by vegetation. 

Fine/sand stream substrate Stream substrate not gritty (silt/clay/muck < 0.06 mm 

diameter) to gritty (up to ladybug sized; 2 mm diameter) 

substrate.

Glide Water moving slowly along stream channel, with smooth, 

unbroken surface; low turbulence.

Gran analysis A mathematical procedure used to determine the 

equivalence points of a titration curve for acid 

neutralizing capacity. 

Gravel stream substrate Fine to coarse gravel (ladybug to tennis ball sized; 2 mm 

to 64 mm diameter) size substrate.

Ground water Water in a saturated zone within soil or rock. 
APPENDIX B. GLOSSARY

199

Hindcast To estimate the probability of some past event or 

condition as a result of rational study and analysis of 

available data. 

Hydrologic(al) event Pertaining to increased water flow or discharge resulting 

from rainfall or snowmelt. 

Hydrologic flow paths Surface and subsurface routes by which water travels 

from where it is deposited by precipitation to where it 

drains from a watershed. 

Hydrology The science that studies the waters of the earth—their 

occurrence, circulation, and distribution; their chemical 

and physical properties; and their reaction with their 

environment, including their relationship to living things. 

Hypolimnion The layer of water in a thermally stratified lake that lies 

below the thermocline, is noncirculating, and remains 

perpetually cold.

Index of Biotic Integrity (IBI) Provides assessment of biological condition based on a 

combination of metrics. 

Invasive species A species aggressively expanding its range and 

population density into a region in which it is not native, 

often through outcompeting or otherwise dominating 

native species. 

Labile monomeric aluminum Operationally defined as aluminum that does not pass 

through a cation exchange column; assumed to represent 

inorganic monomeric aluminum (Ali).

Leaching The removal of soil elements or applied chemicals by 

water movement through the soil. 

Macrophyte Rooted aquatic plant.

MAGIC Model of Acidification of Groundwater in Catchments – a 

watershed ion balance model.

Mitigation Generally described as amelioration of adverse impacts 

caused by a stressor such as acidic deposition at the 

source (e.g., emissions reductions) or the receptor (e.g., 

lake liming). 

Model An abstraction or representation of a system, generally on 

a smaller scale. 
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200

Monomeric aluminum Aluminum that occurs as a free ion (Al3+), simple 

inorganic complexes (e.g., Al(OH)n

3-n

, AlFn

3-n

), or simple 

organic complexes, but not in polymeric forms; 

operationally defined as extractable aluminum measured 

by the pyrocatechol violet method or the methyl-isobutyl 

ketone method (also referred to as the "oxine" method) is 

assumed to represent total monomeric aluminum. 

Monomeric aluminum can be divided into labile and nonlabile components using a cation exchange column.

Non-labile monomeric aluminum Operationally defined as aluminum that passes through a 

cation exchange column and is then measured by one of 

the two extraction procedures used to measure 

monomeric aluminum; assumed to represent organic 

monomeric aluminum (Alo). 

Occult deposition The removal of gases and particles from the atmosphere 

to surfaces by fog or mist.

Organic acids Heterogeneous group of acids generally possessing a 

carboxyl (-COOH) group or phenolic (C-OH) group. 

Parameter (1) a characteristic factor that remains at a constant value 

during the analysis, or (2) a quantity that describes a 

statistical population attribute. 

pH The negative logarithm of the hydrogen ion activity. The 

pH scale is generally presented from 1 (most acidic) to 14 

(most alkaline); a difference of one pH unit indicates a 

tenfold change in hydrogen ion activity. 

Phytoplankton The plant-like forms of plankton. These single-celled 

organisms are the principal agents of photosynthetic 

carbon fixation in some fresh waters. 

Plankton Small (often microscopic) plant-like or animal species 

that spend part or all of their lives in open water.

PnET-BGC Photosynthesis and Evapotranspiration Biogeochemistry 

Model: a model of water, carbon, and nitrogen balance, 

coupled with a biogeochemistry model.

Pool In ecological systems, the supply of an element or 

compound, such as exchangeable or weatherable cations 

or adsorbed sulfate, in a defined component of the 

ecosystem. 
APPENDIX B. GLOSSARY

201

Population For the purpose of this report, 1) the total number of lakes 

or streams within a given geographical region or the total 

number of lakes or streams with a given set of defined 

chemical, physical, or biological characteristics; or 2) an 

assemblage of organisms of the same species inhabiting a 

given ecosystem. 

Precision A measure of the capacity of a method to provide 

reproducible measurements of a particular analyte (often 

represented by variance). 

Primary Productivity All forms of production accomplished by plants.

Quality assurance A system of activities for which the purpose is to provide 

assurance that a product (e.g., data base) meets a defined 

standard of quality with a stated level of confidence. 

Quality control Steps taken during sample collection and analysis to 

ensure that data quality meets the minimum standards 

established in a quality assurance plan. 

Rapid Water movement along stream channel is rapid and 

turbulent; surface with intermittent “white water” with 

breaking waves; characterized by a continuous rushing 

sound.

Reachwide sample All kick net samples collected at the 11 transects 

combined into a single composite sample. 

Riffle Water moving along stream channel, with small ripples, 

waves, and eddies; waves not breaking, and surface 

tension is not broken; characterized by a “babbling” or 

“gurgling” sound. 

Scenario One possible deposition sequence following 

implementation of a control or mitigation strategy and the 

subsequent effects associated with this deposition 

sequence. 

Sensitivity For this report, the degree to which a system is affected, 

either adversely or beneficially, by an effect of NOx

and/or SOx pollution (e.g. acidification, N-nutrient 

enrichment, etc.). The effect may be direct (e.g., a change 

in growth in response to a change in the mean, range, or 

variability of N deposition) or indirect (e.g., changes in 

growth due to the direct effect of N consequently altering 

competitive dynamics between species and decreased 

biodiversity).
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Species richness The number of species occurring in a given ecosystem, 

generally estimated by the number of species caught and 

identified using a standard sampling regime. 

Specific conductance The conductivity between two plates with an area of 1 

cm2 across a distance of 1 cm at 25o C. Provides an index 

of the ionic strength of a water sample. 

Steady state The condition that occurs when the sources and sinks of a 

property (e.g., mass, volume, concentration) of a system 

are in balance (e.g., inputs equal outputs; production 

equals consumption). 

Stream flow Water flow within a river channel, for example, expressed 

in m3/s or cfs (cubic feet per second). A synonym for 

river discharge. 

Subpopulation Any defined subset of the target population. 

Support reach The length of stream to be sampled at a sampling 

location.

Total monomeric aluminum Operationally defined as the simple unpolymerized form 

of aluminum present in inorganic or organic complexes. 

Turnover The interval of time in which the density stratification of 

a lake is disrupted by seasonal temperature variation, 

resulting in entire water mass becoming mixed. 

Variable A quantity that may assume a numeric value during 

analysis. 

Watershed The geographic area from which surface water drains into 

a particular lake or point along a stream. 

X-site Stream sampling location.

Zooplankton The animal forms of plankton. Zooplankton include 

crustaceans, rotifers, pelagic (open water) insect larvae, 

and aquatic mites.
203

APPENDIX C. 

SAMPLING PROCEDURE FOR SURFACE 

WATER CHEMISTRY 

Robert Musselman

US Forest Service, Rocky Mountain Research Station

This sampling procedure provides guidance in container handling for stream sampling projects 

conducted by the Rocky Mountain Research Station. It is intended for training purposes and may 

be augmented as needed with requirements from formal Standard Operating Procedures (SOPs).

LOCATE SAMPLE COLLECTION SITE

Streams are always sampled upstream from any manmade structure such as a bridge, culvert, or 

flume. Lakes are sampled at their outlet unless the chemistry of the lake profile is needed. 

Approach the sampling location from downstream if possible. Choose a sampling spot that is fastmoving, at least 15–20 cm (6–8 inches) deep if possible, and where you can reach it from a solid 

place on the stream bank (a rocky, not soft or spongy, spot) or from a large rock.

Collect from the same sampling site each time if collections are repeated over time. Check the 

previous collection field notes for exact sampling location. A GPS identification of the location 

longitude/latitude/elevation is helpful.

TAKE WATER TEMPERATURE

Place a thermometer in the water near the sampling point—preferably downstream. Avoid 

disturbing the bottom at the sample site. Allow the thermometer to equilibrate (reach stream 

temperature and stop changing readings) and record the temperature. It may take a couple of 

minutes for the thermometer to equilibrate. Digital thermometers equilibrate faster and it is easier 

to determine when they have reached water temperature.

LABEL BOTTLE CORRECTLY

Sample bottles are typically brown, 250 ml (about 1 cup volume), high-density polyethylene 

(HDPE) or low-density polyethylene (LDPE) plastic, unless otherwise provided. Before 

immersion of the sample bottle, use a black permanent marker to write on the dry bottle the 

sample location (geographic area name and stream or lake name—e.g., Indian Peaks Wilderness, 
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204

Blue Lake), date and year, time of day (indicate daylight or standard time as MDT or MST for 

Mountain Daylight Savings Time or Mountain Standard Time), water temperature (add o

F or oC),

and sampler's first and last names.

PUT ON GLOVES AND RINSE BOTTLE AND GLOVES

After the bottle is labeled, put on the gloves (powder-free latex or nitrile) and rinse the capped 

bottle and the gloves downstream to remove any possible contaminants. Be careful not to touch 

your face, the ground, clothing, or anything except the bottle and cap after the gloves and bottle 

are rinsed. Fingers contain contaminants such as nitrates, and insect repellents, sunscreen, and 

cigarette smoke are particularly troublesome as contaminants. Avoid breathing directly in/on the 

bottle or cap during sample collection.

PREPARE BOTTLE FOR SAMPLING

First, rinse the inside of the bottle, the inside of the cap, and then rim of the bottle with the 

deionized water (DIW) that comes in the bottle. To rinse, remove the cap with index finger and 

thumb, palm facing up. If the cap is not held correctly when removed, there is an increased 

possibility of pouring water over your hand and into the cap or bottle when rinsing. 
APPENDIX C. SAMPLING PROCEDURE FOR SURFACE WATER CHEMISTRY

205

To remove the cap for rinsing: DON’T have your palm facing the bottle. DO have your palm facing 

AWAY from the bottle.

Once the cap is removed, turn your palm down to face the cap up, fill the inside of the cap with 

DIW, and empty it over the rim and threads of the bottle, pointing the mouth of the bottle 

downward and being careful not to pour water over your finger and/or thumb or back into bottle.

WRONG 

WRONG 

RIGHT! 

RIGHT! 
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206

To hold cap for rinsing: DON’T have your palm facing up; DO have your palm facing DOWN.

To collecting rinse water in cap, keep fingers and palm BELOW and OUT OF THE WAY. 

Rinse the rim of the bottle with water from the cap, with fingers and palm OUT OF THE WAY and the 

bottle opening DOWN.

WRONG 

RIGHT! 

RIGHT! 

RIGHT! 
APPENDIX C. SAMPLING PROCEDURE FOR SURFACE WATER CHEMISTRY

207

Rinse the cap and bottle rim about six times, then completely empty the bottle of any remaining 

DIW. Pour discarded DIW downstream or off-stream on land away from the sampling site. After 

the final rinse, place cap back on and tighten it 1⁄4 turn, hold the cap on the bottle with index 

finger of the hand holding the bottle. Now the sample rinses can be collected.

RINSE THE BOTTLE WITH SAMPLE WATER

After DIW rinsing, the bottle is rinsed again with water from the sample location to remove any 

DIW left in the bottle. This ensures that the bottle will eventually contain only sample water for 

analysis. Collect water for the rinses in the same way as collecting the sample afterward to ensure 

good habits in practice.

When collecting water on the left side of stream (facing upstream), hold the bottle with your right 

hand, cap pointing upstream. Remove cap with the index finger and thumb of your left hand, with 

your palm placed under the bottle (palm and other fingers should not be upstream of the bottle). 

When collecting water on the right side of stream (facing upstream), hold the bottle with your left 

hand, cap pointing upstream. Remove the cap with the index finger and thumb of your right hand,

with your palm placed under the bottle (palm and other fingers should not be upstream).

Whether you are on the right or left side of the stream (facing upstream) and whether you are 

holding the bottle in your right hand or left hand, always be careful that the hand holding the 

bottle and your palm and fingers of the other hand tightening and loosening the cap are not 

upstream of the bottle opening. 

Move to your sampling point and reach as far into stream as possible. Hold the cap on the bottle 

with your index finger. 

To handle the sample bottle in preparation for sampling, 

hold the cap on with your index finger.

Immerse the bottle completely about 10 cm (4 inches) deep or half-way to the bottom if the 

stream is shallow (be sure to write the depth the sample was collected on the field notes after the 
NATIONAL PROTOCOLS FOR SAMPLING AIR-POLLUTION-SENSITIVE WATERS

208

sample is collected and secured). If the stream is too shallow to immerse the bottle fully, immerse 

it as far as possible and collect as much sample water as possible, being very careful not to touch 

the bottom where sediments can be disturbed and making sure that no surface film flows into the 

bottle. A syringe (appropriately rinsed at least three times with DIW) may be used to fill the 

bottle if necessary, but it is best to collect the sample directly into the bottle. If a syringe is used,

be sure to document this in the field notes.

Place the bottle flat on its side underwater, pointing the mouth of the immersed bottle upstream. 

Place the thumb and forefinger of your free hand on the cap before removing the index finger 

holding the cap on the bottle. Remove the cap with your thumb and index finger, with the palm of 

that hand placed under or over the bottle to make sure the palm is not upstream of the bottle.

To remove cap under water: DON’T have your hand upstream of the bottle; 

DO keep your palm UNDER or BESIDE the bottle.

Be very careful that any water entering bottle does not touch your gloves on its way toward the 

bottle by making sure your hands and fingers are not held upstream of the bottle when the cap is 

removed. Move the cap enough to let water enter the bottle. Fill the bottle about half full. It is not 

necessary to fill the bottle completely for rinses. Place the cap back on loosely (1⁄4-turn) 

underwater, holding cap on with index finger of the hand holding the bottle. Always remove and 

replace the cap underwater.

Remove the immersed and capped bottle from the stream and shake it. It is okay if some water 

leaks out rinsing the rim during shaking. The rinse procedure is then the same as described for 

DIW. 

Rinsing must be conducted downstream of the sample site. Rinse the cap and rim three times,

being careful that the rinse water does not touch your gloves. The bottle opening should be 

pointed downward during rinses to avoid rinse water flowing back into the bottle. This is easier if 

the bottle is only half-filled (or half-dumped) for rim/cap rinses. After the three separate cap/rim 

rinses, pour out any remaining rinse-water downstream of the sample point. Do not touch the 

bottle mouth or the inside of the cap.

WRONG 

RIGHT! 
APPENDIX C. SAMPLING PROCEDURE FOR SURFACE WATER CHEMISTRY

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COLLECT THE SAMPLE

After three stream-water rinses, collect the final sample on the fourth immersion. Use the same 

procedure as before, but fill the bottle completely. 

Tip the bottle mouth up but still underwater to remove all air bubbles from the bottle before 

capping. If necessary because the stream is shallow, squeeze the bottle slightly as the cap is 

tightened so that no air remains in bottle. Be sure that the cap is on tight, first underwater and 

again when the bottle is taken out of the stream.

Be especially careful not to contaminate the sample with surface film or bottom sediment. If this 

happens, discard the sample and start the procedure over with three more stream-water rinses. 

If the stream is too shallow to immerse bottle fully when tipped up for filling, collect as much as 

possible, being very careful not to touch the surface or bottom. Note the sampling depth on field 

notes.

SEAL AND STORE BOTTLE FOR TRANSPORTATION

Once the sample is collected, seal the sample bottle immediately in a zipper-lock bag, place the 

bag in a cooler, and keep the bag cold with frozen ice-packs or snow (if using snow, place the 

snow in two nested zipper-lock bags). Do not place snow or ice in the same bag as the sample. Do 

not expose sample bottles to the sun.

COMPLETE MISCELLANEOUS POST-SAMPLING TASKS

After all samples are secured in the cooler, remove the gloves, rinse them in the stream, and place 

them in a zipper-lock bag. Before leaving the field site, make sure the field data sheet is 

completed. Note any noteable weather conditions such as wind or rain. Measure the air 

temperature (the thermometer must be shaded) and record it. When you return from the field, 

remove the gloves from their bag, rinse the gloves (with DIW if available), and dry them out for 

reuse.

SHIPPING INSTRUCTIONS

Samples are filtered in the lab, so no further processing is typically necessary in the field. Handdelivery to the lab is preferred, either by the sampling team or through lab pick-up arrangements. 

Keep samples cool during shipment. Ship them overnight to the pre-assigned lab in a hard-sided

cooler with frozen icepacks via FedEx or UPS. Labs are not typically open on weekends or 

holidays, so do not ship the samples unless they can arrive at the lab on a workday, Monday 

through Friday. If you need to store the samples before shipping them, keep them refrigerated at 

39-40o

F (4o C) but do not allow them to freeze. Ship them using an afternoon pickup for next-day 

arrival if possible to shorten shipping time. 
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COLLECT DUPLICATE AND BLANK SAMPLES

Collect a duplicate sample if so instructed. Generally, every 10th to 15th sample collected is 

replicated (duplicated). Sample sites chosen for duplicate sampling are selected at random among 

streams or lakes sampled. If a duplicate is required for your site, repeat the same procedures as 

with the normal stream samples. The duplicate is the second of the two samples collected. Write 

“DUPL” on the sample bottle before the sample is collected. Duplicates document the 

repeatability of individual sample collections and reproducibility of laboratory results. 

Take a field blank sample bottle to the field if so instructed. The field blank (FB) remains 

unopened. Field blanks are included in the analysis to quantify chemicals from non-sample 

sources such as water bottles, DIW, filter paper, handling procedures, etc. Write the name of the 

sample study area, stream or lake name, date and year, sampler's name, and “FB” on the bottle. 

The FB bottle is never opened in the field; however, the extra FB bottle can be used in place of a

sample bottle in an emergency: for example, if a sample bottle or its cap is lost or contaminated. 

The FB is not as vital to the analysis as the samples. 

ACKNOWLEDGEMENTS

Thanks to Andrea Holland-Sears for the photos taken during a training session August, 2011, to 

Dave Richie for providing the demonstration for those photos, and to Cass Cairns for the rimrinsing photo.

INFORMATION SOURCES FOR WILDERNESS LAKE SAMPLING

Where to find blank field sampling forms:

Go to: <http://www.fs.fed.us/air/> and click on: Forms.

Where to find field sampling protocols:

Go to: <http://www.fs.fed.us/air/> and click on: Sampling Protocols.

Where to obtain water bottles and where to ship samples:

Louise O'Deen 

Lab Manager 

USFS/USGS Water Chemistry Laboratory

2150 Centre Avenue, Building C 

Fort Collins, CO 80526-8118

Shipping notes:

Ship your samples by overnight UPS or FedEx Monday through Thursday for workday arrival. 

Samples should be shipped in a cooler with icepacks. Do not ship on Friday. The lab is not open 

weekends. Keep samples refrigerated and ship Monday. 

For more information, contact: 

Louise O’Deen 

Email: Louise_O'Deen@usgs.gov

Phone: (970) 226-9190 
211

APPENDIX D. 

STANDARD OPERATING PROCEDURES FOR 

FIELD SAMPLING ACTIVITIES 

D-1 STANDARD OPERATING PROCEDURE FOR STREAM 

SAMPLING

This Standard Operating Procedure (SOP) provides guidelines for stream sampling within the FS 

ARM program. It is intended as a Generic SOP, suitable for adoption as a stand-alone procedure, or 

for modification to fit local program needs. It is divided into individual sections that cover pre-trip 

activities, sampling site documentation, stream sampling and sample handling, measurement of 

stream discharge, post-trip activities, and needed equipment and supplies.

D-1.1 Pre-Trip Activities

Field teams conduct a number of activities in their office or at a base site. These include tasks that 

must be completed both before departure to the sampling site and after return from the site. This 

section describes pre-trip procedures for office and base site activities that should be carried out in 

support of stream sampling. 

Pre-departure activities include development of sampling itineraries, instrument calibration if 

appropriate, equipment checks and repair, supply inventories, and sample container preparation. 

Procedures for these activities are described in the following sections. An example checklist for 

materials and supplies is given in Table D-1. Use this checklist to ensure that equipment and supplies 

are organized and available at the stream site in order to conduct the activities efficiently. Remember 

to take any safety equipment required by your unit (e.g., hard hats, radios, or cell phones).
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Table D-1. Checklist of materials and supplies for stream sampling site visits.

Standard Items: 

Collection permits and entry permits, if required

Site documentation forms (for new sites)

Clipboard

Site documentation reports (compiled as folders for existing sites)

Stream sampling record forms

Insulated container with ice or frozen refrigerant (packed in sealed plastic bags or other containers)

Small insulated container (with ice) for hike-in sites

Watch for recording time

Digital field camera with free memory and extra charged battery

GPS unit with extra batteries

Compass

Field thermometer (with string attached)

Pre-processed sample bottle(s) with completed sample label attached. Include a second bottle if sampling at that site is 

to be replicated. Put each bottle in a clean plastic zipper-lock bag.

Plastic gloves in sealed plastic bag

60 mL plastic syringes (with Luer type tip) with completed sample labels attached. Plastic container with snap-on lid to 

hold filled syringes

Syringe valves (Mininert® with Luer type adapter, or equivalent, available from a chromatography supply company)

Water Chemistry labels (if not already filled out and attached to sample containers at base site)

Soft-lead pencils and write in rain type pens for filling out field data forms and notebook entries

Fine-tipped indelible markers for filling out labels

Roll or box of tape strips

Field operations and methods documents

First aid kit

Backpack

Extra zipper-lock bags

Optional Items (may be required for specific studies):

60 mL glass bottles with septum caps and with completed sample labels attached

Calibrated multiparameter sonde, data logger and cable, with extra batteries

Calibration standards, quality control check samples, DIW, rinse bottles, waste tray and container, calibration cup, and 

sensor guard for sonde (multiple sensors combined in a unit that is lowered into the water)

Sonde calibration and post-calibration record forms

Measurement tape

Waders or high-top water proof boots for wading

Clear packaging tape to cover labels

Dissolved oxygen (DO)/temperature meter with probe

DO repair kit containing additional membranes and probe filling solution

Conductivity meter with probe
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

213

PREPARATION OF SAMPLE CONTAINERS

Before leaving the base location, package the sample containers, typically two sample bottles and two 

60 mL syringes for each site to be sampled, plus backup bottles and syringes in the event that one is 

lost or contaminated. Make sure to have plastic containers for transport of filled syringes from the 

field to the laboratory. Fill out a set of water chemistry sample labels and attach a completed label to 

each sample bottle and/or syringe. Make sure the syringe labels do not cover the volume gradations 

on the syringe. Place each sample bottle in a separate zipper lock bag. Finally, make sure that ice and 

or refrigerant for shipment to the lab is frozen or freezing so that it will be ready when you return 

from the field to the base location with the samples.

DAILY ITINERARIES 

Field sampling efforts should include a Project Leader who guides activities in the field, and a Project 

Coordinator who remains in the office during the sampling effort. The Project Leader reviews each 

site folder to ensure that it contains the appropriate maps, contact information, copies of access 

permission letters (if needed), and access instructions. Additional activities can include confirming 

the best access routes, calling landowners or local contacts (if applicable), confirming lodging or 

camping plans and locations (with directions), and coordinating rendezvous locations with 

individuals who must meet with field teams before accessing a site. This information is used to 

develop an itinerary. 

The Project Leader should provide the Project Coordinator with a schedule for each day of sampling. 

Schedules include departure time, estimated duration of sampling activities, routes of travel, and 

estimated time of arrival at the sampling site(s) and return to the base site. Changes that might be 

made to the itinerary should be relayed by the Project Leader to the Project Coordinator as soon as 

possible. Miscommunications can result in the initiation of expensive search and rescue procedures 

and disruption of carefully planned schedules. 

INSTRUMENT CHECKS AND CALIBRATION 

If appropriate, each field team should test and/or calibrate field instruments before departure for the 

sampling site. Such testing may be appropriate for dissolved oxygen (DO) meters, global positioning 

system (GPS) units, and perhaps other instrumentation. Batteries should be checked before departure 

for field sites. Extra batteries should be carried. 

Field personnel should check the inventory of supplies and equipment before departure using projectspecific site-visit checklists. Meters, probes, and sampling gear should be packed for transport to the 

field in such a way as to minimize physical shock and vibration during transport. Rafts or float tubes 

should be packed for transport so as to minimize the potential for puncture by any sharp object.

D-1.2 Site Documentation 

BACKGROUND

This section describes SOPs for establishing and documenting sampling sites on small well-mixed 

streams or lake outlets. This procedure applies to new sites for which approximate locations have 

been designated based on program objectives and sampling design. It also applies to previously 

established sites for which current or updated site documentation is needed. 
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OBJECTIVE

The objective of this procedure is to establish and document new sampling sites and to update 

documentation for established sites, providing:

• Site descriptions and notes;

• Travel and access descriptions and notes;

• Site coordinates obtained in the field using a GPS unit;

• Site and access-related photos; and

• Placement or confirmation of numbered site tags (where applicable).

For established sites, existing site documentation will be evaluated for clarity and improved as 

needed based on conditions observed in the field.

MATERIAL NEEDED FOR USE IN FIELD FOR SITE DOCUMENTATION

1. Available site documentation records for previously established sites:

a. Site location maps, topographic maps, and road maps;

b. Site descriptions and access notes;

c. Site tag numbers and tag tree descriptions (where applicable);

d. Site coordinates; and

e. Site photos.

2. Preliminary site documentation for new sites:

a. Site location maps, topographic maps, and road maps indicating approximate site 

locations; and

b. General site descriptions and access notes.

3. General material for site documentation:

a. Regional-scale topographic and road maps;

b. Stream water sampling site documentation forms on waterproof paper;

c. Clipboard or field notebook and pens for use with waterproof paper;

d. GPS unit with replacement batteries;

e. Digital camera with charged battery and charged replacement battery;

f. Site tags, aluminum nails, and hammer (if applicable);

g. Measuring tape;

h. Blaze orange material for flagging tag trees in photos (if applicable);

i. Gate keys (if needed); and 

j. Cell phone with numbers of project staff and management agency offices.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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SEQUENCE OF INITIAL SITE DOCUMENTATION ACTIVITIES

1. Initiate the Stream Sampling Site Documentation and Sample Record Forms.

a. Complete the header information on each form (1-5). See Stream Form instructions in 

Appendix E for details on completing the forms correctly.

2. Select or locate using GPS the specific sampling site (applies to new sites).

a. The approximate or preliminary location of new site locations will be indicated on 

topographic maps. The sample collection team must still determine the exact point on the 

stream to be sampled. 

b. Avoid establishing sites where streams may not be well mixed, such as locations in close 

proximity to inflowing tributaries or braided channels. Also avoid locations that may be 

influenced by runoff from disturbed areas, roads, trails, drainage ditches, or other sources 

of inflow. Select sites that are upstream rather than downstream of potentially altered 

inflow. As a general rule, select sites that are at least 25 m above or below confluence 

points or inflow.

c. The best point to sample will be where the water is flowing fast or falling, where there 

are no eddies, and where the depth is at least 8 inches (20 cm). Ideally the sampling point 

is one that can be reached while kneeling on the stream bank or on stable rocks 

downstream from the sampling point. If possible, avoid standing in the water to reach the 

sample point. 

3. Obtain coordinates at the site using the GPS unit. These coordinates will be part of the site 

identification information entered into the national database (NRM Air) and must be 

documented exactly the same on future visits to the site.

a. The unit position format should be set to Decimal Degrees (hddd.ddddd). The Datum 

should be set to NAD83. Distance and elevation should be set to meters. Wide Area 

Augmentation System (WAAS) should be enabled. 

b. When “Mark Waypoint” is selected, the default GPS site ID (a number) should be 

changed to the actual Site ID.

c. Before saving the coordinates, note the estimated accuracy of the measurement on the 

Site Documentation Form 1. 

d. Save the coordinates in the GPS unit’s memory and record both the coordinates and the 

elevation on the Sampling Site Documentation Form 1. Do not rely solely on the GPS to 

store the coordinates.

e. Confirm that the waypoint has been saved in the GPS unit.

4. Enter the approximate stream depth and width on the Sample Record Form 3.

a. Enter the approximate average values for stream depth and width observed in the 

sampling site area (about 5 m upstream and downstream of the sampling site) on the 

sample site documentation date.

5. Enter site description information on the Site Documentation Form 2.

a. For existing sites, enter information to improve and update existing site description 

information.
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b. Generally describe the site, referring to proximity to landmarks (trails, bridges, 

tributaries, trees, landscape features, or other relatively permanent features). Add any 

additional information, including detailed stream bank and stream structure descriptions,

that will help future sample collectors indentify the site. Also, add any information here 

that might be relevant to water and stream quality, such as cleared land, roads, 

construction, logging, development, or any earth disturbance, etc. observed above the site 

in the watershed or in the stream.

c. As a general convention, the right bank and left bank of a stream are determined based on 

looking downstream. When documenting field observations on the forms, indicate 

whether the observation was made looking downstream or upstream.

6. Enter travel and access directions on the Site Documentation Form 1.

a. For existing sites, improve and update existing travel and access information.

b. Travel and access notes should be sufficient to guide future sample collectors to the site 

without reliance on GPS units. Not all future sample collectors will necessarily have GPS 

units.

c. Access notes should refer to trails, roads, and permanent landmarks, providing distances 

and, where helpful, compass-based directions. Backtrack if necessary to determine 

distances. Linear distances and directions from the established site waypoint can be 

determined using the GPS unit. 

d. In cases where a parking location is not immediately adjacent to the sampling site, use 

the GPS unit to obtain the coordinates for the parking location and record these in the 

travel and access information area of the Site Documentation Form 1.

e. For complicated or long walk-ins, use the GPS unit to record and save a track. But again, 

do not rely on future sample collectors having access to a GPS unit.

f. Sketch the route on the back of the Site Documentation Form for scanning and saving as 

a jpg image if that would be helpful to future sample collectors.

7. Obtain site and access photos.

a. For site documentation, if the camera allows, set the camera's picture size at 3

megapixels. This will create picture files of about 550-650 kilobytes (Kb). Larger, higher 

resolution files are not needed for site documentation work. Switch to higher resolution if 

you are taking pictures for other purposes.

b. Photos should be obtained providing views looking downstream and upstream of the 

sampling site, views of the tag tree (if applicable, with blaze orange material attached), 

and views of other distinguishing features in relation to the site (e.g., trails, roads, notable 

rocks, trees, landforms, bridges, and signs). Photos should also be obtained to show 

important aspects of site access (e.g., parking area and forks in the trail).

c. All photos should be listed on Site Documentation Form 2, including the filename, date, 

and description. Enter this information at the time that the pictures are taken. Do not rely 

on memory for later entry of photo descriptions. The entered description should serve as 

the photo caption for site documentation reporting.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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D-1.3 Stream Sampling 

Water chemistry data are used to characterize acid-base status, trophic condition, and to classify 

streams based on their water chemistry. Samples for analysis of most parameters are collected into 

plastic bottles. Syringe samples or samples collected into glass bottles with septum caps are preferred 

for collection of sample aliquots for laboratory analysis of pH and DIC where practical. Syringes and 

septum caps are used to protect samples from exposure to the atmosphere because the measured 

values for these parameters can change if the stream water sample equilibrates with atmospheric CO2

subsequent to collection.

Stream samples are obtained at a single sampling location below the water surface in the portion of 

the stream cross section that appears visually to represent the greatest amount of flow or,

alternatively, at mid-channel in an area of flowing water. Spatial variability across the channel of a 

single stream is expected to be minimal in relatively small wadeable streams as compared to the 

variability expected among sites, so a composite water chemistry sample is not required. 

At each stream, optional on-site water data and streamside measurements are made using field meters 

and recorded on Sample Record Form 4. Stream water is collected in one or more bottles and two 60 

mL syringes or glass bottles with septum caps that are stored on ice in darkness and shipped or driven 

to the analytical laboratory as quickly as possible after collection. Overnight express mail to the 

laboratory is required for these samples because the syringe or glass bottle samples need to be 

analyzed, and some or all of the bottled sample needs to be stabilized (by filtration and/or 

acidification) within a short period of time (typically 72 hours) after collection. Check with the 

analytical laboratory in advance of sampling regarding applicable holding times for the parameters to 

be measured.

These SOPs describe the process for routine sampling and data collection at water quality monitoring 

sites on streams. Water samples are collected for lab analysis with optional on-site measurements of 

selected water quality parameters (i.e., water temperature, specific conductance, pH, DO, and 

turbidity) using a multi-parameter instrument (sonde). 

This section describes procedures to be followed for data collection at established water quality 

monitoring sites. The sites may be part of a synoptic sampling or fixed long-term sample site program 

for which water quality data and water samples are collected on a scheduled periodic basis.

DOCUMENTATION OF DATA AND SAMPLE COLLECTION 

The Stream Sampling Record Forms 3, 4, and 5 are used to document sample collection and record 

all field data. These forms are used to record the following information:

• The organization, site ID, and site name;

• The date and arrival time for the site visit and specific times of measurements obtained;

• The name, contact information, and affiliation of the individual who is the Collector of Record 

and responsible for protocol adherence during the site visit;

• Suggested revisions or amendments to site documentation and travel directions;

• A listing of site-related photographs taken, including file name, date, and descriptions;

• Qualitative descriptions of weather, stream discharge level and appearance, and other factors that 

might influence water quality during the site visit; 

• Air temperature;

• Results for all water quality data collected on-site, including:

a. Numerical results, units, and measurement time; and

b. Instruments used.
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• Identification of calibration and post-calibration sensor check records,

• Results for all discharge data collected, including:

a. Location of measurement site relative to the sampling and data collection site;

b. Numerical results, units, and specific time of measurement;

c. Methods identification; and

d. Identification of discharge record files.

• A listing of all samples collected, including:

a. Collection time;

b. Types of samples collected and number of replicates; and

c. Method of delivery to analytical lab.

SEQUENCE OF ACTIVITIES FOR DATA COLLECTION 

Collectors are advised to avoid entering or disturbing the stream or stream bank at, or upstream of, 

the collection site before sample collection and completion of water quality data documentation. The 

typical sequence of activities on arrival at the sampling site is as follows:

1. Confirm the site location based on information in the Site Documentation Forms, including 

coordinates, photos, and access notes.

2. Initiate completion of the Sample Record Forms. 

3. Complete Site Information and General Observations sections of the Stream Sampling Record 

Form 3.

4. Enter information needed to improve or correct the site description and travel directions 

provided on the Sample Record Form 3. 

5. Obtain any photographs needed to improve site documentation and enter file names, dates, and 

descriptions.

6. Note any factors (other than weather and discharge conditions) that might affect water quality 

(e.g., bank or upstream disturbance or debris in the water).

If desired, on-site measurement of one or more parameters can be made using a multiparameter water quality sonde (a hand held instrument with a probe containing multiple sensors 

for measuring various physical parameters) that is lowered into the water. Such measurements 

might include temperature, pH, DO, specific conductance, and/or turbidity. The procedures for 

such in situ data collection will vary with the specific field instrument, but in general require the 

following steps:

• Initiate water quality sonde field calibration and calibration checks. 

• Record results on a water quality instrument calibration and post-calibration record form. 

• Confirm that sensor check criteria are met. If criteria are not met, recalibrate, perform 

sensor maintenance, or replace sonde or sensors as needed to meet the criteria. 

Deploy the water quality sonde for the period required to obtain stabilization. Enter the results 

and time of measurement in the On Site Water Quality Data section of the Stream Sampling 

Record Form 4.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

219

7. Collect water samples and complete the Water Sample and Replicates section of the Sample

Record Form 3. Enter any on-site data into that section of the Sample Record Form 4. 

8. Complete the Chain of Custody Form (see Appendix E for the form and instructions).

9. Check to make sure that all of the information recorded on the sample label(s), chain of custody 

form, and stream sampling record forms match. 

10. Obtain discharge measurements or stage height data, if required. Indicate method, time of 

measurement, result, name of the record file, and location of measurement relative to the data 

and sample collection site in the Stage and Discharge Data section of the Sample Record Form 

5. Note that discharge gaging may be conducted at the same time as other site visit activities if 

the discharge measurement site is downstream of the water quality and data collection site. 

Also note that measurements of discharge or stage height are considered optional.

SAMPLE COLLECTION

In the field, make sure that all labels are filled out correctly (see Appendix E for detailed instructions) 

and that the labels on the bottles (and syringes, if used) are securely attached. Carefully avoid 

disturbance of water upstream of the sampling point before sample collection. This means: do not 

walk in the upstream water or on upstream rocks of the sample site. 

Collect a water chemistry sample from the middle of the stream channel at the sampling site, unless 

no water is present at that location. Throughout the collection process, it is important to take 

precautions to avoid contaminating the sample. Wear gloves provided in sample bag. Rinse all 

sample containers three times with stream water before filling them with the sample. Many streams 

have a very low ionic strength and can be contaminated quite easily by perspiration from hands, 

sneezing, smoking, insect repellent, sunscreen, or chemicals used when collecting other types of 

samples (e.g., formalin or ethanol). Make sure that none of the water sample contacts your hands 

before going into the sample bottle or syringe. The chemical analyses conducted using the syringe 

samples can be affected by equilibration with atmospheric carbon dioxide; thus it is essential that no 

outside air contact the syringe samples during or after collection. 

Document the information from the sample bottle/syringe label on Stream Sample Record Form 3. 

Note any problems related to possible contamination in the form comments section.

General stream sample collection procedures for water chemistry are as follows. See additional detail 

under Section 1.4.2.

Collection into Bottle

1. Select sample location in a flowing portion of the channel near the middle of the stream.

2. Put on gloves provided in the sample bag.

3. Always keep the empty sample bottles capped when it enters and leaves the water.

4. Rinse sample bottle and lid three times with stream water, dumping rinse water on the bank or 

downstream of sampling location.

5. Fill the sample bottle(s) completely, holding the bottle in a tilted position approximately at the 

midpoint between the water surface and the streambed, being careful not to disturb any 

sediment before or while collecting the sample.
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6. Cap the bottle underwater. Bottles should be filled so that no air bubbles remain in the bottle 

and capped very tightly while submerged. This may require tipping the bottle up and squeezing 

the sides of the bottle slightly underwater to help force out any bubbles. This procedure is of 

extreme importance if a septum cap is to be used.

7. Put the sample bottle into a clean plastic zipper-lock bag.

8. Place the sample bottle(s) in a cooler (on ice or in stream water) and shut the lid. This may be a 

soft cooler for packing out of the field to the vehicle or a hard cooler in the vehicle. If a cooler 

is not available, place the bottle(s) in an opaque garbage bag and immerse it in the stream.

Collection into Syringe

1. Rinse the syringe three times with water from the sampling location.

2. Slowly fill the syringe with sample, avoiding generation of air bubbles, until it is two-thirds to 

three-fourths full. This will help to ensure that the plunger remains inserted far enough into the 

filled syringe so that it will not be likely become dislodged during transport.

3. Expel any air by tilting the syringe upward and depressing the plunger to force the air out.

4. Repeat procedure using a second syringe.

5. Place the filled syringes into plastic container for transport.

Collection from a Very Shallow Stream

If the stream is too shallow to collect a sample using standard procedures, the following approach can 

be used using a new clean syringe at each site:

1. Rinse the syringe three times with stream water, downstream of the sample site, as usual.

2. Use the syringe to put stream water in the sample bottle and rinse the sample bottle three times.

3. Finally, use the syringe to fill the bottle to the brim with stream water at the sample site. Cap 

the bottle and proceed as usual.

SAMPLE COLLECTION PROCEDURE

The sample should be collected on a step-by-step basis as follows:

1. Remove the gloves from the plastic bag and put them on. 

2. Remove the sample bottle from the plastic bag. Do not put the bag on the ground.

3. Check to ensure that the correct labels are affixed to each sample bottle and syringe.

4. Rinse the sample bottle in the stream at a location at least 2-3 feet downstream of the sample 

collection point. Always keep the empty sample bottle capped when entering and leaving the 

water. The bottle and cap should be rinsed three times. For each rinse, fill the bottle and then 

pour the rinse water over the inside of the cap, held bottom-side up in the other hand. Pour the 

rinse water downstream of the rinsing and sampling points and avoid stirring-up the streambed 

debris during the process.

5. After the rinsing is completed, move upstream to the sampling point and collect the sample by 

submersing the tilted bottle or syringe to a depth midway between the sediment and the water 

surface. If collecting in a bottle, remove the cap and fill up the bottle as completely as possible. 

Cap the bottle while underwater making sure no air bubbles remain. This can be done by tilting 

the bottle up and squeezing the sides slightly before putting on the cap. This procedure is of 

extreme importance if a septum cap is to be used. While collecting the sample, avoid stirring-
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

221

up streambed debris that might be collected with the sample. Try to avoid generating large 

bubbles in the bottle while it is being filled. Make sure the sample does not flow over sampler’s 

gloves or the outside of the bottle during collection. This can often be best achieved by 

sampling rapidly flowing or falling water. If debris may have entered the sample bottle, discard 

the contents downstream, re-rinse the bottle (or use a clean back-up bottle), and collect a new 

sample. 

6. Immediately after collecting the sample, return the bottle to its plastic bag. Seal the bag. 

7. If a sample is to be collected into a syringe, submerge a 60-mL syringe halfway into the stream 

and withdraw a 15-20 mL aliquot. Pull the plunger to its maximum extension and shake the 

syringe so the water contacts all surfaces. Point the syringe downstream and discard the water 

by depressing the plunger. Repeat this rinsing procedure two more times.

8. Submerge the syringe into the stream again and slowly fill the syringe with a fresh sample. Try 

not to get any air bubbles in the syringe. 

9. Invert the syringe (tip pointing up), and cap it with a syringe valve or stopcock. Tap the side of 

the syringe lightly to detach any trapped air bubbles. With the valve open, expel the air bubbles 

and a small volume of water, leaving the syringe between two-thirds and three-fourths full. 

Note that the syringe is transported only partially full to avoid dislodging the plunger during 

transport. Close the syringe valve. If any air bubbles were drawn into the syringe during this 

process, discard the sample and fill the syringe again (step 8).

10. Repeat steps 7 through 9 with a second (back-up) syringe. Place the syringes together in a 

separate plastic bag and place in a plastic container, which is then placed into the cooler (or 

stream water if that method of cooling is used while still in the field).

11. Complete Stream Sample Record Form 3 while at the sample site. 

12. Inspect all equipment, and clean off any plant and animal material before moving to the next 

sample location. This effort ensures that introductions of nuisance species do not occur between 

streams. Inspect, clean, and handpick plant and animal remains from any footwear or 

equipment that may have contacted stream water.

SAMPLE HANDLING

1. Place the bagged sample on double-bagged ice or refrigerant immediately after collection. 

Note: do not put ice in the plastic bag that contains the sample bottle or in the plastic container 

that contains the syringes. Ice or refrigerant should be double bagged in plastic bags to avoid 

possible leakage and contamination of the samples. Samples can be held in a soft-sided cooler 

until returned to the vehicle.

2. The large sample cooler can be left in the collection team’s vehicle. The sample can be 

transferred to the larger cooler upon return to the vehicle.

3. For sites that are not close to road access, the collection team should make arrangements to 

keep the sample on ice after collection and during the return hike. One approach would be to 

use a small soft-pack cooler or other container that will fit in a backpack. Ice, snow, or 

refrigerant could be placed in a plastic bag in the cooler or container (double bag to avoid 

leakage and contamination of samples). Samples are transferred to the larger cooler at the 

vehicle.

4. The samples should be kept in the dark and on ice until delivery to the lab. The ice may need to 

be replenished during sample transit. Do not place the sample bottle in a refrigerator or cooler 

with food or in any container that is not clean. Ship the samples as soon as possible, preferably 
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within 24 hours of sampling. Ship early in the week to ensure the lab receives the samples 

during the work-week.

5. Note that we do not recommend filtration in the field. If, however, a program filters the 

samples in the field for chlorophyll a measurement, it is important to record on the sampling 

record form the volume of water filtered. Record this information in the Notes section of the 

form. The filter is then sent to the analytical laboratory for determination of chlorophyll a

content. 

POST SAMPLING ACTIONS

1. Completely fill out all stream sampling forms. Refer to Appendix E for instructions on filling 

out the forms.

2. Complete the Chain of Custody Form.

3. Check to make sure that all of the information on the sample label(s), Chain of Custody Form, 

and all stream sampling forms is consistent. 

4. Transport the samples back to the vehicle in a soft cooler on ice or snow. 

5. After carrying the samples to the vehicles, place the bottle(s) and syringes in a cooler and 

surround them with 1 gallon re-sealable plastic bags filled with ice. Double bag the ice to avoid 

getting cooling water into sample bags.

FIELD MEASUREMENTS

Determine stream temperature with a field thermometer (one that does not use mercury). Determine 

specific conductance and dissolved oxygen concentration using field meters (optional). Follow 

instructions provided below. Record the measured values on the Stream Sample Record Form. 

Measuring for Specific Conductance

1. Check the batteries and electronic functions (e.g., zero and red line) of the conductivity meter 

as instructed by the operating manual.

2. If you haven’t tested the meter at a base location recently, insert the probe into the RINSE 

container of the quality control check sample (QCCS) and swirl for three to five seconds. 

Remove the probe, shake it off gently, transfer it to the TEST container of QCCS, and let it 

stabilize for 20 seconds. If the measured conductivity is not within 10% of the theoretical 

value, repeat the measurement process. If the value is still unacceptable, do not use the meter 

until it can be inspected, problem(s) diagnosed, and repaired.

3. Submerge the probe in an area of flowing water near the middle of the channel at the same 

location where the water chemistry sample was collected. Record the measured conductivity 

and any pertinent comments about the measurement on the Field Measurement form.

Measuring for Dissolved Oxygen and Temperature

1. Inspect the probe for outward signs of fouling and for an intact membrane. Do not touch the 

electrodes inside the probe with any object. Always keep the probe moist by keeping it inside 

its calibration chamber.

2. Check the batteries and electronic functions of the meter as described in the operating manual.

3. Calibrate the oxygen probe in water-saturated air as described in the operating manual. Allow 

at least 15 minutes for the probe to equilibrate before attempting to calibrate. Try to perform 
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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the calibration as close to stream temperature (not air temperature) as possible by using stream 

water to fill the calibration chamber before equilibration.

4. After the calibration, submerge the probe in midstream at mid-depth at the same location where 

the water chemistry sample was collected. Face the membrane of the probe upstream and allow 

the probe to equilibrate. Record the measured DO and stream temperature on the Field 

Measurement form. Record the time that the DO and temperature measurements were made in 

24 hour units (e.g., 14:23) on the Sample Record Form 4. If the DO meter is not functioning, 

measure the stream temperature with a field thermometer and record the reading on the Sample 

Record Form 4 along with any pertinent comments.

NOTE: Older model dissolved oxygen probes require a continuous movement of water (0.3 to 0.5 

m/s) across the probe to provide accurate measurements. If the velocity of the stream is appreciably 

less than that, agitate the probe in the water as you are taking the measurement.

Measuring for Temperature Only (if no field meters are being used)

1. Place a field thermometer (±1o C accuracy) beneath the surface of the stream at the approximate 

depth of sample collection in an area of flowing water at or near where the water chemistry 

samples were collected.

2. Record the stream temperature (estimated to the nearest 0.1o C) on the Field Measurement 

form. Record the time the temperature measurement was made in 24 hour units (e.g., 14:23) on 

the field form, along with any pertinent comments (e.g., measurement taken in sun or shade).

Steps below describe the equipment cleaning procedures to be followed after measurements are 

taken. Inspect all equipment and clean off any plant and animal material. This effort ensures that 

introductions of nuisance species do not occur between streams. 

Cleaning Equipment After Sampling

1. Clean any equipment that may have contacted surface water for biological contaminants. If you 

are moving between sites on the same day, do this before moving to the next site.

2. Clean and dry other equipment before storage. Rinse coolers with water to clean off any dirt or 

debris on the outside and inside.

3. Inventory equipment and supply needs and relay orders to the Project Coordinator.

4. Remove dissolved oxygen meters, other instrumentation, and GPS units from carrying cases 

and set up for pre-departure checks and calibration. Examine oxygen membranes of DO meters 

for cracks, wrinkles, or bubbles. Replace if necessary.

5. Recharge batteries overnight if possible. Replace other batteries as necessary.

6. Recheck field forms from the day's sampling activities. Make corrections and completions 

where possible and initial each form after review.

POST-TRIP ACTIVITIES

Upon return to a lodging or office location after sampling, the team should review all labels and 

completed data forms for accuracy, completeness, and legibility. A final inspection should be made 

of all samples. If information is missing from the forms or labels, the Project Leader should attempt, 

if possible, to fill in the information accurately. The Project Leader should initial all data forms after 

review. If samples are missing or not properly labeled, it may be necessary to reschedule the site for 

complete sampling. Other post-sampling activities include inspection and cleaning of sampling 

equipment, inventory and sample preparation, sample shipment, and communications.
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EQUIPMENT CLEANUP AND CHECK 

Inspect, clean, and handpick plant and animal remains from any vehicle, footwear, or equipment that 

may have contacted stream water.

SHIPMENT OF SAMPLES AND FORMS

Upon completion of data and sample collection, the samples and forms should be shipped or 

transported to the analytical laboratory in as short a time as is reasonably possible. Call or email the 

lab to alert them that samples are in transit and tell them what date to expect delivery. Samples should 

be maintained in insulated containers with refrigerant after collection and during transport. The Chain 

of Custody Record Form should be maintained and kept with the samples until the samples are 

logged-in at the analytical laboratory. If samples are to be shipped to the laboratory, an overnight 

shipping service should be used, and shipping should be avoided when samples would be delayed by 

transit over a weekend or holiday period. 

The field team should ship samples to the lab as soon as possible after collection. Water samples 

should be shipped in coolers packed with ice. Line each shipping cooler with a large plastic bag. 

Inside, package the ice separately within numerous (as many as feasible) zipper-lock plastic bags and 

ensure that the ice is fresh before shipment. Use block ice when available or “blue ice” packaged 

refrigerant. Block ice should be sealed in two large plastic bags. White or clear bags will allow for 

labeling with a dark indelible marker; label all bags of ice as “ICE” with an indelible marker to 

prevent misidentification by couriers of any leakage of water as a possible hazardous material spill.

To prepare the sample bottles and syringes for shipping, line the shipping cooler with a 30-gal plastic 

bag. Place another plastic bag in the cooler, and place the samples in the second bag. Place filled 

syringes and/or glass bottles in sturdy containers to prevent damage during transport. Ensure that all 

label entries are complete and close the bag of samples. Place bags of ice around it. Then close the 

cooler liner (outer garbage bag). Ship water samples on the day of collection whenever possible. If 

that is not possible, they should be shipped the next day.

PROCESSING SITE DOCUMENTATION DATA AND INFORMATION 

A file system and database with reliable backup should be established for storage of site records and 

files, map images, and photos. Processing site documentation data and information include the 

following steps:

1. Retrieve site coordinates (and any tracks) from the GPS unit using the GPS software. Delete 

any extra coordinate sets (waypoints) and save the file as a *.gdb file.

2. Retrieve photos from the camera. 

3. Enter or revise the site record in the site documentation database.

a. Enter site coordinates obtained in the field.

b. Enter or revise the site description, travel and access directions. 

c. Enter or revise the tag and tag tree information as needed. If no changes were made, note 

that the tag placement was confirmed on the particular date. Note that tree tags may not 

be applied in a wilderness setting. 

d. Add new photos as JPG images with captions to the site record. 

4. Create site maps providing both detailed and broader information for access and orientation. 

Annotate maps and pictures with text and arrows when it would be helpful. Note that the 
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

225

accuracy of maps may vary and the coordinate-based point on the maps, as well as other 

information, may be misleading. Add clarifying notes. Save these maps as JPG images in the 

site record. Add captions as appropriate. Enter site and visit data into NRM Air and attach the 

JPG images to the data records.

D-1.4 Stream Discharge

Stream discharge is equal to the product of the mean current velocity and vertical cross sectional area 

of flowing water. It reflects the volume of water per unit time that passes a particular location (a line 

drawn at a right angle to the stream channel) on the stream. Discharge measurements can be helpful 

for assessing trends in stream water chemistry that are sensitive to stream flow differences. Stream 

discharge information is also useful in interpreting the representativeness of water chemistry data and 

some physical habitat information. 

The location selected for measuring stream discharge should be as close as possible to the location 

where chemical samples are collected. Variability in stream discharge within the reach of interest is 

expected to be small compared to variability in stream discharge among streams, so multiple 

determinations at a site are not required. 

No single method for measuring discharge is applicable to all types of stream channels. The preferred 

procedure for obtaining discharge data for small streams is based on “velocity-area” methods (e.g., 

Rantz 1982, Linsley et al. 1982). For streams that are too small or too shallow to use the equipment 

required for the velocity-area procedure, an alternative method, the timed filling procedure, is 

presented. It is based on timing the filling of a bucket of known volume with water. 

VELOCITY-AREA PROCEDURE

Because velocity and depth typically vary greatly across a stream, accuracy in field measurements is 

achieved by measuring the mean velocity and flow cross-sectional area of many increments across a 

channel (Figure D-1). Each increment gives a subtotal of the stream discharge, and the whole is 

calculated as the sum of these parts. Discharge measurements are made at only one carefully chosen 

channel cross section within the sample reach. It is important to choose a channel cross section that is 

as much like a canal as possible. A glide area with a U-shaped channel cross section that is free of 

obstructions provides the best conditions for measuring discharge by the velocity-area method. You 

may remove rocks and other obstructions to improve the cross-section before any measurements are 

made. However, because removing obstacles from one part of a cross-section affects adjacent water 

velocities, you must not change the cross-section once you commence collecting the set of velocity 

and depth measurements.

Figure D-1. Layout of a channel cross-section for obtaining discharge data by the velocity-area 

procedure.
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The procedure for obtaining depth and velocity measurements is outlined below (Rantz 1982). 

Record the data from each measurement in the Stream Discharge section of the Stream Sampling 

Record Form, giving the distance from the left bank (facing downstream), water depth, measured 

velocity, and any required flags or notes for each measurement increment.

1. Locate a cross-section of the stream channel for discharge determination that has most of the 

following qualities:

• Segment of stream above and below cross-section is straight.

• Depths mostly greater than 15 cm and velocities mostly greater than 0.15 m/s. Do not 

measure discharge in a pool.

• “U” shaped, with a uniform streambed free of large boulders, woody debris or brush, and 

dense aquatic vegetation.

• Flow is relatively uniform, with no eddies, backwaters, or excessive turbulence.

2. Lay the surveyor’s rod (or stretch a meter tape) across the stream perpendicular to its flow with 

the “zero” end of the rod or tape on the left bank, as viewed when looking downstream. Leave 

the tape tightly suspended across the stream, at the bankfull mark or higher. Adjust the tape 

with the aid of a small bubble level suspended from the rod or tape so that it is, and remains 

throughout the period of measurement, level.

Physical indicators of the bankfull stage include: 1) top of highest depositional features, 2) 

break in the slope of the bank or a change in particle size, 3) staining of rocks, and 4) exposed 

root hairs below an intact soil.

3. Attach the velocity meter probe to the calibrated wading rod. Check to ensure the meter is 

functioning properly and the correct calibration value is displayed. Calibrate (or check the 

calibration) the velocity meter and probe as directed in the meter’s operating manual. Place an 

X in the VELOCITY AREA box on the Stream Discharge Form.

4. Divide the total wetted stream width into 15 to 20 equal-sized intervals. To determine interval 

width, divide the width by 20 and round up to a convenient number. Intervals should not be less 

than 10 cm wide, even if this results in less than 15 intervals. The first interval is located at the 

left margin of the stream (left side, looking downstream) and the last interval is located at the 

right margin of the stream (right side, looking downstream).

5. Stand downstream of the rod or tape and to the side of the first interval point (closest to the left 

bank, looking downstream).

6. Place the wading rod in the stream at the interval point and adjust the probe or propeller so that 

it is at the water surface. Place an X in the appropriate DISTANCE UNITS and DEPTH 

UNITS boxes on the Stream Discharge Form. Record the distance from the left bank and the 

depth indicated on the wading rod on the Stream Discharge Form.

Note that, for the first interval, distance equals 0 cm, and in many cases depth may also equal 0 

cm. For the last interval, the distance will equal the wetted width (in cm) and depth may again 

equal 0 cm.

7. Stand downstream of the probe or propeller to avoid disrupting the stream flow. Adjust the 

position of the probe on the wading rod so that it is at 0.6 of the measured stream depth below 

the surface of the water. Face the probe upstream at a right angle to the cross-section, even if 

local flow eddies hit at oblique angles to the cross-section.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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8. Wait 20 seconds to allow the meter to equilibrate, then measure the velocity. Place an “X” in 

the appropriate VELOCITY UNITS box on the Stream Discharge Form. Record the value on 

the Stream Discharge Form. Note for the first interval, velocity may equal 0 because depth will 

equal 0. Note that negative velocity readings are possible; when recording negative values, 

assign a flag to denote they are indeed negative values.

• For the electromagnetic current meter (e.g., Marsh-McBirney), use the lowest time constant 

scale setting on the meter that provides stable readings.

• For the impeller-type meter (e.g., Swoffer 2100), set the control knob at the mid-position of 

DISPLAY AVERAGING. Press RESET then START and proceed with the measurements.

9. Move to the next interval point and repeat steps 6 through 8. Continue until depth and velocity 

measurements have been recorded for all intervals. Note for the last interval (at the right 

margin), depth and velocity values may equal 0.

10. At the last interval (the right margin), record a Z in the FLAG field on the field form to denote 

the last interval sampled.

TIMED FILLING PROCEDURE

In channels too small for the velocity-area method, discharge can sometimes be determined directly 

by measuring the time it takes to fill a container of known volume. “Small” is defined as a channel so 

shallow that the current velocity probe cannot be placed in the water or where the channel is broken 

up and irregular due to rocks and debris and a suitable cross-section for using the velocity area 

procedure is not available. The timed filling method can be extremely precise and accurate but 

requires a natural or constructed spillway of free-falling water. Because obtaining data by this 

procedure can result in channel disturbance and can stir up a lot of sediment, wait until after all 

biological and chemical measurements and sampling activities have been completed before 

proceeding. 

Choose a cross-section of the stream that contains one or more natural spillways or plunges that 

collectively include the entire stream flow. A temporary spillway can be constructed using a portable 

V-notch weir, plastic sheeting, or other materials (i.e., rocks or wood) that are available onsite. 

Choose a location within the sampling reach that is narrow and easy to block when using a portable 

weir. Position the weir or constructed spillway in the channel so that the entire flow of the stream is 

completely rerouted through its notch (Figure D-2). Impound the flow with the weir, making sure that 

water is not flowing beneath or around the side of the weir. Use mud or stones and plastic sheeting to 

get a good waterproof seal. The notch must be high enough to create a small spillway as water flows 

over its sharp crest.

The timed filling procedure is presented in below. Make sure that the entire flow of the spillway is 

going into the bucket. Record the time it takes to fill a measured volume on the Stream Discharge 

section of the Stream Sampling Record Form. Repeat the procedure five times. Discharge will be 

calculated as an average of these five measurements. If the cross-section contains multiple spillways, 

you will need to do separate determinations for each spillway. Clearly indicate which time and 

volume data replicates should be averaged together for each spillway; use an additional Stream 

Sampling Record Form if necessary. On the additional form, record a flag value (e.g., F1) on all lines 

in the Timed Filling section, and explain that the flag means an additional spillway was measured in 

the comment section.
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Figure D-2. Use of a portable weir in conjunction with a calibrated bucket to obtain an estimate of stream 

discharge.

To determine stream discharge through the time-filling procedure:

1. Choose a cross section that contains one or more natural spillways or plunges, construct a 

temporary spillway using on-site materials, and/or install a portable weir using a plastic sheet 

and on-site materials.

2. Place an X in the TIMED FILLING box in the stream discharge section of the Stream 

Discharge form.

3. Position a calibrated bucket or other container of known volume beneath the spillway to 

capture the entire flow. Use a stopwatch to determine the time required to collect a known 

volume of water. Record the volume collected (in liters) and the time required (in seconds) on 

the Stream Discharge form.

4. Repeat step 3 a total of five times for each spillway that occurs in the cross section. If there is 

more than one spillway in a cross-section, you must use the timed-filling approach on all of 

them. Additional spillways may require additional data forms.

Note: If measuring discharge by this procedure will result in significant channel disturbance or will 

stir up sediment, do not determine discharge until all biological and chemical measurement and 

sampling activities have been completed.

EQUIPMENT AND SUPPLIES

Table D-2 shows the list of equipment and supplies necessary to measure stream discharge. Use this 

checklist to ensure that equipment and supplies are organized and available at the stream site in order 

to conduct the activities efficiently.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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Table D-2. Equipment and supply checklist for measuring stream discharge.

Quantity Item 

1 Surveyor’s telescoping leveling rod (7 m long, metric scale, round cross-section)

1 50-m fiberglass measuring tape and reel

1 Small bubble level to make sure that the tape is level

1 Current velocity meter, probe, and operating manual

1-2 Extra batteries for velocity meter

1 Top-set wading rod (metric or English scale) for use with current velocity meter

1 Portable weir with 60° “V” notch (optional)

1 Plastic sheeting to use with weir (optional)

1 Plastic bucket (or similar container) with volume graduations or known total volume

1 Stopwatch

1 Clipboard

Soft (#2) pencils

Stream Discharge forms (one per stream plus extras if needed for timed filling procedure or 

additional velocity-area intervals)

1 copy Field operations and methods documents

1 set Laminated sheets of procedure tables and/or quick reference guides for stream discharge

D-1.5 Acknowledgments

This SOP is based partly on material developed in the following publications: 

Herlihy, A.T. 2006. Section 5. Water chemistry. In: Peck, D.V., A.T. Herlihy, B.H. Hill, R.M. 

Hughes, P.R. Kaufmann, D.J. Klemm, J.M. Lazorchak, F.H. McCormick, S.A. Peterson, P.L. 

Ringold, T. Magee, and M. Cappaert. Environmental Monitoring and Assessment ProgramSurface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams. 

EPA/620/R-06/003. U.S. Environmental Protection Agency, Office of Research and 

Development, Washington, DC. pp. 85–98.

Kaufmann, P.R. 2006. Section 6. Stream Discharge. In: Peck, D.V., A.T. Herlihy, B.H. Hill, R.M. 

Hughes, P.R. Kaufmann, D.J. Klemm, J.M. Lazorchak, F.H. McCormick, S.A. Peterson, P.L. 

Ringold, T. Magee, and M. Cappaert. Environmental Monitoring and Assessment ProgramSurface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams. 

EPA/620/R-06/003. U.S. Environmental Protection Agency, Office of Research and 

Development, Washington, D.C. pp. 99–110.

Merritt, G.D.,V.C. Rogers, and D.V. Peck. 1997. Section 3. Base site activities. In: Baker, J.R., D.V. 

Peck, and D.W. Sutton (eds.). Environmental Monitoring and Assessment Program Surface 

Waters: Field Operations Manual for Lakes. Report No. EPA/620/R-97/001. U.S. 

Environmental Protection Agency, Washington DC.
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D-2 STANDARD OPERATING PROCEDURE FOR LAKE 

SAMPLING

This SOP provides guidelines for lake sampling within the FS ARM program. It is intended as a 

general SOP, suitable for adoption as a stand-alone procedure, or for modification to fit local program 

needs. It is divided into individual sections that cover pre-trip activities, sampling site documentation, 

index site location, lake sampling, lake assessment, post-trip activities, and needed equipment and 

supplies.

D-2.1 Pre-Trip Activities 

Field teams conduct a number of activities before departing for the field sampling. These include 

tasks that must be completed both before departure to the sampling site. This section describes 

procedures for office and base site pre-trip activities that should be carried out in support of lake 

sampling. 

Pre-departure activities include development of sampling itineraries, instrument calibration, 

equipment checks and repair, supply inventories, and sample container preparation. Procedures for 

these activities are described in the following sections. 

PREPARATION OF SAMPLE CONTAINERS

Generally, the analytical laboratory supplies properly washed bottles and syringes used in the 

sampling program. In order to do this, the laboratory needs to know in advance what analytes will be 

measured in the samples to be collected. Ensure that the proper number, type, and size sampling 

containers are provided. It is wise to carry a few back-ups.

Before leaving the base location, package the sample containers (typically two sample bottles and 

two 60 mL syringes or glass bottles with septum caps) for each site to be sampled (plus back-up). Fill 

out a set of water chemistry sample labels. Attach a completed label to each sample bottle and 

syringe. Make sure the syringe labels do not cover the volume gradations on the syringe.

DAILY ITINERARIES 

Field sampling efforts should include a Project Leader who guides activities in the field and a Project 

Coordinator who remains in the office during the sampling effort. The Project Leader reviews each 

site folder to ensure that it contains the appropriate maps, contact information, copies of access 

permission letters (if needed), and access instructions. Additional activities can include confirming 

the best access routes, calling landowners or local contacts (if applicable), confirming lodging or 

camping plans and locations (with directions), and coordinating rendezvous locations with 

individuals who must meet with field teams before accessing a site. This information is used to 

develop an itinerary. The Project Leader should provide the Project Coordinator with a schedule for 

each day of sampling. Schedules include departure time, estimated duration of sampling activities, 

routes of travel, and estimated time of arrival at the sampling site(s) and return to the base site. 

Changes that might be made to the itinerary should be relayed by the Project Leader to the Project 

Coordinator as soon as possible. Miscommunications can result in the initiation of expensive search 

and rescue procedures and disruption of carefully planned schedules.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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INSTRUMENT CHECKS AND CALIBRATION 

Each field team should test and/or calibrate field instruments such as, thermometers, DO meters, and

GPS units, before leaving for the sampling site. Batteries should be checked before departure for field 

sites. Extra batteries should be carried. 

Field personnel should check the inventory of supplies and equipment before departure using sitevisit checklists. Meters, probes, and sampling gear should be packed for transport to the field in such 

a way as to minimize physical shock and vibration during transport. Rafts or float tubes should be 

packed for transport so as to minimize the potential for puncture by any sharp object. 

SUPPLY INVENTORIES

Develop a checklist of equipment and supplies that will be needed to conduct lake sampling. Check 

off each item as it is packed and loaded for transport to the field. 

A preliminary list of equipment and supplies required to collect lake samples and associated field 

data is presented in Table D-3. Use and revise this checklist to ensure that equipment and supplies are 

organized and available at the lake sampling site in order to conduct the activities efficiently.
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Table D-3. Checklist of equipment and supplies for sampling water chemistry and Secchi depth.

Quantity Item 

Standard Items:

1 Field thermometer

1-2 Sample bottle(s) with completed sample label attached (in clean plastic bag). Include a second bottle 

if sampling at that site is to be replicated

2-4 60 mL plastic syringes (with Luer type tip) or glass bottles with septum caps with completed sample 

labels attached

1 Plastic container with snap-on lid to hold filled syringes

2-4 Syringe valves (Mininert® with Luer type adapter, or equivalent, available from a chromatography 

supply company)

1 Cooler with 4 to 6 plastic bags (1-gal) of ice or a medium or large opaque garbage bag to store the 

water sample at shoreline

1 Lake Sampling Record Form

1 set Water Chemistry labels (if not filled out and attached at base site)

2-4 Soft-lead pencils and write-in-rain-pens for filling out field data forms and notebook entries

2-4 Fine-tipped indelible waterproof markers for filling out labels

1 copy Field operations and methods documents

2-4 Plastic gloves stored in a secure plastic bag

1 Survey grade GPS unit and compass

1 Digital camera with extra memory cards and batteries

1 Backpack with waterproof cover (if site is not accessible by vehicle)

1 Van Dorn sampler with messenger and cable

1 Raft or float tube with pump for inflating, oars, paddles, or flippers

1-2 Personal flotation devices for each person in the boat or float tube

1 First aid kit

1 Locally determined safety equipment

1 Secchi disk and line (with depth increments)

1 Tape measure

Optional Items:

roll/box Clear packaging tape to cover labels (tape strips)

1 Dissolved oxygen (DO)/temperature meter with probe

1 DO repair kit containing additional membranes and probe filling solution

1 Conductivity meter with probe

1 250-mL or 500-mL plastic bottle of conductivity QCCS labeled RINSE (in plastic bag)

1 250 mL or 500-mL plastic bottle of conductivity QCCS labeled TEST (in plastic bag)
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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D-2.2 Site Documentation 

BACKGROUND

This section describes SOPs for establishing and documenting sampling sites on small to mediumsize lakes, primarily those situated in relatively remote backcountry locations. This procedure applies 

to new sites for which approximate locations have been designated based on program objectives and 

sampling design. It also applies to previously established sites for which current or updated site 

documentation is needed. 

Sampling the correct lake is critical to most lake study sampling designs. It is also important to 

identify, to the extent possible, the index site (deepest point) on a lake. On arriving at a target lake, 

the GPS unit is a valuable tool for identifying and verifying the location. Nevertheless, site 

verification must be supported by all available information (e.g., maps, photos, signs, GPS, and 

expected lake size and shape). Do not sample the lake if there is reason to believe it is the wrong one. 

Contact the Project Coordinator to resolve discrepancies.

OBJECTIVE

The objective of this procedure is to establish and document new lake sampling sites and to update 

documentation for established sites, providing:

• Site descriptions and notes;

• Travel and access descriptions and notes;

• Site coordinates obtained in the field using a GPS unit; and

• Site and access-related photos.

For established sites, existing site documentation will be evaluated for clarity and improved as 

needed based on conditions observed in the field.

MATERIAL NEEDED FOR USE IN THE FIELD FOR SITE DOCUMENTATION

1. Available site documentation records for previously established sites:

a. Site location maps, topographic maps, and road maps;

b. Site descriptions and access notes;

c. Site coordinates; and

d. Site photos.

2. Preliminary site documentation for new sites:

a. Site location maps, topographic maps, and national forest maps, indicating approximate site 

locations; and

b. General site descriptions and access notes.

3. General material for site documentation:

a. Regional-scale topographic and road maps;

b. Lake sampling site documentation forms on waterproof paper;

c. Clipboard or field notebook and pens for use with waterproof paper;

d. GPS unit and replacement batteries;

e. Digital camera with charged battery and charged replacement battery;

f. Gate keys (if needed); and

g. Cell phone with numbers of project staff and management agency offices.
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SEQUENCE OF SITE DOCUMENTATION ACTIVITIES

1. Complete the header information on each form (1-4) before entering the field. See Lake Form 

instructions in Appendix E for details on completing the forms correctly.

2. Select or locate the specific sampling site using the GPS unit (applies to new sites).

a. The approximate or preliminary location of new site locations will be indicated on 

topographic maps. The sample collection team must still determine the exact point on the 

lake to be sampled. 

b. The preferred sampling site location is over the deepest area of the lake, which is often, 

but not always, near mid-lake. This is designated as the “index” site. If it is not feasible to 

access the index site, or for conducting some types of screening studies, it is acceptable 

to sample from the principal outlet stream (designated “outlet” sample) rather than at the 

index site. If there is no available outlet stream or if the outlet stream is not flowing at a 

sufficient rate to collect a representative sample, then it can be acceptable to collect a 

sample by reaching into the lake from an appropriate location along the lake shore 

(designated “shoreline” sample). 

3. Obtain coordinates at the site using the GPS unit. These coordinates will be part of the site 

identification information entered into the national database (NRM Air).

a. The unit position format should be set to Decimal Degrees (hddd.ddddd). The Datum 

should be set to NAD83. Resolution should be expressed in meters. Wide Area 

Augmentation System (WAAS) should be enabled. 

b. When “Mark Waypoint” is selected, the default GPS site ID (a number) should be 

changed to the actual Site ID.

c. Before saving the coordinates note the estimated accuracy of the measurement and enter 

on the Site Documentation Form 1. 

d. Save the coordinates in the GPS unit memory and record the coordinates in decimal 

degrees, the datum, and the elevation (preferably in meters) on the Sampling Site 

Documentation Form 1. Use NAD83 to conform to Forest Service standards. Do not rely 

solely on the GPS unit to store the coordinates.

e. Confirm that the waypoint has been saved in the GPS unit.

f. It can be very helpful to establish and document benchmarks on shoreline rocks. 

4. Enter (if applicable) the approximate lake water level on the Sampling Site Documentation 

Form 2.

5. Enter site description information on the Site Documentation Form 2.

a. For existing sites, enter information to improve and update existing site description 

information.

b. Generally describe the site, referring to the proximity of landmarks (trails, bridges, 

tributaries, trees, shoreline features, landscape features, or other relatively permanent 

features). Add any additional information that will help future sample collectors indentify 

the site. Also, add any information here that might be relevant to water quality, such as 

cleared land, mining (ongoing or historical), roads, construction, logging, heavy grazing, 
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

235

development, or any earth disturbance observed on or near the shoreline, above the lake 

in the watershed, or along the inlet stream(s).

6. Enter travel and access directions on the Sampling Site Documentation Form 1. 

a. For existing sites, improve and update existing travel and access information.

b. Travel and access notes should be sufficient to guide future sample collectors to the site 

without reliance on GPS units. Not all future sample collectors will necessarily have GPS 

units.

c. Access notes should refer to trails, roads, and permanent landmarks, providing distances 

and, where helpful, compass-based directions. Backtrack if necessary to determine 

distances. Linear distances and directions from the established site waypoint can be 

determined using the GPS unit. 

d. In cases where a parking location is not immediately adjacent the sampling site, use the 

GPS unit to obtain the coordinates for the parking location and record them in the travel 

and access information entry area of the Sampling Site Documentation Form.

e. For complicated or long walk-ins, use the GPS unit to record and save a track. But, again, 

do not rely on future sample collectors’ access to a GPS unit.

f. Sketch the route on the back of the Sampling Site Documentation Form for scanning and 

saving as a JPG image if that would be helpful.

7. Make a sketch of the lake on Sampling Site Documentation Form 2. Mark on the sketch the 

launch and sampling site locations. 

8. Obtain site and access photos

a. Photos should be obtained providing views of the sampling site and the shoreline and 

views of other distinguishing features in relation to the site (bridges, roads, notable rocks, 

trees, landforms, signage, etc.) Photos should also be obtained to show important aspects 

of site access (parking area, forks in the trail, etc.).

b. All photos should be listed on Form 2, including their filenames, date, and descriptions. 

Enter this information at the time that the pictures are taken. Do not rely on memory for 

later entry of photo descriptions. The entered description should serve as the photo 

caption for site documentation reporting.

LAKE VERIFICATION AT THE LAUNCH SITE

Record directions to the sampling site and a description of the launch location for lake sampling on 

the Lake Sampling Site Documentation Form in the site information folder. This information will be 

important in the future if the site is revisited by another sampling team. Provide information about 

signs, road numbers, gates, landmarks, and any additional information you feel will be useful to 

another sampling team in locating this site. It is also helpful to describe the road distance traveled (in 

miles) between turns and hiking distance and/or time traveled to reach the sampling or launch site. 

Additional details can also be helpful. What landmarks are in the vicinity of the site? Is the trailhead 

well marked?

If a GPS fix is obtained, record the location in decimal degrees and the type of satellite fix (2D or 3D)

for the site. Compare the site information folder map coordinates recorded for the site with the GPS 

coordinates displayed at the site. Check to see if the two sets of coordinates are within a distance that 
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is approximately equal to the precision of the GPS receiver without differential correction of the 

position fix. If a GPS fix is not available, do not record fix information but try to obtain the 

information at a later time during the visit. A fix may be taken at any time during a site visit and 

recorded on the form. If this is the first visit to this lake, mark the location of the launch site with an 

“L” on the lake outline that is provided on the Lake Sampling Site Documentation Form. In addition 

to the GPS unit, use as many of the following methods as possible to verify the location and identity 

of the site:

1. Obtain confirmation from a local person familiar with the area.

2. Identify confirming trails, roads, and signs.

3. Compare lake shape to that shown on the topographic map included in the site information 

folder.

4. Determine lake position relative to identifiable topographic features shown on the map.

5. Compare visual evaluation of lake area with available mapped information. 

If this is not the first visit to this lake and if the lake shape on the map sketch that appears on the Lake 

Sampling Site Documentation Form and on the USGS map do not correspond with each other or with 

the actual lake shape as seen in the field, check “Not Verified” and provide comments on the form. 

The lake should not be sampled if there are clear, major differences in lake shape or lake area. 

INDEX SITE LOCATION

Locate the sampling site in what is approximately the deepest portion of the lake. There are different 

ways to do this, as follows:

1. If the deepest location had been determined and documented on a previous trip to this lake, 

based on that documentation and use of GPS unit and/or mapped lake features, navigate to the 

sampling location.

2. If the sampling location has not previously been documented, locate the deepest part of the lake 

based on visual examination of the lake shape and surrounding topography, coupled with 

reconnaissance on foot and/or by boat for up to about one-half hour. Use visual cues and/or 

soundings with a weighted line to locate what appears to be the deepest part of the lake. 

Once the sampling location has been selected at what appears to be the deepest part of the lake, 

determine the GPS coordinates and record them on the Lake Sampling Site Documentation Form. 

Mark the sample site with an “x” in the lake drawing. A checklist for lake verification is given in 

Table D-4.

Table D-4. Lake verification checklist.

Item 

Site information folder for lake to be sampled

Clipboard

Lake Sampling Site Documentation Form

Field notebook

GPS unit with manual, extra battery pack

50 m line to attach to rock anchor
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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D-2.3 Lake Sampling 

These procedures cover collection of lake water samples and measurement of Secchi depth 

(transparency). The lake sampling procedures assume collection of the primary sample from the 

deepest part of the lake. Measurement of Secchi depth and collection of the deep water index sample 

will require use of a boat or float tube. If it is not possible to sample the lake by boat or float tube, the 

next best option is to sample at the principal outlet stream. If a lake outlet sample is to be collected 

instead of a sample in deep water, follow the procedures outlined in the stream sampling SOP and 

sample the outlet stream as close to the lake as is practical. 

If neither a deep water sample nor an outlet sample can be collected, the third option is to sample 

from the shoreline, satisfying as many of the following criteria as possible:

• As close to the outlet as possible;

• From a bedrock outcropping or otherwise rocky area; and

• From the deepest accessible point.

Water must be deep enough so that surface scum and sediments are not collected into the bottle. Take 

samples from a wind-exposed area so that the water is relatively well-mixed. Avoid sampling in 

locations having emergent vegetation and/or downed logs or other woody debris.

DOCUMENTATION OF DATA AND SAMPLE COLLECTION

The Lake Sample Record Forms 3 and 4 are used to document sample collection and field data. See 

Appendix E for complete instructions for filling out the forms correctly. These forms are used to 

record the following information:

• The site ID and site name;

• The date and arrival time for the site visit and specific times of measurements obtained;

• The name, contact information, and affiliation of the individual who is the Collector of Record 

and responsible for protocol adherence during the site visit;

• Suggested revisions or amendments to site documentation and travel directions;

• A listing of site-related photographs taken, including file name, date, and descriptions;

• Qualitative descriptions of weather, lake level and appearance, and other factors that might 

influence water quality during the site visit;

• Air temperature;

• Results for all water quality data collected on-site, including:

a. Numerical results, units, and measurement time; and

b. Instruments used and methods identification;

• Identification of calibration and post-calibration sensor check records;

• A listing of all samples collected, including:

a. Collection time; and

b. Types of samples collected and number of replicates.

SEQUENCE OF ACTIVITIES FOR DATA COLLECTION 

The typical sequence of activities on arrival at the lake sampling site is as follows:
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1. Confirm the site location based on information in the Lake Sampling Site Documentation 

Form, including coordinates, photos, and access notes.

2. Initiate completion of the Lake Sample Record Forms (see Appendix E for complete form 

instructions). 

3. Complete Site Information and General Observations sections of the Lake Sample Record 

Forms.

4. Enter information needed to improve or correct the site description and travel directions 

provided on the Site Documentation Form 3. 

5. Obtain any photographs needed to improve site documentation and enter file names, dates, and 

descriptions.

6. Note any factors (other than weather and lake level) that might affect water quality (e.g., 

shoreline or watershed disturbance or debris in water).

7. Collect water samples and complete the Water Sample section of the Sample Record Form 3. 

Enter any on site water data (optional) into Sample Record Form 4. 

8. Complete the Chain of Custody Form. 

9. Check to make sure that all of the information recorded on the sample label(s), Chain of 

Custody Form, and the lake sampling forms is consistent. 

10. Obtain lake level measurements, if required.

SAMPLE COLLECTION

Deep Water Index Sample

Collect a water sample at the index site using a Van Dorn water sampler from 1.5 m depth (0.5 m if 

lake depth is less than 2.0 m), using the procedure described below. From the Van Dorn sampler, fill 

the required number of syringes and/or glass bottles, and one or two 500 ml or 1000 ml sample 

bottles. Procedures for collecting these samples are presented below. Prior to filling syringes and 

sample bottles, check the labels on these containers to ensure that all written information is legible 

and that each container has the same (and correct) site identification number. It can also be a good 

idea to place clear packing tape over the label and identification code, covering the label completely

to seal it against water damage. Record the identification code assigned to the sample set (the 

syringes and bottles collected from the same site are considered one sample) and have the same code 

on the Lake Sampling Record Form. Also record the depth from which the sample was collected 

(usually 1.5 m or 0.5 m) on the form. Enter a flag code and provide comments on the Sample 

If desired, in situ measurement of one or more parameters can be made using a multiparameter 

water quality sonde. Such measurements might include temperature, pH, DO, specific 

conductance, and/or turbidity. The procedures for such in situ data collection will vary with the 

specific field instrument, but in general require the following steps:

• Initiate water quality sonde field calibration and calibration checks. 

• Record results on a water quality instrument calibration and post-calibration record form. 

• Confirm that sensor check criteria are met. If criteria are not met, recalibrate, perform sensor 

maintenance, or replace sonde or sensors as needed to meet the criteria. 

Deploy the water quality sonde for the period required to obtain stabilization. Enter the results and 

time of measurement in the in situ Water Quality Data section of the Lake Sampling Record Form.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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Collection Form if there are any problems in collecting the sample or if conditions occur that may 

affect sample integrity. Store samples in the appropriate containers in the dark, and verify that they 

are carefully packed with plenty of ice bags and properly positioned, sealed, and labeled in the 

sample coolers. Recheck all forms and labels for completeness.

To use the Van Dorn sampler:

1. Open the Van Dorn sampler by pulling the elastic bands and cups back and securing the 

latches. Make sure that the mechanism is cocked so that it will be tripped by the messenger 

weight. Make sure that all valves are closed. Inspect the line for fraying, especially where it 

connects to the Van Dorn sampler. Do not place hands inside or on the lip of the container; this 

could contaminate samples. To reduce chances of contamination, wear powder-free latex 

laboratory gloves.

2. Attach the free end of the messenger line to the boat. This is important to prevent accidental 

loss of the equipment overboard. Rinse the open sampler by immersing it in the water column 

three times.

3. Lower the sampler to 1.5 m below the surface (0.5 m in lakes < 2 m deep).

4. Trip the sampler by releasing the messenger weight so that it slides down the line.

5. Raise the full sampler out of the lake. Set it on a clean, flat surface in an upright position. To 

avoid contamination, do not set the sampler in the bottom of the boat. Applying some body 

weight to the top of the Van Dorn sampler often will seal minor air leaks and preserve the 

sample integrity. If air enters the Van Dorn sampler, discard the sample and obtain another 

(repeat steps 1 to 5).

Use the following procedure for syringe and sample bottle collection. In doing so, wear powder-free 

surgical gloves while collecting syringe and bottle samples. Syringes may be chilled before use to 

reduce the occurrence of air bubbles in the sample. 

Fill one bottle for each routine lake water sample. Fill a second bottle, with its own unique Sample 

ID/Barcode, if this site is to be replicated for QA/QC purposes.

1. Make sure that the sample bottle(s) and 60 ml syringes have the same site identification code 

number (which identifies a single lake) and that the labels are completely covered with clear 

tape. Record the identification code number on the Sample Collection Form.

2. Unscrew the valve at the top of the Van Dorn sampler. Fit a pre-labeled syringe to the fitting.

3. Slowly draw a 20 mL aliquot into the 60 pre-labeled syringe. Pull the plunger back so that the 

water contacts all inner surfaces of the syringe. Expel the water from the syringe. Repeat this 

rinse procedure twice more (there are three rinses for each syringe sample).

4. Reattach the syringe to the Van Dorn sampler and slowly draw 60-mL of water into the 

syringe. If air enters the Van Dorn sampler during this process, dispose of the sample and 

obtain another Van Dorn sample.

5. Place the syringe valve on the syringe tip. Press the green button toward the syringe.

6. Hold the syringe with the tip and valve pointed skyward. Tap the syringe to gather air bubbles 

to the top. Expel all air from the syringe by depressing the plunger and then close the one-way 

stopcock to seal the syringe with 40 to 50 mL of sample water remaining. Place a piece of tape 

around the stopcock so that it cannot freely open and package securely to prevent damage 

during transport (e.g. inside a paper towel cardboard roll). Secure in a container with packing 

material and ice.
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7. Repeat steps 2 to 5 for one to three additional syringes. There should be a total of two syringes 

for each routine water sample if the protocol specifies that water for both DIC and pH be 

collected in syringes (four syringes if sample is being replicated).

8. Place the syringes in the solid plastic container and place in the cooler. Use ice contained in 

sealed 1-gal plastic bags to maintain the sample below 4 μC.

9. Unscrew the top valve of the Van Dorn sampler. Unscrew the lid of the pre-labeled sample 

bottle.

10. Open the bottom valve of the Van Dorn sampler and partially fill the sample bottle with water 

(approximately 50 mL).

11. Screw the lid on the bottle. Shake the bottle so that the water inside contacts all sides. Discard 

the water. Repeat this rinse procedure twice more. Collection of the water sample in the bottle 

should be preceded by three rinses.

12. Open the Van Dorn valve and completely fill the bottle.

13. Compress the plastic bottle to remove any residual head space. Seal the cap tightly. Holding the 

glass bottle (if applicable) level; fill it completely to the top. Seal the cap tightly.

14. Place bottle in a cooler with sealed 1-gal plastic bags of ice. Note the depth from which the 

sample was collected on the Sample Collection Form.

Shoreline Sample

Only collect a shoreline sample if the study objective is to perform a rough screening to identify 

probable lake chemical conditions or if it is not feasible to collect either a deep water or outlet sample 

from the subject lake. Collect the shoreline sample as follows. In the field, make sure that the labels 

all have the sample ID number (barcode), and that the labels on the bottles and syringes are securely 

attached. Carefully avoid disturbance of water or sediment in the vicinity of the sampling point before

sample collection. This means not walking in the water or on loose rocks. If you must walk out to 

obtain a clean sample, wait for the sediment to settle before collecting the sample. 

Collect a water chemistry sample, as described below, in the deepest water possible. Throughout the 

collection process, it is important to take precautions to avoid contaminating the sample. Rinse all 

sample containers three times with lake water before filling them with the sample. Many lakes have a 

very low ionic strength and can be contaminated quite easily by perspiration from hands, sneezing, 

smoking, insect repellent, sunscreen, or chemicals used when collecting other types of samples (e.g., 

formalin or ethanol). Make sure that none of the water sample contacts your hands before going into 

the sample bottle or syringe. The chemical analyses conducted using the syringe or septum bottle 

samples can be affected by equilibration with atmospheric carbon dioxide; thus, it is essential that no 

outside air contact the syringe samples during or after collection. 

Shoreline Collection into Bottle

1. Rinse the sample bottle and lid three times with water. Discard the rinse on the shore away 

from the sample location.

2. Check to ensure that the correct labels are affixed to each sample bottle and syringe. Fill the 

sample bottle(s), holding the bottle in a tilted position approximately at the midpoint between 

the water surface and the lakebed, being careful not to disturb any sediment before or while 

collecting the sample. Try to avoid generating large bubbles in the bottle while it is being filled. 

If a septum cap is being used, place the cap on the bottle under the surface of the water to avoid 

any contact of the sample with the air.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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3. Place the sample bottle(s) in a cooler (on ice or in a bag in the lake) and shut the lid. If a cooler 

is not available, place the bottle(s) in an opaque garbage bag and immerse it in the stream.

Shoreline Collection into Syringe

1. Submerge a 60-mL syringe halfway into the lake and withdraw a 15-20 mL aliquot. Pull the 

plunger to its maximum extension and shake the syringe so the water contacts all surfaces. 

Point the syringe away from the lake and discard the water by depressing the plunger. Repeat 

this rinsing procedure two more times.

2. Submerge the syringe into the lake again and slowly fill the syringe with a fresh sample. Try 

not to get any air bubbles in the syringe. If more than one to two tiny bubbles are present, 

discard the sample and draw another one.

3. Invert the syringe (tip pointing up), and cap it with a syringe valve. Tap the syringe lightly to 

detach any trapped air bubbles. With the valve open, depress the plunger to expel the air 

bubbles and a small volume of water, leaving between 40 and 50 mL of sample in the syringe. 

Close the syringe valve. If any air bubbles are drawn into the syringe during this process, 

discard the sample and fill the syringe again.

4. Repeat steps 1 through 3 with a second syringe. Fill a total of four syringes if the lake sample is 

to be replicated. Place the syringes together in the cooler or temporarily (until time to depart 

from the lake) in the opaque plastic bag immersed in lake water with the sample bottle(s).

Post Sampling Actions

1. Completely fill out all lake sampling forms. Refer to Appendix E for instructions on filling out 

the forms.

2. Complete the Chain of Custody Form.

3. Check to make sure that all of the information recorded on the sample label(s), Chain of 

Custody Form, and Lake Sampling Record Form match.

4. Place each filled bottle into a zipper-lock bag; place the filled syringes into a plastic box with 

snap-on lid. After carrying the samples to the vehicles, place the (bagged) bottle(s) and (boxed) 

syringes in a cooler and surround them with 1-gallon re-sealable plastic bags filled with ice. 

Double bag the ice to avoid getting cooling water (melted ice) into the sample bags.

The sample should be collected on a step-by-step basis as follows:

1. Remove the gloves from the plastic bag and put them on. 

2. Remove the sample bottle from the plastic bag. Do not set the bag on the ground or any dirty 

surface.

3. Rinse the sample bottle in the lake at a location at least 10 feet away from the sample collection 

point. The bottle and cap should be rinsed three times. For each rinse, fill up the bottle and then 

pour the rinse water over the inside of the cap, held bottom-side up in the other hand. Pour the 

rinse water away from the lake sampling point and avoid stirring-up lakebed debris during the 

process.

4. After the rinsing is completed, move to the sampling point and collect the sample by 

submersing the tilted bottle or syringe to a depth midway between the sediment and the water 

surface. Fill the bottle completely. While collecting the sample, avoid stirring-up lakebed 

debris that might be collected with the sample. If debris may have entered the sample bottle, 

discard the contents (at a different location), re-rinse the bottle (or use a clean back-up bottle), 

and collect a new sample. 
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5. Immediately after collecting the sample, place the lid on the bottle (tightly) and return the bottle 

to its plastic bag. Seal the bag. 

6. Complete the Water Sample Collection section of the Lake Sampling Record Form while at the 

sample site.

Record the information from the sample label on the Lake Sampling Record Form. Note any 

problems related to possible contamination in the comments section of the form.

1. Place the sample on ice or refrigerant immediately after collection. Note: do not put ice in the 

plastic bag that contains the sample bottle.

2. For sites that are close to road access, the large sample cooler can be left in the collection 

team’s vehicle. The samples can be placed in the cooler upon return to the vehicle.

3. For sites that are not close to road access, the collection team should make arrangements to 

keep the samples on ice after collection and during the return hike. One approach would be to 

use a small soft-pack cooler or other container that will fit in a backpack. Ice, snow, or 

refrigerant should be placed in a small plastic bag in the cooler or container (double bagged to 

avoid leakage and contamination of samples). 

4. The samples should be kept in the dark and on ice until delivery to the lab. The ice may need to 

be replenished during sample transit. Avoid letting the sample bottle float in melted ice water. 

Do not place the sample bottle in a refrigerator or cooler with food or in any container that is 

not clean. Ship the samples to the laboratory as soon as possible, preferably within 24 hours of 

sampling.

FIELD MEASUREMENTS

Anchor the boat if possible. After achieving a stable position and determining the site depth, measure 

Secchi disk transparency using the procedures below. Record the depth of disk disappearance and the 

depth of reappearance on the Lake Sampling Record Form. If the Secchi disk is visible at the bottom 

of the lake, check the "clear to bottom" space on the form. Comment on the form if there are any 

conditions that may affect this measurement (e.g., surface scum, suspended sediments, or weather 

conditions).

Other field measurements might be made depending on the study. These could include measurements 

at the sample site and/or measurements at other locations or depths. They might include dissolved 

oxygen, specific conductance, and/or other parameters.

To measure for transparency using the disk:

1. Remove sunglasses unless they are prescription lenses.

2. Clip the calibrated chain (should already be in 0.5-m increments) to the Secchi disk. Make sure 

the chain is attached so that depth is determined from the upper surface of the disk.

3. Lower the Secchi disk over the shaded side of the boat until it disappears.If the disk is visible 

to the lake bottom, check the appropriate space on the form.

4. Read the depth indicated on the chain. If the disappearance depth is <1.0 m, determine the 

depth to the nearest 0.01 m by marking the chain at the nearest depth marker and measuring the 

remaining length with a tape measure. Otherwise, estimate the disappearance depth to the 

nearest 0.1 m. Record the disappearance depth on the Lake Sampling Record Form.

5. Slowly raise the disk until it reappears and record the reappearance depth on the form.

6. Note any conditions that might affect the accuracy of the measurement in the comments field.
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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D-2.4 General Lake Assessment

Standard operating procedures are summarized here for the site assessment conducted at lake 

sampling locations. The purpose of this assessment is to record site characteristics that may aid in the 

interpretation of the chemical and/or biological data collected from the lake. 

Team members should complete the Lake Assessment portions of the Site Documentation Forms at 

the end of lake sampling, recording all observations from the lake that were noted during the course 

of the visit. This Lake Assessment is designed as a template for recording pertinent field 

observations. It is not intended to be comprehensive, and any additional observations should be 

recorded in the comments section. The Assessment consists of three major sections: General Lake 

Hydrologic Information, Shoreline Characteristics, and Qualitative Macrophyte Survey. Each is 

described below.

GENERAL LAKE HYDROLOGIC INFORMATION

Observations regarding the general characteristics of the lake are described in Table D-5. The 

hydrologic lake type is an important variable for defining subpopulations for acidic deposition 

effects.

Table D-5. General lake information noted during lake assessment. (Source: Herlihy 1997.)

Item Description

Hydrologic Lake Type Note if there are any stream outlets from the lake, even if they are not flowing. If no lake outlets were 

observed, record the lake as a seepage lake. If the lake was created by a man-made dam (not that a dam is 

present just to raise the water level), record the lake as a reservoir. Otherwise record the lake as a drainage 

lake.

Outlet Dams Note the presence of any dams (or other flow control structures) on the lake outlet(s). Differentiate between 

artificial (manmade) structures and natural structures (beaver dams). Describe in detail the observed flow 

control structure, providing measurements if possible. Note the material from which the structure is made.

Lake Level If a lake level reference point is established, examine the lake shoreline for evidence of lake level changes 

(e.g., “bathtub ring”). If there are none, check "zero"; otherwise try to estimate the extent of vertical changes in 

lake level from the present conditions based on other shoreline signs.

SHORELINE CHARACTERISTICS

Shoreline characteristics of interest during the lake assessment are described in Table D-6. To 

estimate the extent of major vegetation types, limit the assessment to the immediate lake shoreline 

area (i.e., within 20 m of the water). Also estimate the percentage of the immediate shoreline that has 

been developed or modified by humans.

Table D-6. Shoreline characteristics observed during lake assessment. (Source: Herlihy 1997.)

Characteristics: Description

Forest/Shrub Deciduous, coniferous, or mixed forest, including shrub and sapling vegetation

Agriculture Cropland, orchard, feedlot, pastureland, or other horticultural activity

Open Grass Meadows, lawns, or other open vegetation

Wetland Forested and nonforested wetlands (submerged terrestrial vegetation)

Barren Nonvegetated areas such as beaches, sandy areas, paved areas, and exposed rock

Developed Immediate shoreline area developed by human activity; this includes lawns, houses, stores, malls, marinas, 

golf courses, or any other human-built land use

Shoreline Modifications Actual shoreline that has been modified by the installation of riprap, revetments, piers, or other human 

modifications
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QUALITATIVE MACROPHYTE SURVEY

Macrophytes (aquatic plants large enough to be seen without magnification) can be important 

indicators of lake trophic status. The most important macrophyte indicator for assessment purposes is 

often the percentage of the lake area covered with macrophytes. For both "emergent/floating" and 

"submergent" coverage, choose one of the four percentage groupings (0 to 25 percent, 25 to 50 

percent, 50 to 75 percent, or 75 to 100 percent) that best describes the lake. In some cases, it will be 

fairly easy to estimate the percentage from observations made during sampling. In other cases, it will 

be an educated guess, especially if the water is turbid. After recording the areal percentage of 

macrophyte coverage, record the density of the plants in the observed macrophyte beds as dense, 

moderate, or sparse. Finally, provide any qualitative description (genera present, if known; dominant 

type—floating, emergent, or submergent) of the macrophyte beds that would be useful for 

interpreting the trophic status of the lake. All activities described in this subsection are recorded on 

the Lake Assessment portion of the Lake Sampling Site Documentation Form 1.

D-2.5 Post-Trip Activities

DATA FORMS AND SAMPLE INSPECTION

After all lake sampling and chain of custody forms are completed, one team member must review the 

data forms and sample labels for accuracy, completeness, and legibility. Confirm that the SiteID is 

correct on the forms, as well as the date of the visit. Verify that all information has been recorded 

accurately, the recorded information is legible, and any flags are explained in the comments section. 

Ensure that written comments are legible and use no "shorthand" or abbreviations. After reviewing 

the lake forms, the reviewer should initial the lower right corner of each page of the form. Ensure that 

all samples are labeled, all labels are completely filled in, and each label is covered with clear plastic 

tape.

LAUNCH SITE CLEANUP

If a boat or inflatable raft or float tube was used for lake sampling, inspect it for evidence of weeds 

and other macrophytes. Clean the boat or raft as completely as possible before leaving the launch site 

to minimize the possibility of transporting aquatic plant fragments or aquatic animals to other lakes 

where these species may not already occur. Clean up all waste material at the launch site and dispose 

of or transport it out of the site. 

PROCESSING SITE DOCUMENTATION DATA AND INFORMATION

A file system and database with reliable backup should be established for storage of site records and 

files, map images, and photos. Processing site documentation data and information include the 

following steps:

1. Retrieve site coordinates (and any tracks) from the GPS unit using the GPS software. Delete 

any extra coordinate sets (waypoints) and save the file.

2. Retrieve photos from the camera. 

3. Enter or revise the site record in the database.

a. Enter site coordinates obtained in the field.

b. Enter or revise the site description, travel and access directions. 

c. Add new photos as JPG images with captions to the site record. 

4. Create site maps providing both detailed and broader information for access and orientation. 

Annotate maps and pictures with text and arrows when it would be helpful. Note that the 
APPENDIX D. STANDARD OPERATING PROCEDURES FOR FIELD SAMPLING ACTIVITIES

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accuracy of maps varies and the coordinate-based points on the maps, as well as other 

information, may be misleading. Add clarifying notes. Save these maps as JPG images in the 

site record. Add captions as appropriate.

5. Enter site and visit data into NRM Air. Images and site maps can be attached to the site or visit 

database records.

Acknowledgements 

This SOP is based partly on material developed in the following publications:

Baker, J.R., and D.V. Peck. 1997. Section 4. Lake verification and index site location. In: Baker, J.R., 

D.V. Peck, and D.W. Sutton (eds.). Environmental Monitoring and Assessment Program 

Surface Waters: Field Operations Manual for Lakes. Report No. EPA/620/R-97/001. U.S. 

Environmental Protection Agency, Washington DC. 69p.

Baker, J.R., D.V. Peck, and D.W. Sutton, eds. 1997. Environmental Monitoring and Assessment 

Program Surface Waters: Field Operations Manual for Lakes. Report No. EPA/620/R97/001. U.S. Environmental Protection Agency, Washington DC. 69p.

Herlihy, A.T. 1997. Section 9. Final lake activities. In: Baker, J.R., D.V. Peck, and D.W. Sutton 

(eds.). Environmental Monitoring and Assessment Program Surface Waters: Field Operations 

Manual for Lakes. Report No. EPA/620/R-97/001. U.S. Environmental Protection Agency, 

Washington DC. 204p.

Linsley, R.K., M.A. Kohler, and J.L.H. Paulhus. 1982. Hydrology for Engineers. McGraw-Hill Book 

Co. New York. 508p.

Merritt, G.D.,V.C. Rogers, and D.V. Peck. 1997. Section 3. Base site activities. In: Baker, J.R., D.V. 

Peck, and D.W. Sutton (eds.). Environmental Monitoring and Assessment Program Surface 

Waters: Field Operations Manual for Lakes. Report No. EPA/620/R-97/001. U.S. 

Environmental Protection Agency, Washington DC.

Rantz, S.E. 1982. Measurement and Computation of Streamflow: Volume 1. Measurement of Stage 

and Discharge. U.S. Geological Survey Water-Supply Paper 2175. U.S. Government Printing 

Office, Washington, DC. Available at 

<http://pubs.usgs.gov/wsp/wsp2175/pdf/WSP2175_vol1a.pdf>.

247

APPENDIX E. 

DATA ENTRY FORMS AND INSTRUCTIONS 

FOR FIELD SAMPLING ACTIVITIES 

E-1 STREAM SAMPLING FORMS AND INSTRUCTIONS

There are two types of records used for collecting stream water samples: one documents sample site 

characteristics and the other documents that actual water samples. Site characteristics are documented 

on two forms called Stream Sampling Site Documentation Forms 1 and 2. These forms are used to 

document new sampling sites and to clarify and update existing site information.

The Stream Sample Records Forms 3, 4, and 5 are used to record each sample site visit. These forms 

are to be completed each time you visit a sample site and collect data.

Site information is required at the top of each form, but varies slightly.

FORM HEADER INFORMATION

Form headers provide basic site information. Each form header must be completely filled out in order 

for the water chemistry and field data to be associated to the correct site in a project and the NRM Air 

database. The minimum information needed is underlined in bold below. This information also 

includes the GPS information (latitude/longitude) on Form 1, under Site Verification.

Headers for Forms 1 and 3

1. Monitoring Project Name. Document the name of the project the monitoring site is assigned 

to (e.g., R8 Wilderness Stream Water Chemistry Survey or R9 Dry River Watershed 

Monitoring 2002). To ensure all the lab data is associated with the correct project and site it is 

very important to document the project name as it is displayed in NRM Air. If this is a new 

project document on the forms exactly what will be entered into NRM Air as the project name.

2. Forest Name. Document the National Forest where the sample will be collected.

3. Wilderness Name (if applicable). If the sample site is in a wilderness area, document the 

wilderness name.

4. Stream Name (USGS). Document the official USGS stream name where the sample site is 

located. In some cases, the stream will not have a name; in this case, enter “Unnamed Stream” 

or “Unnamed Tributary to [name of stream]”.

5. Stream Name (Local). Document the local stream name. In most cases this will be the same as 

the USGS stream name. In some cases, an unnamed USGS stream may have a local name 

associated with it.
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Site Name. Document the specific site name as there may be more than one site on a single 

stream there is a need for a unique sample site name. This unique site name should be 

developed at the project level. For example, you have a stream named “Mill Creek” with three

different sample site locations, you may call each site “Mill Creek SS1,” “Mill Creek SS2,” and 

“Mill Creek SS3.”

6. Site ID. Each sample site should have a unique Site ID, developed at the project level. The 

unique ID can be a mix of letters and numbers in any logical sequence (be sure to document the 

naming convention). The Site ID can contain characters for the project’s national forest and 

region. Using the example above, for the site name “Mill Creek SS1” on the Smokey Forest in 

Region 10, you may abbreviate the names and numbers in the ID: for example, in the Site ID

“R10SFMILLCKSS1”, R10 = Region 10, SF = Smokey Forest, MILLCK = Mill Creek, SS1 = 

Sample Site 1.

7. Date of Visit. Enter the date that you visited the sample site. Check the space according to 

whether this is the first, “Initial” visit to this site to establish it as part of a survey or monitoring 

effort or if this site has been visited and documented previously and therefore this is a 

“Subsequent” visit.

8. Field Team Leader Information. Enter the name, affiliation, phone number, and email 

address of the field person responsible for the sampling.

9. Access Information. Check all spaces that apply to the type of access to this site. Document 

any travel directions (attach a map if available), estimated time of travel, and any additional 

information that may be helpful in getting to this site.

Headers for Forms 2, 4, and 5

Note that each of these Forms has space near the top to enter information that identifies the stream, 

sample site, and the date of sampling. This is an abbreviated version of the Basic Site Information 

described above. It is important to fully complete this “header” information on each form to ensure 

that identifiers are consistent and related data are kept together.

The instructions for completing the following four attributes are described under Basic Site 

Information, above.

• Stream Name (Local): see item 4 above.

• Date of Visit: see item 7 above.

• Site Name: see item 5 above.

• Site ID: see item 6 above.

STREAM SAMPLING – SITE DOCUMENTATION FORM 1

Site Verification, GPS Information, and Tagging

Most of this data will be collected and recorded at the sample site.

1. Stream Verified. Does the available data match conditions observed on the ground sufficiently 

to verify that the intended sampling site has been located? Check Yes or No. Check all methods 

used to verify that you have located the correct site.

2. GPS Information. After establishing the sample site location, record the coordinates from the 

GPS unit: Latitude and Longitude in decimal degrees (if possible to six decimal places); 

include the Datum of the collected coordinates (e.g., NAD83). Record the Elevation in feet or 

meters, and, if the GPS unit has the capability, record its given accuracy in feet or meters.
APPENDIX E. DATA ENTRY FORMS AND INSTRUCTIONS FOR FIELD SAMPLING ACTIVITIES

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These coordinates will be the ones used in site identification and will be documented on the 

forms and in the NRM Air database. Each time the site is visited, use the same coordinates.

3. Site Tag. Record whether a metal tree tag has been affixed to a prominent tree in proximity to 

the sampling site. (Many wilderness areas do NOT allow the use of metal tags.) If a tag has 

been used, indicate the tree species to which the tag is attached. If the species is not known, 

record “unknown.” Record whether the tag was affixed to the tree on a previous trip (an 

existing tag) or newly placed on this trip (a new tag). Describe the tree and its location relative 

to the sampling site. Include, in the description, the height above the ground and compass 

bearing from the tag to the sampling site.

Site Assessment

The site assessment data will be collected from field observations (within 20 meters of the 

streambank) and recorded at the sample site.

1. Streambank Character. Estimate the percentages of vegetation types along the streambank 

(i.e., forest/shrub, herbaceous, wetland, or barren). Also look at the observed land use in the 

same area and identify shoreline modifications (e.g., a dock or riprap), development, and/or 

agricultural use. Use the following classes for these estimates (record the percent range on the 

form):

a. Rare (< 5%)

b. Sparse (5-25%)

c. Moderate (25-75%)

d. Extensive (> 75%)

2. The vegetation Dominant Age Class (forested areas only). If the site is located within a 

forested area, estimate the dominant age class of the trees. Check the appropriate age class 

space (i.e., 0-10 years, 10-25 years, 25-50 years, or > 50 years).

3. Dominant plant species. Observe the dominant plant species within the area of the sample site

and, if known, record them on Form 1.

4. Beaver activity. Document observed beaver activity; check the appropriate spaces on the form.

Watershed Assessment

Most of this information can be obtained in the office through geographic information system (GIS)-

based analysis before visiting the sample site. It can be helpful to have this information completed 

before the first sampling visit. This information will help to locate the targeted sample location. The 

form is useful for verifying in the field that the office GIS-based analysis appears to be a valid 

identification of watershed characteristics.

1. Vegetation cover. Using ortho-corrected or georeferenced aerial imagery in a GIS, estimate the 

different vegetation types, including exposed rock and tallus. Document this as a percentage of 

the watershed area above the sample site. Verify in the field whether this appears to be a valid 

identification.

2. Lithology. If GIS information on bedrock lithology is available, indicate the primary lithology 

(e.g., granitic, volcanic, or metamorphic). If there is more than one lithology, make note of 

additional significant lithology.

3. Watershed area. Using GIS, digitize the watershed area above the sample site (the watershed 

size will vary based on the sample site location). Calculate the watershed area in acres or 
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hectares. It will be helpful to determine the watershed area and boundary before determining 

other watershed assessment attributes.

4. Primary watershed aspect. The primary aspect of the watershed is determined by estimating 

the direction the watershed is facing based on the direction of stream flow. Indicate the aspect 

in degrees, with North as 0 ̊, East as 90 ̊, South as 180 ̊, and West as 270 ̊.

5. Average watershed slope. A simple way of calculating average slope percentage within the 

watershed boundary is to use digital elevation model (DEM) GIS files and Zonal Stats in GIS 

Spatial Analyst.

6. Stream order. Identify the stream order at the sampling site location using the Forest Service’s

National Hydrography Dataset (NHD).

STREAM SAMPLING – SITE DOCUMENTATION FORM 2

Photo Log

Take two photos at the sampling site, looking both upstream and downstream. On the initial visit to 

the site, it can be helpful to take additional photos to visually record stream and site characteristics 

(e.g., land use and vegetation). Record the photo ID or file name, date photographed, and description 

of the photo. It is recommended that these photos be attached to the site documentation forms in the 

project files in hardcopy once they are printed.

Description and Sketch of Site

Record a description and make a sketch of the site. This information can help in locating and 

characterizing the site, and may include access and tree tag information, land marks, and land use 

indicators.

Additional Notes

Add any additional information that may help to identify, locate, or describe this site.

STREAM SAMPLING – SAMPLE RECORD FORM 3

When collecting water chemistry data, care must be taken to protect samples from contamination. It 

is recommended that water samples be taken before the general observations (e.g., stream width and 

depth) are measured; this will ensure stream bottom sediments will not be stirred up before sampling, 

potentially contaminating the water samples.

Basic Site Information—Suggested Revisions

Examine information recorded on the Stream Sampling Site Documentation Forms for agreement

with site characteristics. Indicate any suggested revisions or updates to the site documentation form. 

Place a check mark next to all types of information requiring revision and explain the suggested 

revision in the space provided.

Water Samples and Replicates

1. Time Sampled. Indicate time of sample collection using a 24-hour clock (e.g., 4 p.m. is 

“1600”). Note that the time recorded on the bottle(s) and syringe(s) for the replicates should 

differ from the time recorded for the normal (regular) sample. This is important! The 

recommended protocol is to separate the sampling times for normal and replicate samples by 

one minute.
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2. Sample ID. Enter the unique identification code assigned to the sample (developed at the 

project level). These Sample ID codes are designed to be familiar to the project personnel and 

can represent the specific site, sample, type, and date, etc. For example, the Site ID combined 

with the sample type and/or date (e.g., SITEIDREG for a regular sample or SITEIDREP for a 

replicate sample). Note: Barcodes provided by ARML are a unique identifier and have a 

specific location for placement on the form (see item 6 below).

The sample ID represents a sample of water (bottle(s) and/or syringes) intended to represent 

conditions at a particular location, on a particular day, at a particular time. Note that multiple 

containers (bottle and/or syringes) obtained within one time window represent the SAME 

sample and receive the SAME ID code (and barcode). Replicated samples will receive different 

ID codes (and barcodes).

3. Sample Type. Record the type of sample collected (i.e., regular, replicate, or field blank).

4. Bottle Type. Record the type of bottle (or syringe) used for sample collection (i.e., plastic, 

glass, or syringe).

5. Number of Bottles. Record the number of samples (aliquots) collected for the normal (regular) 

sample and any replicates that may have been collected.

6. Bar Code. The barcode (a unique number/letter ID) will be prepared and provided by the 

ARML. Each year the lab provides new barcodes specific for that year. The barcode sheets will 

have multiple stick-on copies of the same barcode. These can be organized before field 

sampling and subsequently be affixed to the Stream Sampling Record Form, the Chain of 

Custody Form, and to each container (bottle or syringe) for the sample.

7. Collection Location. If the sample was not collected at the intended (targeted) location, 

explain the reason for changing the sample location.

General Observations

1. Air Temperature. Record the air and water temperature to the nearest degree and document 

the time (24hr) when it was measured. Record whether it is expressed in °C or °F.

2. Weather Condition (current and previous). Check the spaces that best describe the collectionday weather up to the time of sampling and the average weather over the previous three days (if 

known).

3. Stream Depth. Record the stream depth measured at sample site in mid-channel; check the 

space for the appropriate measurement units.

4. Stream Width. Record the stream width measured at the sample site; check the space for the 

appropriate measurement units.

5. Observed Discharge Level. Indicate the observed level of discharge in the stream at the time 

of sampling. Check the appropriate space (i.e., no flow, low flow, normal flow, or flood).

6. Delivery Method. Record the delivery method type.

STREAM SAMPLING – SAMPLE RECORD FORM 4

On-Site Water Data (optional)

If on-site water data were collected, record the measured values—the time measurements were taken, 

air temperature, and water temperature (at sample site). Check the space for the appropriate 

measurement units. Note: For field instrument data, express DO in units of mg/L and, if possible, 

percent DO. Correct specific conductance to 25o C. Record the measurements, instruments, and 

methods used for on-site water data collected.
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Photo Log

Take two photos of the sampling site looking both upstream and downstream. It can be helpful to take 

additional photos to visually record stream and site characteristics (e.g., land use and vegetation). 

Record the photo ID or file name, date photographed, and description of the photo. It is 

recommended that these photos be attached to the site documentation forms in the project files in 

hardcopy once they are printed.

Additional Notes

Add any additional information that may help to identify, locate, or describe this sample site.

STREAM SAMPLING – SAMPLE RECORD FORM 5

This form is for collecting stream water stage and discharge data.

General Information

Record the time this data was obtained and indicate what methods were used to collect an estimate of 

stream stage or discharge (check appropriate spaces). Complete the appropriate stage or discharge 

section on the form (Stage Measurement Only, Velocity-Area Procedure, or Timed Filling 

Procedure).

Stage Measurement Only

If stage measurements (estimates) were made in the field, record the measured value and indicate the 

unit of measure. Describe the location of measurement. Indicate if a rating curve has been developed 

with which to estimate discharge from stage measurements at this location, and indicate what the 

stage is referenced to (i.e., fixed staff gage or permanent landscape feature).

Discharge Measurement by Velocity-Area Procedure

If the velocity-area procedure was used to measure discharge, check “Yes” and indicate the units of 

measurement for water depth and velocity. Record the approximate width of the stream at the 

sampling location. Record the water depth and velocity in each of up to 20 evenly spaced intervals of 

the stream cross section.

Discharge Measurement by Timed Filling Procedure

If the timed filling method was used to measure discharge, check “Yes” and indicate the units of 

measurement for time and water volume. Record the time and volume measurements for five separate 

trials at each of up to three spillway locations.
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E-2 LAKE SAMPLING FORMS AND INSTRUCTIONS

There are two types of records used for collecting lake water samples: one documents lake 

characteristics and the other documents the actual water samples. Site characteristics are documented 

on two forms called Lake Sampling Site Documentation Forms 1 and 2. These forms are used to 

document new sampling sites and to clarify and update existing site information.

The Lake Sampling Record Forms 3 and 4 are used to record each sample site visit. These forms are 

to be completed each time you visit a sample site and collect data.

Site Information is needed at the top of each form, but varies slightly.

FORM HEADER INFORMATION

Form headers provide basic site information. Each form header needs to be completely filled out in 

order for the water chemistry and field data to be associated to the correct site in a project and the 

NRM Air database. The minimum information required to import this data into the NRM Air 

database is underlined in bold below. This also includes the GPS information (latitude/longitude) on 

Form 1 under Site Verification.

Headers for Forms 1 and 3

1. Monitoring Project Name. Document the name of the project the monitoring site is assigned 

to (e.g., R2 Long-Term Lake Monitoring or R2_R4 Wind River Mountains Bulk Deposition 

Monitoring). To ensure all the lab data is associated with the correct project and site, it is very 

important to document the project name as it is displayed in NRM Air. If this is a new project 

document on the forms exactly what will be entered into NRM Air as the project name.

2. Forest Name. Document the National Forest where the sample will be collected.

3. Wilderness Name (if applicable). If the sample site is in a wilderness area, document the 

wilderness name.

4. Lake Name (USGS). Document the lake name where the sample site is located. In some cases 

the lake will not have a name; in this case, document “Unnamed Lake” or “Unnamed Tributary 

to [name of lake]”.

5. Lake Name (Local). Document the local lake name. In most cases this will be the same as the 

USGS lake name. In some cases, an unnamed USGS lake may have a local name associated 

with it.

6. Site Name. Document the specific site name as there may be more than one site on a single 

lake there is a need for a unique sample site name. This unique site name should be developed 

at the project level. For example, you have a lake named “Deep Lake” with two different 

sample site locations (e.g., in the outlet and in the middle of the lake), you may call one site 

“Deep Lake Outlet” and the other site “Deep Lake Mid.”

7. Site ID. Each sample site should have a unique identifier (ID) and Site ID, developed at the 

project level. The unique ID can be a mix of letters and numbers in any logical sequence (be 

sure to document the naming convention). The Site ID can contain characters for the project’s 

national forest and region. Using the example above, for the site name “Mill Creek SS1” on the 

Smokey Forest in Region 10, you may abbreviate the names and numbers in the ID: for 

example, in the Site ID “R10SFMILLCKSS1”, R10 = Region 10, SF = Smokey Forest, 

MILLCK = Mill Creek, SS1 = Sample Site 1.
APPENDIX E. DATA ENTRY FORMS AND INSTRUCTIONS FOR FIELD SAMPLING ACTIVITIES

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8. Date of Visit. Enter the date you visit the sample site. Check the space according to whether 

this is the first “Initial” visit to this site to establish it as part of a survey or monitoring effort or 

if this site has been visited and documented previously and, therefore, this is a “Subsequent” 

visit.

9. Field Team Leader Information. Enter the name, affiliation, phone number, and email 

address of the field person responsible for the sampling.

10. Access Information. Check all spaces that apply to the access of this site. Document any travel 

directions (attach a map if available), estimated time of travel, and any additional information 

that may be helpful in getting to this site.

Headers for Forms 2 and 4

Note that each of these forms has space near the top to enter information that identifies the lake, 

sample site, and the date of sampling. This is an abbreviated version of the Basic Site Information 

described above. It is important to fully complete this “header” information on each form to ensure 

that identifiers are consistent and related data are kept together.

The instructions for completing the following four attributes are described under Basic Site 

Information above.

• Lake Name (Local): see item 4 above.

• Date of Visit: see item 7 above.

• Site Name: see item 5 above.

• Site ID: see item 6 above.

LAKE SAMPLING – SITE DOCUMENTATION FORM 1

Site Verification, GPS Information, and Tagging

1. Lake Verified. Does the available data match conditions observed on the ground sufficiently to 

verify that the intended sampling site has been located? Check Yes or No. Check all methods 

used to verify that you have located the correct site.

2. GPS Information. After establishing the sample site location record the coordinates from the 

GPS unit—Latitude and Longitude in decimal degrees (if possible to six decimal places)—

include the Datum of the collected coordinates (e.g., NAD83). Record the Elevation in feet or 

meters, and if the GPS unit has the capability, record its accuracy in feet or meters. These 

coordinates will be the one used for site identification and will be documented on the forms and 

in the NRM Air database. Each time the site is visited, use the same coordinates.

Lake Assessment

The lake assessment data will be collected from field observations and recorded at the lake.

1. Shoreline Character. Estimate the percentages of vegetation types within 20 meters of the 

shoreline (i.e., forest/shrub, herbaceous, wetland, or barren). Also, look at the observed land 

use in the same area, identify shoreline modifications (e.g., dock or riprap), development,

and/or agricultural use. Use the following classes for these estimates (record the percent range 

on the form):

a. Rare (< 5%)

b. Sparse (5-25%)

c. Moderate (25-75%)

d. Extensive (> 75%)
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2. Hydrologic Lake Type. Record the hydrologic type of the lake from the following:

a. Reservoir (artificial, human-made dam).

b. Drainage (outlet stream present; may or may not be flowing at time of visit).

c. Seepage (no outlet stream present, regardless of whether or not it is flowing at the time of 

visit). If a dam is present, select the appropriate type (e.g., artificial, augmented, or 

natural). Record the number of inlets to and outlets from the lake (if present).

3. Macrophyte Observation. From a quick visual survey of the lake, estimate the percent of lake 

area covered by “emergent/floating” and “submergent” macrophytes (rooted aquatic plants 

large enough to be seen without magnification). Select the appropriate percent class (i.e., 0-

25%, 25-50%, 50-75%, or >75%). Record the average density of the macrophyte community 

(i.e., absent, sparse, moderate, or dense). Identify (if known) the one to three most prevalent 

macrophyte species.

4. Lake Trophic State. Eutrophication is the process whereby a body of water becomes overenriched in nutrients, resulting in increased productivity of algae or aquatic plants (biomass) 

and sometimes also decreased dissolved oxygen levels. Based on the amount of biomass in the 

lake, record your estimation of the lake’s trophic state from the following classes:

a. Oligotrophic (low biomass production). Oligotrophic lakes are most common in cold 

regions underlain by resistant igneous rocks (especially granitic bedrock).

b. Mesotrophic (moderate or intermediate level of biomass productivity). These lakes are 

commonly clear water lakes and ponds with beds of submerged aquatic plants and 

medium levels of nutrients.

c. Eutrophic (high biomass productivity due to excessive nutrients, subject to algal blooms).

Eutrophic waters commonly lack fish species like trout, which require cold, welloxygenated waters.

d. Hypereutrophic (very high biomass productivity —nutrient-rich). Lakes are characterized 

by frequent and severe nuisance algal blooms and low transparency.

Watershed Assessment

Most of this information can be obtained in the office through geographical information system 

(GIS)-based analysis of aerial photography before visiting the sample site. It can be helpful to have 

this information completed before the first sampling visit. This information will help to locate the 

targeted sample location. The form is useful for verifying in the field that the office GIS-based

analysis appears to be a valid identification of watershed characteristics.

1. Vegetation cover. Using ortho-corrected or georeferenced aerial imagery in a GIS, estimate the 

different vegetation types including exposed rock and tallus. Document this as a percentage of 

the watershed area above the sample site. Verify in the field whether this appears to be a valid 

identification.

2. Lithology. If GIS information on bedrock lithology is available, indicate the primary lithology 

(e.g., granitic, volcanic, or metamorphic). If there is more than one lithology, make note of 

additional significant lithology.

3. Watershed area. Using GIS, digitize the watershed area above the sample site (the watershed 

size will vary based on the sample site location). Calculate the watershed area in acres or 

hectares. It will be helpful to determine the watershed area and boundary before determining 

other watershed assessment attributes.
APPENDIX E. DATA ENTRY FORMS AND INSTRUCTIONS FOR FIELD SAMPLING ACTIVITIES

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4. Watershed aspect. The primary aspect of the watershed is determined by estimating the 

direction the watershed is facing based on the direction of stream flow. Indicate the aspect in 

degrees with North as 0 ̊, East as 90 ̊, South as 180 ̊, and West as 270 ̊.

5. Average watershed slope. A simple way of calculating average slope percent within the 

watershed boundary is to use digital elevation model (DEM) GIS files and Zonal Stats in GIS 

Spatial Analyst.

6. Stream order. Identify the stream order at the sampling site location using the Forest Service’s

National Hydrography Dataset (NHD).

LAKE SAMPLING – SITE DOCUMENTATION FORM 2

Establish Lake Level Reference Point

On the initial visit (or subsequent visit) to the lake, establish a reference point for measuring lake 

level.

If it is allowed, permanently monument the lake level location by marking a shoreline rock (e.g., 

inserting a bolt). Check the appropriate space (Yes or No), indicating whether there is an established

permanent marker. If a permanent monument is not allowed, identify a landmark (e.g., “bathtub 

ring,” rock formation, or outcrop) from which to measure lake level. Record in the space provided the 

type of monument or landmark used to mark the reference point. Describe the lake level reference 

point location, with measurements and photos, on Form 2 under Description and Sketch of Lake and 

Sample Location and under Photo Log.

Measure the lake level at the time that the reference point is established; record the distance from lake 

level to the reference point—include measurement units (feet, inches, meters, or centimeters).

Description and Sketch of Lake and Sample Location

Record a description and sketch of the lake. This information can help in locating and characterizing 

the lake, and may include lake access, land marks, and land use indicators. Make sure to include 

sampling locations (e.g., mid-lake, shoreline, or outlet), lake level reference point location, and photo 

points.

Photo Log

Take photos of the lake sampling site to help identify the precise location of the sampling location. 

On the initial visit to the site, it can be helpful to take additional photos to visually record lake and 

site characteristics (e.g., land use and vegetation). Record the photo ID or file name, date 

photographed, and description of the photo. It is recommended that these photos be attached to the 

site documentation forms in the project files in hardcopy once they are printed.

Additional Notes

Add any additional information that may help to identify, locate, or describe this site.
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LAKE SAMPLING – SAMPLE RECORD FORM 3

Basic Site Information—Suggested Revisions

Examine information recorded on the Lake Sampling Site Documentation Forms for agreement with 

site characteristics. Indicate any suggested revisions or updates to the site documentation form. Place 

a check mark next to all types of information requiring revision; explain the suggested revision in the 

space provided.

Water Samples and Replicates

1. Time Sampled. Indicate time of sample collection, using a 24-hour clock (e.g., 4:00 PM is 

“1600”). Note that the time recorded on the bottle(s) and syringe(s) for the replicates should 

differ from the time recorded for the normal (regular) sample. This is important! The 

recommended protocol is to separate the sampling times for normal and replicate samples by 

one minute.

2. Sample ID. Enter the unique identification code assigned to the sample (developed at the 

project level). These Sample ID codes are designed to be familiar to the project personnel and 

can represent the specific site, sample, type, and date, etc. For example, the Site ID combined 

with the sample type and/or date (e.g., SITEIDREG for a regular sample or SITEIDDUP for a 

duplicate sample). Note: Barcodes provided by the ARML are a unique identifier and have a 

specific location for placement on the form (see item 6 below).

The sample ID represents a sample of water (bottle(s) and/or syringes) intended to represent 

conditions at a particular location, on a particular day, at a particular time. Note that multiple 

containers (bottle and/or syringes) obtained within one time window represent the SAME 

sample and receive the SAME ID code (and barcode). Replicated samples will receive different 

ID codes (and barcodes).

3. Sample Depth. Record the depth at which the sample was collected. Indicate the units of 

measure (ft or m).

4. Sample Depth Zone. Record the limnetic zone (i.e., epilimnion or hypolimnion) in which the 

sample was collected. If the limnetic zone is not known record either “surface” or “deep”.

5. Sample Type. Record the type of sample collected (i.e., regular, replicate, or field blank).

6. Bottle Type. Record the type of bottle (or syringe) used for sample collection (i.e., plastic, 

glass, or syringe).

7. Number of Bottles. Record the number of sample aliquots (bottles or syringes) collected for 

the normal (regular) sample and any replicates that may have been collected.

8. Bar Code. The barcode (a unique number/letter ID) will be prepared and provided by ARML.

Each year the lab provides new barcodes specific for that year. The barcode sheets will have 

multiple stick-on copies of the same barcode. These can be organized before field sampling and 

subsequently be affixed to the Lake Sampling Record Form, the Chain of Custody Form, and to 

each container (bottle or syringe) for the sample.

9. Collection Location. If the sample was not collected at the intended (targeted) location, 

explain the reason for changing the sample location.
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General Observations

1. Record the Time Obtained (24hr) when the general observations were made.

2. Air Temperature and Water Temperature. Record the air and water temperature collected to 

the nearest degree; check the units in °C or °F. Document the instrument used for measurement 

(e.g., certified thermometer or Clinefinder).

3. Weather conditions (current and previous). Check the spaces that best describe the collection 

day weather up to the time of sampling, and the average weather over the previous three days 

(if known).

4. Lake Level Change. Measure the distance from lake level to the reference point (see initial or 

previous years established reference point for location) and record the measurement plus or 

minus (+ / -) the reference point—include measurement units (i.e., ft, in, m, or cm). If there is 

no change from the reference point, mark the “zero change” space on the form.

5. Observed Discharge Level. Indicate the observed level of discharge from the lake at the time 

of sampling. Check the appropriate space (i.e., no flow, low flow, normal flow, high flow, or 

flood).

LAKE SAMPLING – SAMPLE RECORD FORM 4

Transparency Data

Record two depth measurements using the Secchi disk, 1) when the disk disappears and 2) when the 

disk reappears; select the appropriate unit of measurement. If the Secchi disk was visible to the 

bottom of the lake, check the space for “Yes” beside Clear to Bottom?

Depth/Temperature Profile

1. Method/Instrument. Document the EPA/SM/USGS method used for the measurements, as 

well as the make and model of the equipment used for the profile.

2. Index Location and Depth. Indicate whether the depth profile was measured at the index site 

or not. If the profile was collected in a different location, explain the reason. Record the total 

depth at the location of the profile and select the appropriate units.

3. Measurements. Collect the depth and measurements for that depth; record the depth and 

measurement values in the profile table for each parameter. Check the appropriate space 

indicating units of measure (temperature and DO). Indicate whether conductivity measurements 

are corrected to 25oC.

On-Site Water Data (optional)

If on-site water data were collected, record the measured values: the time measurements were taken, 

air temperature and water temperature (at sample site). Check the space for the appropriate 

measurement units. Note: Express DO in units of mg/L and, if possible, percent DO. Correct specific 

conductance to 25o C. Record the measurements, instruments, and methods used for on-site water 

data collected.

Photo Log

Take photos of the lake sampling site that will help identify its precise location. It may be helpful to 

take additional photos to visually record lake and site characteristics (e.g., land use and vegetation). 

Record the photo ID or file name, date photographed, and description of the photo. It is 

recommended that these photos be attached to the site documentation forms in the project files in 

hardcopy once they are printed.
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E-3 CHAIN OF CUSTODY FORM AND INSTRUCTIONS

1. Page ___ of ___. Page number(s) of total number of chain of custody forms sent. 

2. Forest/Wilderness/Park/Other (Circle one). Circle one of these options and write the name 

of the national forest, wilderness area, national park, or other area (e.g., specific unit within the 

national forest) in the space provided. Provide the name and affiliation of the project contact 

individual. 

3. Address and Phone Number. Provide address and phone number of the office of the national 

forest, wilderness, national park, or other area. In the address, please include the city, state, and 

zip code. 

4. Shipped to (Lab Name and Address). Name, address, and email of the laboratory to which 

the water samples and original Chain of Custody Form will be sent. 

5. Lab Phone #. Phone number of the laboratory to which the water samples and original Chain 

of Custody Form will be sent. 

6. Lab Contact and Email. Contact person and email address in the analytical laboratory. 

7. Shipped by: UPS/Fed Ex/USPS/Other. Identify the carrier that you used. (Remember to 

consider the arrival date of the shipped samples because, on weekends and government 

holidays, there may not be anyone to receive samples at the laboratory). In general, you should 

try to ship samples on Monday, Tuesday, or Wednesday in order to arrive before the weekend. 

8. Shipping #. Tracking number assigned to the shipment by the carrier. 

9. Sampled Date. Date sample was taken (mm/dd/yyyy). 

10. Sample Time. Time sample was taken (24-hr: ####). 

11. Site/Sample ID. The unique identification number assigned to the sample in the field based on 

the specific protocol. Refer to the lake or stream form instructions for more detail on 

developing a site ID before field collection (see header instructions, item 6).

12. Site/Sample Location. Document the lake or stream sample site name. Refer to the lake or 

stream form instructions for more detail on developing a site name before field collection (see 

header instructions item five). Provide the latitude and longitude in decimal degrees. 

13. Sample Type. Document the type of sample collected, such as normal (regular) water sample, 

field blank, replicate (duplicate), or split. 

• Replicate samples are collected at the same location as the normal water sample but at 

slightly different times (typically, one minute apart). Replicates are usually collected for QA

purposes or as backup samples should the normal sample be lost or damaged. 

• A field blank is a prepared sample of DIW that is carried into the field and then shipped to 

the laboratory with the samples.

• A field split is the second sample bottle when a normal sample has been split in the field into 

two bottles. The first bottle is labeled as the normal sample; the second is labeled as the field 

split (S).

14. Filtered (Y/N). Where? Was this sample filtered in either the field or field laboratory? If so, 

where?

15. Preserved (Y/N/Type). Was this sample preserved in the field and, if so, with what kind of 

preservative (e.g., H2SO4)?
APPENDIX E. DATA ENTRY FORMS AND INSTRUCTIONS FOR FIELD SAMPLING ACTIVITIES

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16. Analyses Requested. Document instructions for the laboratory, requesting the type of analyses 

to be performed (e.g., ANC, pH, conductivity, major cations, anions, etc.). You may write 

“same as usual” or leave the space blank if you have an agreement with the laboratory 

concerning routine analyses. 

17. Bar Code. The barcode is a unique identifier provided by ARML for tracking purposes. The 

multiple (4) stick-on copies of the barcode ID labels are prepared to be affixed in the field to 

multiple forms and sample containers.

18. Comments. Any extra remarks or instructions are placed in this space. 

19. Received/Relinquished by:

• Print Name. Printed name of sampler relinquishing the samples to another person for 

shipment to the laboratory or directly to the laboratory. 

• Signature. Sampler’s signature relinquishing the samples to another person for shipment to 

the laboratory or directly to the laboratory. 

• Date & Time Relinquished. Date and time relinquished by the sampler or by person 

shipping samples to the laboratory.

• Date & Time Received. Date and time samples were received from the sampler. 

20. Received at Laboratory by: 

• Print Name. Printed name of laboratory personnel receiving the samples. 

• Signature. Signature of the laboratory personnel receiving the samples. 

• Date. Date the samples were received by the laboratory. 

• Time. Time the samples were received by the laboratory. 

It is extremely important to send this form and accompanying lake or stream sampling record forms

to the laboratory with the samples so that proper connections can be made between field and 

laboratory information and so that relevant data may be entered into the ARM program database.
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271

APPENDIX F. 

LABELING INSTRUCTIONS FOR FIELD 

SAMPLE CONTAINERS 

F-1 WATER SAMPLE

Below is an example of what should be on the label for the water samples. These can be premade 

labels or filled in on a blank label at time of sample collection. Apply a separate label to each sample 

container (e.g., bottle or syringe). Placing one or more strips of packing tape over the label once it is 

on the container can protect the label from damage and subsequent misreading.

Lake or Stream Name. Enter the name of the lake or stream sampled. Provide both the USGS name 

(from a topographical map) and the local name by which the lake or stream is known. 

Site ID Number. Each sample site (location) will be assigned a unique Name and ID number. These 

are generated locally for the project. The identification number will appear on all sample 

bottles used for sampling this lake or stream at this location. It is especially important to have 

a unique Site ID when more than one sample site is located on a given lake or stream. 

Sample Date. Enter the date when visiting the sampling site. 

Sample Time (24-hr). Indicate time of arrival at sampling location. Use 24-hour (military time) 

format. Indicate whether it is recorded in standard local time or daylight savings local time. 

Collected By. Enter the name and affiliation of the field person responsible for the sampling. 
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Sample Type. Check the space indicating whether this is a normal sample, a replicate sample 

(Replicate 1 or Replicate 2), or a field blank or field split sample. 

Barcode. Affix the same numbered bar code represented on the form from the sheets supplied by the 

lab to the associated sample container. It is very important to make sure each sample has a 

unique bar code. Do not use the same bar code on different samples (e.g., normal and 

replicate).

F-2 ZOOPLANKTON SAMPLE

Below is an example of what should be on the label for the zooplankton sample containers. These can 

be premade labels or written on a blank label at time of sample collection. In addition to the label 

affixed to the outside of the sample bottle, a label must be placed inside the bottle of preserved 

sample. This internal label, on waterproof paper, must be filled out with a pencil.

Lake Name. Enter the name of the lake or stream sampled. Provide both the USGS name (from a 

topographical map) and the local name by which the lake or stream is known. 

Site ID Number. Each sample site (location) will have a unique Name and ID number assigned to it.

These are generated locally for the project. The identification number will appear on all 

sample bottles used for sampling this lake or stream at this location. It is especially important 

to have a unique Site ID when more than one sample site is located on a given lake or stream. 

Sample Date. Enter the date when visiting the sampling site. 

Sample Time (24-hr). Indicate time of arrival at sampling location. Use 24-hour (military time) 

format. Indicate whether it is recorded in standard local time or daylight savings local time. 

Collected By. Enter the name and affiliation of the responsible field person. 

Sample Depth. Record the water depth at which the sample was collected. Check whether the depth 

is being recorded in meters or feet. 

Net Mesh Size. Check whether the net used to collect the sample was of mesh size 80 μM, 243 μM, 

or some other size. If the mesh was of a size other than 80 μM or 243 μM, specify the mesh 

size used. 
APPENDIX F. LABELING INSTRUCTIONS FOR FIELD SAMPLE CONTAINERS

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F-3 STREAM BENTHIC MACROINVERTEBRATE SAMPLE

Below is an example of what should be on the label for the macroinvertebrate sample containers.

These can be premade labels or written on a blank label at time of sample collection. In addition to 

the label affixed to the outside of the sample bottle, a label must be placed inside the bottle of 

preserved sample. This internal label, on waterproof paper, must be filled out in pencil. 

Stream Name. Enter the name of the lake or stream sampled. Provide both the USGS name (from a 

topographical map) and the local name by which the lake or stream is known. 

Site ID Number. Each sample site (location) will have a unique Name and ID number assigned to it. 

These are generated locally for the project. The identification number will appear on all 

sample bottles used for sampling this lake or stream at this location. It is especially important 

to have a unique Site ID when more than one sample site is located on a given lake or stream. 

Sample Date. Enter the date when visiting the sampling site. 

Sample Time (24-hr). Indicate time of arrival at sampling location. Use 24-hour (military time) 

format. Indicate whether it is recorded in standard local time or daylight savings local time. 

Collected By. Enter the name and affiliation of the field person responsible for the sampling. 

Number of Kick Net Samples Collected. Record the number of discrete kick net samples that were 

collected and pooled to form this one sample being submitted to the laboratory.

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APPENDIX G. 

EXAMPLE STANDARD OPERATING 

PROCEDURES FOR LAB ANALYSIS 

PROTOCOLS 

G-1 USGS STANDARD OPERATING PROCEDURE FOR 

LABORATORY ANALYSIS OF ONE ANALYTE 

(AMMONIUM) IN DILUTE FRESH WATER – FLOW 

INJECTION ANALYSIS

1. Scope and Application

Analytes: Ammonium

Reporting Limit: 2.0 μmoles/L

Applicable Matrices: This method is used to determine the concentration of ammonium in 

precipitation, dilute surface waters, and soil waters.

Dynamic Range: The analytical range of the determination of ammonium is from 0.5 μmol 

NH4

+

/L (as N) to 35.7 μmol NH4

+

/L (as N). Sample concentrations that exceed this range must be 

diluted and reanalyzed.

2. Summary of Procedure

The ammonium analysis is an automated colorimetric reaction. Samples are systematically 

introduced into the flow-injection analyzer (FIA) reaction manifold. The sample reacts with 

phenol and hypochlorite to form an indophenol blue complex. The color intensity of the blue 

complex is enhanced by mixing with nitroferricyanide. The mixture is heated to 60°C to ensure 

optimal color development. Prior to entrance into the flow cell, the sample flows through a 

debubbler to remove any gas bubbles that develop. Absorbance of the color complex is measured 

at a wavelength of 630 nm.

3. Safety Issues

Chemical Hazards: 

1. All strong acids and bases should be mixed in a fume hood.

2. Gloves, safety glasses, and lab coats should be worn when preparing and performing this 

analysis.
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3. For proper handling techniques for specific chemicals, consult the appropriate Material 

Safety Data Sheets (MSDS).

4. Sample Preservation, Containers, Processing and Analysis Times

Sample Preservation: Samples are frozen after processing. On the day of analysis, samples are 

thawed in warm water or at room temperature.

Containers: Samples are stored in 30-mL polyethylene bottles. The bottles have been rinsed with 

DIW.

Processing and Analysis Times: 

Sample processing: one week

Lab analysis: three months

LIMS entry: one week

5. Reagents and Standards

General Information: All reagents are commercially purchased and should be stored in the 

original container. Date the reagent bottles when received and when opened. Note expiration date, 

if any. No verification of the reagents is necessary. 

Reagents: 

1. Degassed Deionized (DI) Water

a. Milli-Q water is degassed by bubbling with commercial-grade helium for about 2 

minutes.

b. Degassed Milli-Q water is used for carrier water and for preparation of all reagents.

2. Sodium Phenolate

a. Liquid phenol (C5H5OH) should be stored in a designated flammable-storage cabinet. 

b. In a 1,000-mL volumetric flask containing about 500 mL Milli-Q water, add 88-mL 

liquefied phenol.

c. Add 32 g sodium hydroxide (NaOH); swirl to dissolve and allow to cool.

d. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again.

e. Store in a red glass bottle at 4°C; label and date.

f. Prepare every other day.

3. Sodium Hypochlorite

a. Use household bleach containing at least 5.25 percent sodium hypochlorite (NaOCl).

b. Fill a specially designated 1-L polyethylene bottle to the 500 mL mark with Milli-Q 

water and then add bleach to the 1-L mark.

c. Shake thoroughly.

d. Store in a polyethylene bottle at 4°C; label and date.

e. Prepare every other day.

4. Sodium Nitroferricyanide

a. In a 1,000-mL flask containing about 500 mL Milli-Q water, add 3.50 g sodium 

nitroferricyanide (Na2Fe(CN)5NO•2H2O).
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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b. Swirl to dissolve the solid. 

c. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again.

d. Store in a polyethylene bottle at 4°C; label and date.

e. The solution is stable for 3 months.

5. Cleaning Solution

a. In a 1,000-mL volumetric flask containing about 700 mL Milli-Q water, carefully 

add 82.5 mL concentrated hydrochloric acid (HCl). 

b. Allow to cool. 

c. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again.

d. Store in a polyethylene bottle; label and date. 

e. Prepare as needed.

Standards: 

1. Ammonium Standard Stock Solution, 1,000 mg NH4

+

/L (as N)

a. Purchased commercially.

2. Ammonium Standard Substock Solution, 5 mg NH4

+

/L (as N)

a. Pipet 0.5 mL of the ammonium standard stock solution into a designated 100-mL 

volumetric flask containing about 50 mL of Milli-Q water.

b. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again.

c. Store in the volumetric flask at 4°C; label and date.

d. Prepare every other day. 

e. To avoid contamination, aliquots of substock solution must not be withdrawn directly 

from the bottle.

3. Ammonium Working Standards

a. Pipet desired amount of standard substock into a designated 100-mL volumetric 

flask.

b. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again.

c. Store in the volumetric flask at 4°C; label and date.

d. Prepare every other day.
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Working Standard Ammonium Concentration Standard Substock Added (mL) Final Volume (mL)

A 35.70 μmol/L (0.500 mg/L) 10.0 100

B 26.77 μmol/L (0.375 mg/L) 7.5 100

C 17.85 μmol/L (0.250 mg/L) 5.0 100

D 10.71 μmol/L (0.150 mg/L) 3.0 100

E 7.14 μmol/L (0.100 mg/L) 2.0 100

F 3.57 μmol/L (0.050 mg/L) 1.0 100

G 1.78 μmol/L (0.025 mg/L) 0.5 100

4. Ammonium Quality-Control (QC) Stock Solution, 1,000 mg NH4

+

/L (as N)

a. Purchased commercially, this stock must be from a manufacturer or lot different from 

the standard stock.

5. Ammonium Quality-Control (QC) Substock Solution, 5 mg NH4

+

/L (as N)

a. Pipet 0.5 mL of the ammonium QC stock into a designated 100-mL volumetric flask 

containing about 50 mL of Milli-Q water.

b. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again. 

c. Store in the volumetric flask at 4°C; label and date.

d. Prepare every other day.

e. To avoid contamination, aliquots of substock solution must not be withdrawn directly 

from the bottle. 

6. Ammonium QC Samples

a. Pipet desired amount of QC substock into a designated 250-mL volumetric flask.

b. Fill close to final volume with Milli-Q water, mix, then fill to final volume and mix 

again.

c. Store in the volumetric flask at 4°C; label and date.

d. Prepare every other day.

QC Sample Ammonium Concentration QC Substock Added (mL) Final Volume (mL)

High 17.85 mol/L (0.250 mg 12.5 250

Low 7.14 mol/L (0.100 mg/ 5.0 250

6. QC Procedure

1. The standard curve is a linear plot of standard concentration versus peak area. The best-fit 

line is drawn and the curve is accepted if the correlation coefficient is 0.998 or greater.

2. Quality-control samples are analyzed at the start of a run, after every 10 samples during the 

run, and at the end of the run.

3. A quality-control sample is acceptable if the analyzed value is within 10 percent of the QChigh range known value and within 15 percent of the QC-low range known value.
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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4. If one of the QC samples fails the acceptance criteria, the run is stopped and the QC sample 

is re-run. If the QC sample fails again the run is stopped and the instrument is re-calibrated.

Samples associated with the failed QC sample are re-analyzed.

7. Chemical Analysis Procedure:

Instrumentation: 

Lachat QuickChem 8000 flow-injection analyzer

Omnion software v 3.0

Timing: 

Method Cycle Period - 90 seconds

Auto-sampler Timing Sample Period - 41 seconds

Minimum Probe in Wash Period - 41 seconds

Settings Ammonium Channel (seconds)

Time to Valve 26

Load Period 36

Inject Period 53

Expected Inject to Peak Start 24

Expected Peak Base Width 86

Figure G-1. Flow diagram for ammonium manifold.
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Start-Up:

1. Turn the surge protector power on.

2. On the computer desktop, double click the Omnion 3.0 icon. 

3. Check the waste container daily and empty if nearing capacity.

4. Fill DIW reagent bottle with degassed Milli-Q water.

5. Place pump tubes in appropriate reagent containers- see diagram. Cover reagent bottle 

openings with parafilm.

6. Push down cartridges on pump until they click into place. Pull tighteners on pump 

cartridges all the way to right and then forward one click.

7. Pump speed setting should be 35. Turn pump on.

8. Allow reagents and Milli-Q water to pass through system for at least 10 minutes.

9. Check digital readout on heater- it should read 60°C.

10. Observe module connections for any leaks or clogs.

11. Empty debubbler by turning upside down, then right side up.

Calibration:

1. Click OPEN. Double click the CALIB+QC+NH4.omn file. 

2. At the top of the screen the heading should read Omnion – Run 1 (CALIB+QC+NH4.omn).

3. Fill cups on calibration rack with standards and QC samples.

4. Click green START arrow.

5. If the calibration is within acceptance criteria, the instrument will analyze the QC samples.

If the calibration fails, a message will pop up, with options to proceed. In most cases 

choose Recalibrate.

6. If the calibration and QC samples pass, set up a sample tray. 

Analysis:

1. Take ammonium samples out of the freezer and thaw in warm water.

2. Enter the sample serial numbers (SSNs) into run worksheet. Use the Auto Sample ID 

feature if the SSNs are consecutive.

3. From menu bar select RUN. Click EXPORT WORKSHEET DATA. Note date on run 

worksheet printout.

4. Delete calibration samples from the run worksheet.

5. Fill the auto sampler tubes with the appropriate samples.

6. Empty debubbler.

7. Click green START arrow.

8. Fill QC containers and carrier bottle as needed. 

9. QC samples will be run automatically every 10 samples. If all QC samples pass, the run 

will proceed until the tray is finished.
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10. If a QC sample fails, a message will pop up and analysis will cease. Click the Stop Now 

option. Remove SSNs that have been analyzed from the run worksheet.

11. Click green Start arrow.

12. If QC samples pass, the run will continue. If QC samples fail, remake QC samples from 

substock and/or recalibrate.

13. Review the sample peaks and analysis data as the run progresses. Note any air bubbles, bad 

peaks, and/or samples requiring dilutions or re-runs.

Shut Down:

1. Change the pump setting to override standby.

2. Remove pump tubes from reagent bottles and rinse lines and weights with Milli-Q water.

Place lines in a beaker of Milli-Q water.

3. After several minutes, place pump tube reagent lines in the cleaning solution. Allow to 

pump for about 2 minutes.

4. Rinse and place pump tube reagent lines in Milli-Q water again. Allow to pump for several 

more minutes.

5. Remove lines from beaker and allow to pump air until no more water is moving through 

manifold.

6. Release tension of cartridges by pushing holders on side of pump and push tension 

regulators on top of the cartridges all the way to the left.

7. Exit Omnion.

8. Turn the surge protector off.

Maintenance:

1. All pump tubes should be replaced as they become worn or stretched; the frequency 

depends upon the number of samples analyzed. The sample and wash bath tubes should be 

changed as they become discolored or clogged. Note dates in instrument notebook.

2. Interference filters should be cleaned with lens paper twice a year or whenever they become 

dusty or soiled.

3. Manifold tubing should be replaced as it becomes discolored or clogged; note in instrument 

notebook.

4. Sample tubes are rinsed, soaked in DIW overnight, and oven dried at 60°C between uses.

5. Buildup and clogging in the waste lines may require periodic replacement of the lines; note 

in instrument notebook.

Data Processing and LIMS Entry:

1. In Microsoft Excel, open all files for the run date. They are stored in C:\Program 

Files\Lachat\Omnion\Data. All files are in a comma delimited text format.

2. Edit files as needed and copy/paste them all into the same file.

3. Save as a comma delimited file (.csv file extension) named by date.

4. Print a copy and write the filename on the copy and close file.
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5. Double click Watershed LIMS icon.

6. Click Import Data.

7. Under the Import drop down, choose Ammonium.

8. Choose and open the desired file.

9. Choose Client, Units, type in Test Date, and choose Analyst.

10. Exclude and/or edit any data necessary.

11. Click Client ID to Sample No.

12. Click Set Data.

13. Investigate problems for data that did not transfer or are duplicated.

8. Archiving

Data: Data files are backed-up daily by an automatic back-up program. Hard copies of the runs 

are filed and kept indefinitely. The laboratory LIMS system is backed up daily by automatic 

back-up program.

Samples: Samples are stored at room temperature until data can be verified. Sample bottles are 

cleaned and reused for new samples.

G-2 UNIVERSITY OF VIRGINIA STANDARD OPERATING 

PROCEDURE FOR LABORATORY ANALYSIS:

SHENANDOAH WATERSHED STUDY/VIRGINIA 

TROUT STREAM SENSITIVITY STUDY

R.F. Webb, R., F.A. Deviney, and S.W. Maben (unpublished manuscript)

G-2.1 Metrohm Titrando 809 Titration System: pH and ANC

This document describes the current laboratory procedure for measuring pH and ANC using the 

Metrohm Titrando 809 Titration System. Analytical methods are based on the methods published 

in U.S. Environmental Protection Agency (1987).

Summary: pH is measured first while the sample is not stirred. The measurement of ANC is a 

multi-step process. The stirrer comes on, and the sample is titrated with 0.01 N HCl to endpoints 

of pH 4.5 and 4.2, and the pH and volume of acid are recorded for each endpoint. Then eight 

equal-volume aliquots of acid are added, and the pH recorded after each addition. Finally the 

sample is titrated to a pH 3.5, and the pH and volume of acid added is recorded. These 11 pHvolume data pairs are used to calculate the Gran1 ANC.

Section I: Standards

Use purchased NIST-traceable pH 4 and 7 buffer solutions to calibrate the electrode. Use 

purchased NIST-traceable 0.01 N HCl volumetric solution for the titrant and to prepare the pH 

4.60 QCS (Quality Control Sample).
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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Preparation of pH 4.60 QCS (Quality Control Sample):

All glassware should be Class A volumetric laboratory glassware which has been initially cleaned 

by acid washing and afterwards cleaned by multiple rinsings with DI (deionized) water.

• Using a volumetric pipette, add 5 mL of 0.01 N HCl to a 2000 mL volumetric flask partially 

filled with DIW. Dilute to the line with DIW and mix well.

• Store in volumetric flask at room temperature.

Section II: Operation

Analyzing Samples:

1. Log in to Titrator computer

2. Turn on 730 Sample Changer with on/off switch near the bottom on the side

3. After Sample Changer has initialized, open Tiamo software

4. Click “OK” to “Prepare dosing device” reminder

5. Enter your name as “User” and the date as “Remark”

6. Check level of titrant (0.01 N HCl). Fill reservoir if needed

7. Fill beaker in rack position 13 half-full with fresh DIW

8. Open Manual Control under Tools menu or by clicking on icon (looks like a pointing hand)

9. Move sample changer to rack position 13:

a. Choose Tower 1

b. Select Move tab

c. Enter Rack position: 13, then click Start

d. Enter Lift position: 190 mm, then click Start

10. Prepare Dosing device:

a. Choose Dosing device 2 (800)

b. Select Prepare tab

c. Click Start (Answer yes to Splash warning). This will take a few minutes.

11. Click “Sample table  Import data” to import a spreadsheet which has been generated by 

the SWAS/VTSSS LIMS (Laboratory Information Management System) to create the 

Series list or follow instructions to create Series list manually (see section at end entitled 

“To create Series List manually”)

12. Flush tubing: 

a. Select Add fixed volume tab

b. Enter Volume: 6 mL

c. Enter Dosing rate: maximum

d. Enter Filling rate: 5 mL/min

e. Click Start (While the 6 mL is being delivered, make sure any bubbles in the tubing 

are being flushed out.) This will take a few minutes.

13. Fill the following Sample Changer positions: (Rinse each beaker 10 times with DIW, then 

with a little of the solution.)
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a. Rack position 1 = 100 mL pH 7 buffer (approximate volume)

b. Rack position 2 = 100 mL pH 4 buffer (approximate volume)

c. Rack position 14 = Diluted pH 7 buffer for electrode storage (half full)

14. Raise tower head to install electrode:

a. Choose Tower 1

b. Select Move tab

c. Enter Lift position: 0 mm, then click Start

15. Prepare electrode:

a. Carefully remove electrode from storage sleeve

b. Open fill hole

c. Replace filling solution in electrode (3M KCl) weekly or more often as necessary by 

using a transfer pipette to remove old filling solution and then filling electrode with 

new filling solution to just below the fill hole

d. Shake downward gently to remove air bubbles if filling solution was changed

e. Rinse bottom half of electrode with DIW

f. Connect electrode and place in tower head

16. Place electrode in diluted pH 7 buffer:

a. Choose Tower 1

b. Select Move tab

c. Enter Rack position: 14, then click Start

d. Enter Lift position: 190 mm, then click Start

17. Fill Sample Changer positions with samples according to Series list. (You can fill a few 

positions ahead and then start the Series. Continue to fill the rest once the Series has 

started.)

a. Rinse beaker 10 times with DIW

b. Rinse 100 mL measuring volumetric flask 3 times with DIW

c. Rinse 100 mL measuring volumetric flask with a little sample

d. Fill 100 mL volumetric flask with sample as full as possible (will slightly rise over 

top)

e. Dry excess liquid sample from sides of flask so that it won’t drip into beaker

f. Carefully transfer all of sample into beaker

18. Put fresh DIW in beakers in rack positions 12 & 13

19. Rinse electrode:

a. Choose Tower 1

b. Select Move tab

c. Enter Lift position: 0 mm, click Start

d. Enter Rack position: 12, click Start

e. Enter Lift position: 190 mm, click Start

f. Choose Stirrer 1 (Tower)

g. Click Start (let stir for ~ 10 seconds)

h. Click Stop
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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20. Close Manual Control

21. Click Start to begin Determination Series

22. Fill the rest of the Sample Changer positions according to the Series list.

Notes:

a. After the calibration is completed, Rack position 12 can be used for samples.

b. Rack position 13 is always for the rinse water throughout the entire run. Replace with 

fresh DIW occasionally throughout the run.

c. Rack position 14 is always for diluted pH 7 buffer for electrode storage at the end of the 

run.

Shutdown Procedure:

1. Open Manual Control under Tools menu or by clicking on icon

2. Remove electrode:

a. Choose Tower 1

b. Select Move tab

c. Enter Lift position: 0 mm, click Start

d. Remove electrode from tower head and disconnect

e. Rinse bottom half of electrode with DIW

f. Close fill hole

g. Carefully place electrode in storage sleeve filled with electrode storage solution.

Refill sleeve only with electrode storage solution (not 3M KCl electrode filling 

solution).

3. Rinse stirrer and tip:

a. Fill beaker in rack position 12 with fresh DIW

b. Choose Tower 1

c. Select Move tab

d. Enter Rack position: 12, click Start

e. Enter Lift position: 190 mm, click Start

f. Choose Stirrer 1 (Tower)

g. Click Start (let stir for ~ 10 seconds)

h. Click Stop

4. Move stirrer and tip to storage:

a. Fill beaker in rack position 13 with fresh DIW

b. Choose Tower 1

c. Select Move tab

d. Enter Lift position: 0 mm, click Start

e. Enter Rack position: 13, click Start

f. Enter Lift position: 190 mm, click Start

5. Close Manual Control

6. Exit Tiamo
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7. Turn off Sample Changer

8. Log off of computer

9. Rinse all beakers 10 times with DIW and put on rack to dry

Section III: Quality Control

The pH 4.60 QCS is analyzed at the beginning and end of each Series and throughout the Series 

(approximately every 8 samples).

The Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance 

Project Plan of this SOP gives the following precision objective for pH values less than the 

transition value of pH 5.75: + 0.075 pH units.

Therefore the pH value of the QCS should be 4.525 – 4.675. If it is not, stop the run to determine 

the problem. If the QCS fails, all samples run since the last passing QCS must be re-analyzed

To check the measured value of the pH 4.60 QCS during the run:

1. Click on the Database icon on the left side of the Tiamo screen

2. Click on the page arrows to display the results of the current day’s run. (Determinations are 

listed chronologically. It will be at the end.)

3. The pH value is in the RS01 column

4. Note: Click on the Workplace icon on the left side of the Tiamo screen to return to the 

Series list.

To create Series List manually:

1. Double-click in a box on the first available line of Series (has a * on the method box) to 

bring up editing window

2. After entering the information, click on “Apply”, then use the arrows at the bottom to move 

to the next line

3. Set up Series as follows:

a. Begin Series by initiating the calibration with pH 4 & 7 buffers (first line):

i. Method: pH Cal (Select from dropdown list)

ii. Sample position: 1

iii. Station ID: Calibration

iv. Sample size: 100

v. Sample size unit: mL

b. First sample of Series is the pH 4.60 QCS sample:

i. Method: pH and ANC 2 (Select from dropdown list)

ii. Sample position: 3

iii. Station ID: PH 4.60 QCS

iv. Activity ID: QC

v. Sample size: 100

vi. Sample size unit: mL
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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c. Second sample of Series is a QC sample (DIW):

i. Method: pH and ANC 2 (Select from dropdown list)

ii. Sample position: 4

iii. Station ID: DIW

iv. Activity ID: QC

v. Sample size: 100

vi. Sample size unit: mL

d. Begin analyzing samples with the following format:

i. Method: pH and ANC 2 (Select from dropdown list)

ii. Sample position: Enter Sample Changer position for the sample

iii. Station ID: 4-character Station ID for the sample

iv. Date: Date sample was collected

v. Time: Time sample was collected

vi. Activity ID: Only use this field if the sample has a replicate. Use 1 and 2 to 

indicate which replicate it is.

vii. Organization: Organization sample is assigned to (Usually either SHEN for 

samples collected in Shenandoah National Park or UVAVTSSS for samples 

collected for the VTSSS project. Leave blank if unsure.)

e. Do not use Sample Changer positions 13 and 14 for samples

f. Include a pH 4.60 QCS sample approximately every 8 samples

g. Always end Series with a pH 4.60 QCS sample

Method

Sample 

position Station ID Date Time Activity ID Organization ANC ID

pH 

ID

Sample 

size

Sample 

size unit

pH Cal 1 Calibration 100 mL

pH and 

ANC 2

3 PH 4.60 QCS QC 100 mL

pH and 

ANC 2

4 DIW QC 100 mL

pH and 

ANC 2

5 PINE 3/23/2009 15:00 SHEN 100 mL

pH and 

ANC 2

6 NFDR 3/23/2009 16:20 SHEN 100 mL

pH and 

ANC 2

7 STAN 3/24/2009 9:15 1 SHEN 100 mL

pH and 

ANC 2

8 STAN 3/24/2009 9:15 2 SHEN 100 mL

pH and 

ANC 2

9 VT36 4/6/2009 11:15 UVAVTSSS 100 mL

pH and 

ANC 2

10 VT51 4/6/2009 12:10 UVAVTSSS 100 mL

pH and 

ANC 2

11 VT 62 4/7/2009 13:40 UVAVTSSS 100 mL

pH and 

ANC 2

12 PH 4.60 QCS QC 100 mL
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G-2.2 VWR Digital Conductivity Meter 

This document describes the current laboratory procedure for measuring specific conductance 

using the VWR Digital Conductivity Meter. The analytical method is based on the method

published in APHA (2005). The meter automatically corrects the reading to 25 °C.

Section I: Standards

Use purchased NIST-traceable conductivity standards to calibrate the conductivity meter 

following manufacturer’s instructions. Choose standards close to the expected range of samples 

to be analyzed.

Section II: Operation

Analyzing Samples:

1. Turn on conductivity meter to micromho setting

2. Rinse conductivity probe thoroughly with DIW

3. Rinse conical sample beaker 10 times with DIW

4. Fill conical sample beaker with sample

5. Place conductivity probe in sample, gently swirl and then remove probe

6. Discard sample, and then fill conical beaker with fresh sample

7. Place conductivity probe in sample, gently swirl and make sure probe is not touching the 

bottom or sides of beaker

8. Wait for reading to stabilize and then record reading

NOTE: Always measure and record conductivity of DIW at beginning of each session.

Shutdown Procedure:

1. Turn off conductivity meter

2. Rinse conductivity probe thoroughly with DIW

3. Rinse conical sample beaker 10 times with DIW

4. Place conductivity probe in sleeve with VWR Redi-StorTM Conductivity Probe Storage 

Solution

Section III: Quality Control

Calibrate conductivity meter every 6 months with NIST-traceable conductivity standards.

Measure and record the conductivity of DIW as a blank before beginning sample analysis.

G-2.3 Dionex ICS-3000 Ion Chromatography System

This document describes the current laboratory procedure for measuring anions and cations using 

the Dionex ICS-3000 dual system. Current analytes measured by ion chromatography include:

• Anions – chloride, nitrate, sulfate (Phosphate is included for MIDN project samples.)

• Cations – ammonium, calcium, magnesium, potassium, sodium

Analytical methods for anions are based on EPA Method 300.1: Determination of Inorganic 

Anions in Drinking Water by Ion Chromatography (U.S. EPA 1997). Analytical methods for 

cations are based on ASTM Method D 6919 -03: Determination of Dissolved Alkali and Alkaline 
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

289

Earth Cations and Ammonium in Water and Wastewater by Ion Chromatography (ASTM 

International 2003).

Section I: Standards

All glassware should be Class A volumetric laboratory glassware which has been initially cleaned 

by acid washing and afterwards cleaned by multiple rinsings with DI (deionized) water.

Preparation of Anion Calibration Standards:

Anion Calibration Standards Working Stock:

Prepare as follows from prepared or purchased 1000 mg/L stock solutions.

(Throughout this procedure, use only NIST-traceable stock solutions when using purchased stock 

solutions.)

Using a volumetric pipette, add the following to a 500 mL volumetric flask partially filled with 

DIW. Dilute to the line with DIW and mix well.

10 mL of 1000 mg/L chloride stock

15 mL of 1000 mg/L nitrate stock

40 mL of 1000 mg/L sulfate stock

25 mL of 100 mg/L phosphate stock - prepared by diluting 1000 mg/L phosphate stock by a 

factor of 10

The final concentration of the Anion Calibration Standards Working Stock is:

20 mg/L chloride + 30 mg/L nitrate + 80 mg/L sulfate + 5 mg/L phosphate

Anion Calibration Standards:

Dilute the indicated volume of Anion Calibration Standards Working Stock to 500 mL in a 

volumetric flask and mix well to yield standards of the following concentrations in mg/L:

Standard Volume of Stock Cl- NO3

- SO42- PO43-

1 50 mL 2.0 3.0 8.0 0.5

2 40 mL 1.6 2.4 6.4 0.4

3 30 mL 1.2 1.8 4.8 0.3

4 20 mL 0.8 1.2 3.2 0.2

5 10 mL 0.4 0.6 1.6 0.1

6 5 mL 0.2 0.3 0.8 0.05

7 2.5 mL 0.1 0.15 0.4 0.025

8 0 mL 0.0 0.0 0.0 0.0

Preserve all working stocks and standards with chloroform and store refrigerated in prepared 

polyethylene bottles.
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Preparation of Anion Quality Control Standards:

The Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance 

Project Plan of this SOP gives the following Method Detection Limit Objectives:

Chloride: 0.03 mg/L

Nitrate: 0.03 mg/L

Sulfate: 0.05 mg/L

Anion Quality Control Standards Working Stock:

Prepare as follows from purchased or prepared 1000 mg/L stock solutions.

Using a volumetric pipette, add the following to a 1000 mL volumetric flask partially filled with 

DIW. Dilute to the line with DIW and mix well.

15 mL of 1000 mg/L chloride stock

15 mL of 1000 mg/L nitrate stock

25 mL of 1000 mg/L sulfate stock

20 mL of 100 mg/L phosphate stock - prepared by diluting 1000 mg/L phosphate stock by a 

factor of 10

The final concentration of the Anion Quality Control Standard Working Stock is:

15 mg/L chloride + 15 mg/L nitrate + 25 mg/L sulfate + 2.0 mg/L phosphate

Anion MDL (method detection limit) Standard Solution:

Pipette 5 mL of the Anion Quality Control Standards Working Stock into a 1000 mL volumetric 

flask partially filled with DIW. Dilute to the line with DIW and mix well to provide a solution of 

the following concentrations:

0.075 mg/L chloride

0.075 mg/L nitrate

0.125 mg/L sulfate

0.010 mg/L phosphate

Anion Detection Limit Quality Control Check Sample:

Pipette 5 mL of the Anion Quality Control Standards Working Stock into a 500 mL volumetric 

flask partially filled with DIW. Dilute to the line with DIW and mix well to provide a solution of 

the following concentrations:

0.15 mg/L chloride

0.15 mg/L nitrate

0.25 mg/L sulfate

0.02 mg/L phosphate
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291

Anion Calibration Quality Control Check Sample:

Pipette 30 mL of the Anion Quality Control Standards Working Stock into a 500 mL volumetric 

flask partially filled with DIW. Dilute to the line with DIW and mix well to provide a solution of 

the following concentrations:

0.90 mg/L chloride

0.90 mg/L nitrate

1.50 mg/L sulfate

0.12 mg/L phosphate

Preparation of Cation Calibration Standards:

Cation Calibration Standards Working Stock:

Prepare as follows from purchased or prepared 1000 mg/L stock solutions.

Using a volumetric pipette, add the following to a 1000 mL volumetric flask partially filled with 

DIW. Dilute to the line with DIW and mix well.

5 mL of 1000 mg/L ammonium stock

50 mL of 1000 mg/L calcium stock

25 mL of 1000 mg/L magnesium stock

25 mL of 1000 mg/L potassium stock

30 mL of 1000 mg/L sodium stock

The final concentration of the Cation Working Stock is:

5 mg/L ammonium + 50 mg/L calcium + 25 mg/L magnesium + 25 mg/L potassium + 30 mg/L 

sodium

Cation Calibration Standards:

Dilute the indicated volume of Cation Calibration Standards Working Stock to 500 mL in a 

volumetric flask and mix well to yield standards of the following final concentrations in mg/L:

Standard Volume of Stock NH4

+ Ca2+ Mg2+ K+ Na+

1 50 mL 0.5 5.0 2.5 2.5 3.0

2 40 mL 0.4 4.0 2.0 2.0 2.4

3 30 mL 0.3 3.0 1.5 1.5 1.8

4 20 mL 0.2 2.0 1.0 1.0 1.2

5 10 mL 0.1 1.0 0.5 0.5 0.6

6 5 mL 0.05 0.5 0.25 0.25 0.3

7 2.5 mL 0.025 0.25 0.125 0.125 0.15

8 0 mL 0.0 0.0 0.0 0.0 0.0

Preserve all working stocks and standards with chloroform and store refrigerated in prepared 

polyethylene bottles.
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Preparation of Cation Quality Control Standards:

The Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance 

Project Plan of this SOP gives the following Method Detection Limit Objectives:

Ammonium: 0.02 mg/L

Calcium: 0.02 mg/L

Magnesium: 0.01 mg/L

Potassium: 0.04 mg/L

Sodium: 0.02 mg/L

Cation Detection Limit Quality Control Standards Working Stock:

Prepare as follows from purchased or prepared 1000 mg/L stock solutions.

Using a volumetric pipette, add the following to a 1000 mL volumetric flask partially filled with 

DIW. Dilute to the line with DIW and mix well.

10 mL of 1000 mg/L ammonium stock

10 mL of 1000 mg/L calcium stock

5 mL of 1000 mg/L magnesium stock

20 mL of 1000 mg/L potassium stock

10 mL of 1000 mg/L sodium stock

The final concentration of the Cation Quality Control Standards Working Stock is:

10 mg/L ammonium + 10 mg/L calcium + 5 mg/L magnesium + 20 mg/L potassium + 10 mg/L 

sodium

Cation MDL (method detection limit) Standard Solution:

Pipette 5 mL of the Cation Quality Control Standards Working Stock into a 1000 mL volumetric 

flask partially filled with DIW. Dilute to the line with DIW and mix well to provide a solution of 

the following concentrations:

0.05 mg/L ammonium

0.05 mg/L calcium

0.025 mg/L magnesium

0.10 mg/L potassium

0.05 mg/L sodium

Cation Detection Limit Quality Control Check Sample:

Pipette 5 mL of the Cation Quality Control Standards Working Stock into a 500 mL volumetric 

flask partially filled with DIW. Dilute to the line with DIW and mix well to provide a solution of 

the following concentrations:

0.1 mg/L ammonium

0.1 mg/L calcium

0.05 mg/L magnesium
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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0.2 mg/L potassium

0.1 mg/L sodium

Cation Calibration Quality Control Check Sample Working Stock:

Prepare as follows from purchased or prepared 1000 mg/L stock solutions.

Using a volumetric pipette, add the following to a 1000 mL volumetric flask partially filled with 

DIW. Dilute to the line with DIW and mix well.

10 mL of 1000 mg/L ammonium stock

20 mL of 1000 mg/L calcium stock

10 mL of 1000 mg/L magnesium stock

40 mL of 1000 mg/L potassium stock

20 mL of 1000 mg/L sodium stock

The final concentration of the Cation Quality Control Standard Working Stock is:

10 mg/L ammonium + 20 mg/L calcium + 10 mg/L magnesium + 40 mg/L potassium + 

20 mg/L sodium

Cation Calibration Quality Control Check Sample:

Pipette 30 mL of the Cation Quality Control Check Sample Working Stock into a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well to provide a 

solution of the following concentrations:

0.3 mg/L ammonium

0.6 mg/L calcium

0.3 mg/L magnesium

1.2 mg/L potassium

0.6 mg/L sodium

Section II: Operation

Start-up Procedure:

1. Rinse and fill eluent reservoirs, autosampler reservoir and external water pump reservoir 

with fresh DIW. (If system is already on, just fill all reservoirs with DIW.)

2. Log in to IC computer

3. Open Chromeleon software:

a. Toshiba-User_1 is System 1: Anions

b. Toshiba-User_2 is System 2: Cations

4. If the Tabset panel “Anions and Cations Panels” does not open, choose it under the 

Window menu

5. Turn on external water pump by choosing the Detector Compartment tab for the Anion 

system and changing the DC_ACRelay_1 to closed
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6. If both the anion and cation systems are being used, perform steps 7 - 12 for both systems

7. Switch on the pump motor using the Panel (Home tab)

8. Loosen the purge valve screw on the pump one-half turn

9. Switch the pump to Prime using the Panel. The pump will rapidly deliver DIW from the 

reservoir out the waste line. Watch for bubbles coming through the tubing at the pump and 

once all bubbles are gone, switch Prime off and tighten the purge valve.

10. Check that the pressure builds and then levels off around 2000 psi

11. Switch on all the other modules on the Panel:

a. Eluent generator

b. Trap column

c. Suppressor

d. Cell heater

e. Column heater

12. Sometimes the suppressor will not turn on using the Panel. If this happens:

a. Select the Detector Compartment tab

b. Click on Calculate Current

c. Set the cell temperature to 30.0 °C and click OK

d. Click on Calculate Current again

e. Select the suppressor type (either ASRS-4mm for anions or CSRS-4mm for cations) 

and enter the desired eluent concentration

f. If this doesn’t work either, run a DIW sample using a Sequence, and it will set 

everything correctly

13. Prime the autosampler syringe using the Prime button under the Autosampler tab. Then 

flush the syringe using the Flush button.

14. The system will now have to run for several hours until it is stable. For anions, the 

background conductivity should be about 1.0 uS. For cations, the background conductivity 

should be about 0.1 uS. To check the background conductivity, select the Conductivity 

Detector tab on the Panel and see what the Total Signal reading is. After the background 

conductivity reaches these values and is no longer rising or falling, the system is ready to 

use for analysis.

Analyzing Samples:

The following steps have to be performed for both Anions and Cations separately.

1. Open the Browser and create a new Sequence by opening a previous Sequence that used the 

appropriate program and method and saving it under a new name. (Save it under a new 

name; otherwise all the data from the previous sequence will be lost.)

2. Copy the names and primary keys for the samples and QC samples from the spreadsheet 

file generated by the SWAS/VTSSS LIMS (Laboratory Information Management System) 

and save the Sequence. Alternatively, enter the sample names manually.

3. All autosampler vials should be rinsed several times with DIW and preferably filled with 

DIW and stored overnight before use

4. For each sample:
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295

a. Pre-rinse autosampler vial with sample by pouring 2-3 mLs of sample into 

autosampler vial, cap it, shake and then discard sample. Repeat 2-3 times if sample 

volume permits.

b. Fill one-half to two-thirds full with sample and cap vial

c. Load the sample vials into the rack according to the Sequence

5. Start the Sequence with the following steps:

a. Under the Batch menu, choose Start

b. Add the Sequence to be started

c. Click on Ready Check, and if there are any other warnings other than the amount of 

disk space and eluent needed, these will have to be fixed before the Sequence can be 

started

d. Click Start

6. Be sure there is enough DIW in all reservoirs before leaving unattended for long periods of 

time.

Shutdown Procedure:

1. Turn external water pump off by choosing the Detector Compartment tab for the Anion 

system and changing the DC_ACRelay_1 to open. Wait 3-5 minutes for the bubbles to 

clear out of the suppressors before proceeding with step 2.

2. Switch off all the other modules on the Panel for all systems being used:

a. Eluent generator

b. Trap column

c. Suppressor

d. Cell heater

e. Column heater

Section III: Quality Control

Sequence Set-up:

Described on pg. 70 in Figure 6-2 of Paulsen, 1997.

1. Calibration standards

2. Laboratory blank (DIW)

3. Detection limit quality control check sample

4. Calibration quality control check sample

5. Samples (insert quality control check samples at regular intervals)

6. Calibration quality control check sample

7. Calibration standards

Note: Analyze the MDL Standard Solution at least once during each sequence
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Performance Criteria:

Laboratory blank: Value obtained should be less than the specified MDL objective given in the 

Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance Project 

Plan of this SOP

Detection limit quality control sample: Value obtained should be within specified limits: True 

value + MDL objective given in the Measurement Data Quality Objectives Table in the Project 

Laboratory Quality Assurance Project Plan of this SOP

Calibration quality control check sample: Value obtained should be within specified limits: True 

value + precision objective given in the Measurement Data Quality Objectives Table in the 

Project Laboratory Quality Assurance Project Plan of this SOP

G-2.4 Technicon AutoAnalyzer II: Silica – SiO2

This document describes the current laboratory procedure for measuring silica using the 

Technicon AutoAnalyzer II. Analytical methods are based on the methods published in American 

Public Health Association (2005).

Summary: Silica in the sample reacts with molybdate ion in acidic solution to form a colored 

complex. The absorbance of the resulting solution is measured using a spectrophotometer.

Section I: Standards

All glassware should be Class A volumetric laboratory glassware which has been initially cleaned 

by acid washing and afterwards cleaned by multiple rinsings with DI (deionized) water.

Preparation of Silica Calibration Standards:

Silica Calibration Standards Working Stock: 214 mg/L SiO2

Prepare as follows from prepared or purchased 1000 mg/L stock solutions.

(Throughout this procedure, use only NIST-traceable stock solutions when using purchased stock 

solutions.)

Using a volumetric pipette, add 50 mL of 1000 mg/L Si stock solution to a 500 mL volumetric 

flask partially filled with DIW. Dilute to the line with DIW and mix well.

Silica Calibration Standards:

Dilute the indicated volume of Silica Calibration Standards Working Stock to 1000 mL in a 

volumetric flask and mix well to yield standards of the following concentrations in mg/L:

Std. # Volume of stock Concentration (mg/L SiO2)

1 50 mL 10.7

2 40 mL 8.56

3 30 mL 6.42

4 20 mL 4.28

5 10 mL 2.14

6 5 mL 1.07

7 2.5 mL 0.535

8 0 mL 0.00
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Preparation of Silica Quality Control Standards:

The Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance 

Project Plan of this SOP gives the following Method Detection Limit Objective: 0.05 mg/L SiO2

Silica Quality Control Standards Working Stock: 12.84 mg/L SiO2

Prepare as follows from prepared or purchased 1000 mg/L stock solutions.

Two steps:

1. Using a volumetric pipette, add 25 mL of 1000 mg/L Si stock solution to a 250 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well. The 

final concentration of this solution is 214 mg/L SiO2.

2. Using a volumetric pipette, add 30 mL of the 214 mg/L SiO2 solution to a 500 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well. The 

final concentration of this solution is 12.84 mg/L SiO2.

Silica MDL (method detection limit) Standard Solution:

0.1284 mg/L SiO2:

Pipette 10 mL of the 12.84 mg/L Silica Quality Control Standards Working Stock into a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.

Silica Detection Limit Quality Control Check Sample: 

0.2568 mg/L SiO2:

Pipette 10 mL of the 12.84 mg/L Silica Quality Control Standards Working Stock into a 500 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.

Silica Calibration Quality Control Check Sample:

2.568 mg/L SiO2:

Pipette 100 mL of the 12.84 mg/L Silica Quality Control Standards Working Stock into a 500 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.

Section II: Operation

Reagents:

• Ammonium Molybdate reagent: Stable for 2 months refrigerated.

1. Add 40 mL of 2.5 N sulfuric acid (H2SO4) to a cleaned 1000 mL volumetric flask 

partially filled with DIW and mix

2. Add 10.0 g ammonium molybdate ((NH4)6Mo7O24.4H2O) and mix until dissolved

3. Fill to the line with DIW and mix well

4. Filter or remake if cloudy

5. Add 1-2 mL of SLS solution* as a wetting agent

• Oxalic Acid reagent: Prepare daily.

1. Add 17.5 g oxalic acid dihydrate (H2C2O4.2H2O) to a cleaned 250 mL volumetric 

flask partially filled with DIW and mix until dissolved
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2. Fill to the line with DIW and mix well

• Ascorbic Acid reagent: Prepare daily.

1. Add 12.5 mL acetone (CH3COCH3) to a cleaned 250 mL volumetric flask partially 

filled with DIW and mix

2. Add 4.4 g ascorbic acid (C6H8O6) and mix until dissolved

3. Fill to the line with DIW and mix well

4. Add 1-2 mL of SLS solution* as a wetting agent

* To prepare SLS solution: Add 6-8 g of sodium lauryl sulfate to 100 mL deionzed water, mix 

and then add 5 drops concentrated sulfuric acid.

Colorimeter configuration for silica:

Flow cell: 15 mm

Mode: I

Wavelength filter: 660 nm

Sampling rate: 30/hour

Sample:Rinse ratio: 2:1

Start-up Procedure:

1. Rinse and fill reservoirs for autosampler rinse tubing and reagent tubing with fresh DIW

2. Flush autosampler rinse reservoir with DIW using a squirt bottle

3. Mount auto-analyzer flow-rated pump tubing on pump, place platen on pump and turn 

motor on

4. Allow DIW to pump to flush system for at least 10 minutes

5. Place labeled reagent lines into appropriate reagent container. (All reagents should be room 

temperature.)

6. From this point, collect waste in hazardous waste container

7. Turn on strip chart recorder and set to 100 mV

8. Turn on colorimeter

9. Turn the knob on the colorimeter to ZERO and adjust the pen on the strip chart recorder to 

read zero units using the zero knob on the strip chart recorder

10. Turn the knob on the colorimeter to FULL SCALE and adjust the pen on the strip chart 

recorder to read 100 units using the var knob on the strip chart recorder

11. Turn the knob on the colorimeter to NORMAL

12. Turn the baseline adjusting knob on the colorimeter so that it is mid-range by turning knob 

as far as it will go one way and then turn back 5 full turns

13. After reagents have pumped through the colorimeter for about 10 minutes:

a. Turn the STD CAL knob on the colorimeter to the previous run’s setting, if known
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b. Turn the sample light source silver knob so that it is fully open (completely 

clockwise)

c. Adjust the reference light source silver knob until the pen on the strip chart recorder 

is at about 5 units

14. Install pen on strip chart recorder and start paper at 10 mm/min

15. Place a tube of highest concentration standard in the auto sampler and run several times (5-

10):

a. When first peak comes out, adjust STD CAL knob on colorimeter so that the top of 

the peak is at about 95 units

b. After adjustment, check for reproducibility. Successive peaks should be within 2 

units

Analyzing Samples:

1. Create a Sample list using the spreadsheet file generated by the SWAS/VTSSS LIMS 

(Laboratory Information Management System). Alternatively record the sample names 

manually.

2. For each sample:

a. Rinse autosampler tube with DIW

b. Pre-rinse each autosampler tube with sample by filling half-full and discarding.

Repeat if sample volume permits.

c. Fill one-half to two-thirds with sample

d. Load the sample vials into the rack according to the Sample list

3. Throughout the run, check reservoir for autosampler rinse tubing and refill with fresh DIW

as needed

Shutdown Procedure:

1. After the autosampler has sampled the last tube, turn off autosampler

2. After the last peak has been recorded on the stripchart recorder, turn off stripchart recorder, 

and remove and cap pen

3. Turn off colorimeter

4. Remove reagent lines from reagent containers and place in reservoir for reagent tubing 

filled with fresh DIW

5. Pump DIW through all tubing lines for about 15 minutes until reagents are out of lines

6. Turn pump motor off, remove platen from pump, and release flow-rated pump tubing

7. Cap hazardous waste container and dispose of properly

Section III: Quality Control

Sequence Set-up:

Described on pg. 70 in Figure 6-2 of Paulsen, 1997

1. Calibration standards

2. Laboratory blank (DIW)
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3. Detection limit quality control check sample

4. Calibration quality control check sample

5. Samples (insert quality control check samples at regular intervals)

6. Calibration quality control check sample

7. Calibration standards

Note: Analyze the MDL Standard Solution at least once during each sequence

Performance Criteria:

Laboratory blank: Value obtained should be less than the specified MDL objective given in the 

Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance Project 

Plan of this SOP

Detection limit quality control sample: Value obtained should be within specified limits: True 

value + MDL objective given in the Measurement Data Quality Objectives Table in the Project 

Laboratory Quality Assurance Project Plan of this SOP

Calibration quality control check sample: Value obtained should be within specified limits: True 

value + precision objective given in the Measurement Data Quality Objectives Table in the 

Project Laboratory Quality Assurance Project Plan of this SOP

G-2.5 Technicon AutoAnalyzer II: Total and Organic Monomeric Aluminum

This document describes the current laboratory procedure for measuring total and organic 

monomeric aluminum using the Technicon AutoAnalyzer II. Analytical methods are based on the 

methods published in McAvoy et al. (1992).

Summary: Monomeric aluminum in the sample reacts with pyrocatechol violet to form a colored 

complex. The absorbance of the resulting solution is measured using a spectrophotometer.

Section I: Standards

All glassware should be Class A volumetric laboratory glassware which has been initially cleaned 

by acid washing and afterwards cleaned by multiple rinsings with DI (deionized) water.

Preparation of Aluminum Calibration Standards:

Aluminum Calibration Standards Working Stock: 0.50 mg/L Al

Prepare as follows from prepared or purchased 1000 mg/L stock solutions.

(Throughout this procedure, use only NIST-traceable stock solutions when using purchased stock 

solutions.)

Two steps:

1. Using a volumetric pipette, add 25 mL of 1000 mg/L Al stock solution to a 500 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well. The 

final concentration of this solution is 50.0 mg/L Al.

2. Using a volumetric pipette, add 10 mL of the 50.0 mg/L Al solution to a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

301

Aluminum Calibration Standards:

Dilute the indicated volume of Aluminum Calibration Standards Working Stock to 250 mL in a 

volumetric flask and mix well to yield standards of the following concentrations in ug/L:

Std. # Volume of stock Concentration (mg/L Al)

1 50 mL 100

2 40 mL 80

3 30 mL 60

4 20 mL 40

5 10 mL 20

6 5 mL 10

7 2.5 mL 5

8 0 mL 0

Preparation of Aluminum Quality Control Standards:

The Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance 

Project Plan of this SOP gives the following Method Detection Limit Objective: 10 ug/L Al

Aluminum Quality Control Standards Working Stock: 1.0 mg/L Al

Prepare as follows from purchased or prepared 1000 mg/L stock solutions.

Two steps:

1. Using a volumetric pipette, add 25 mL of 1000 mg/L Al stock solution to a 500 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well. The 

final concentration of this solution is 50.0 mg/L.

2. Using a volumetric pipette, add 20 mL of the 50.0 mg/L Al solution to a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well. The 

final concentration of this solution is 1.0 mg/L Al.

Aluminum MDL (method detection limit) Standard Solution:

20 ug/L Al:

Pipette 20 mL of the 1.0 mg/L Aluminum Quality Control Standards +Working Stock into a 1000 

mL volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.

Aluminum Detection Limit Quality Control Check Sample: 

40 ug/L Al:

Pipette 20 mL of the 1.0 mg/L Aluminum Quality Control Standards Working Stock into a 500 

mL volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.

Aluminum Calibration Quality Control Check Sample:

50 ug/L Al:

Pipette 25 mL of the 1.0 mg/L Aluminum Quality Control Standards Working Stock into a 500 

mL volumetric flask partially filled with DIW. Dilute to the line with DIW and mix well.
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Section II: Operation

Reagents:

• Pyrocatechol Violet (PCV) Reagent – Dilute PCV stock 1:10 with DIW:

1. With a graduated cylinder, add 100 mL of stock to a pre-rinsed 1000 mL volumetric 

flask

2. Dilute to the line with DIW and mix well

PCV stock: Add 1.875 g of pyrocatechol violet (C19H14O7S) to a 1000 mL volumetric 

flask partially filled with DIW and mix. Dilute to the line with DIW and mix well.

Store refrigerated in an amber polyethylene bottle.

• Hydroxylamine hydrochloride-Phenanthroline Reagent – Dilute Hydroxylamine 

hydrochloride-Phenanthroline stock 1:10 with DIW:

1. With a graduated cylinder, add 100 mL of stock to a pre-rinsed 1000 mL volumetric 

flask

2. Dilute to the line with DIW and mix well

• Hydroxylamine hydrochloride-Phenanthroline stock: Add 300 g of hydroxylamine 

hydrochloride (NH2OH . HCl) to a 1000 mL volumetric flask partially filled with DIW and 

mix. Add 3 g of 1,10-phenanthroline (C12H8N2) and mix. Dilute to the line with DIW and mix 

well. Store refrigerated in an amber polyethylene bottle.

• Hexamethylene Tetramine Buffer Reagent:

1. Add 200 g of hexamethylene tetramine ((CH2)6N4) to a 1000 mL volumetric flask 

partially filled with DIW and mix

2. Dilute to the line with DIW and mix well

3. Add 1-2 mLs of Triton-X as a surfactant and mix well

Colorimeter configuration for Aluminum:

Flow cell: 50 mm

Mode: I

Wavelength filter: 590 nm

Sampling rate: 10/hour

Sample:Rinse Ratio: 1:3

Start-up Procedure:

1. Rinse and fill reservoirs for autosampler rinse tubing and reagent tubing with fresh DIW

2. Flush autosampler rinse reservoir with DIW using a squirt bottle

3. Mount auto-analyzer flow-rated pump tubing on pump, place platen on pump and turn 

motor on

4. Allow DIW to pump to flush system for at least 10 minutes
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

303

5. Place labeled reagent lines into appropriate reagent container. (All reagents should be room 

temperature.)

6. From this point, collect waste in hazardous waste container.

7. Turn on strip chart recorder and set to 100 mV

8. Turn on colorimeter

9. Turn the knob on the colorimeter to ZERO and adjust the pen on the strip chart recorder to 

read zero units using the zero knob on the strip chart recorder

10. Turn the knob on the colorimeter to FULL SCALE and adjust the pen on the strip chart 

recorder to read 100 units using the var knob on the strip chart recorder

11. Turn the knob on the colorimeter to NORMAL

12. Turn the baseline adjusting knob on the colorimeter so that it is mid-range by turning knob 

as far as it will go one way and then turn back 5 full turns

13. After reagents have pumped through the colorimeter for about 10 minutes:

a. Turn the STD CAL knob on the colorimeter to the previous run’s setting, if known

b. Turn the sample light source silver knob so that it is fully open (completely 

clockwise)

c. Adjust the reference light source silver knob until the pen on the strip chart recorder 

is at about 5 units

14. Install pen on strip chart recorder and start paper at 10 mm/min

15. Place a tube of highest concentration standard in the auto sampler and run several times (5-

10):

a. When first peak comes out, adjust STD CAL knob on colorimeter so that the top of 

the peak is at about 95 units

b. After adjustment, check for reproducibility. Successive peaks should be within 2 

units.

Analyzing Samples:

1. Create a Sample list using the spreadsheet file generated by the SWAS/VTSSS LIMS.

Alternatively record the sample names manually.

2. For each sample:

a. Rinse autosampler tube with DIW

b. Pre-rinse each autosampler tube with sample by filling half-full and discarding.

Repeat if sample volume permits.

c. Fill one-half to two-thirds with sample

d. Load the sample vials into the rack according to the Sample list

3. Throughout the run, check reservoir for auto-sampler rinse tubing and refill with fresh DIW

as needed
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Fractionation Procedure:

To measure the organic fraction of the total monomeric aluminum, the inorganic monomeric 

aluminum must be removed by passing the sample through a cation exchange column. The 

resulting fraction is analyzed to quantify the organic monomeric aluminum. The inorganic 

monomeric aluminum fraction may then be calculated by taking the difference between the total 

monomeric aluminum concentration and the organic monomeric aluminum fraction.

1. Prepare cation exchange resin:

a. Tare a large (1-2 liter) plastic beaker on a lab balance

b. Add the desired amount of Amberlite IR-120+ sodium form ion exchange resin and 

record the mass

c. Calculate 1% of the mass of the sodium form. Add this amount of Amberlite IR-120+ 

hydrogen form ion exchange resin.

d. Add DIW and mix with a stirring rod

e. The composition of the cation exchange resin should be such that the pH of the 

sample does not change significantly after passing through the column. Several 

different compositions of resin may be necessary if the pH range of the samples is 

large (pH 4-7). The lower the sample pH value, the lower the pH of the resin should 

be. Test the resin by fractioning samples of various pH values and observing the pH 

change of the sample. Adjust the composition of the resin by adding more hydrogen 

form (to lower the pH) or sodium form (to raise the pH) of Amberlite IR-120+. In 

this way, prepare various resins appropriate for the samples.

2. Fractionate samples:

a. Rinse 250 mL separatory funnel, stopper with tube, flow regulator and various pieces 

of connecting tubing with DIW at least 3 times

b. Pre-rinse funnel with a few milliliters of sample and discard

c. Close funnel stopcock and fill funnel with 50 – 75 mL of sample (or less if sample 

volume is low)

d. Insert stopper with tube into top of funnel

e. Prepare cation exchange column:

i. Replace poly-fiber in end of column tubing daily

ii. Insert connector into end of column tubing with the poly-fiber in it

iii. Place the other end of the column tubing into a polyethylene beaker filled 

with cation exchange resin (Add DIW as needed to make the resin the proper 

consistency.)

iv. Using a pipette bulb, pull resin into the column tubing

v. Remove pipette bulb, and hold thumb over end of tubing

vi. Remove other end of tubing from resin and quickly hold both ends of tubing 

up so that resin does not flow out
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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vii. Check to see that the column is full of well-packed resin with no air pockets.

If not, remake column.

viii. Use a DIW squirt bottle to bring level of resin to end of connector to create a 

4 cm column

ix. Do not allow water to flow out of column at any time to keep resin well 

packed. If this happens, make a new column.

f. Close the pinch clamp at the end of the tubing connected to the flow regulator

g. Keeping the column in a U-shape so that resin or water does not flow out, connect the 

end of the column with the poly-fiber in it to the flow regulator

h. Connect the other end of the column to the tubing at the bottom of the funnel

i. Confirm that the column is wetted, well-packed and without air pockets

j. Open funnel stopcock

k. Squeeze and tap tubing at end of funnel to remove air and ensure free flow

l. Open pinch clamp and allow at least 20 mL of sample to come through to flush 

column and tubing

m. Check to see that bubbles are coming out of the funnel stopper tube at a consistent 

rate. If not, be sure stopper is tight at top of funnel.

n. The flow rate out of the column should be 4 mL +/- 0.25 mL per 15 seconds. Check 

with graduated cylinder and stopwatch. If it is too slow or too fast, adjust rate by 

opening or closing the flow regulator as needed.

o. Collect sample in polyethylene beaker. Analyze as soon as possible. Store by 

refrigeration only if absolutely necessary.

Shutdown Procedure:

1. After the autosampler has sampled the last tube, turn off autosampler

2. After the last peak has been recorded on the stripchart recorder, turn off stripchart recorder, 

and remove and cap pen

3. Turn off colorimeter

4. Remove reagent lines from reagent containers and place in reservoir for reagent tubing

filled with fresh DIW

5. Pump DIW through all tubing lines for about 15 minutes until reagents are out of lines

6. Turn pump motor off, remove platen from pump, and release flow-rated pump tubing

7. Cap hazardous waste container and dispose of properly
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Section III: Quality Control

Sequence Set-up:

Described on pg. 70 in Figure 6-2 of Paulsen, 1997

1. Calibration standards

2. Laboratory blank (DIW)

3. Detection limit quality control check sample

4. Calibration quality control check sample

5. Samples (insert quality control check samples at regular intervals)

6. Calibration quality control check sample

7. Calibration standards

Note: Analyze the MDL Standard Solution at least once during each sequence

Performance Criteria:

Laboratory blank: Value obtained should be less than the specified MDL objective given in the 

Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance Project 

Plan of this SOP

Detection limit quality control sample: Value obtained should be within specified limits: True 

value + MDL objective given in the Measurement Data Quality Objectives Table in the Project 

Laboratory Quality Assurance Project Plan of this SOP

Calibration quality control check sample: Value obtained should be within specified limits: True 

value + precision objective given in the Measurement Data Quality Objectives Table in the 

Project Laboratory Quality Assurance Project Plan of this SOP

G-2.6 Teledyne-Tekmar Phoenix 8000 TOC Analyzer

This document describes the current laboratory procedure for measuring DOC using the 

Teledyne-Tekmar Phoenix 8000 TOC (total organic carbon) Analyzer. Analytical methods are 

based on the methods published in EPA Method 415.3.

Summary: Inorganic carbon is removed by acidification and sparging with nitrogen gas. Organic 

carbon in the sample is oxidized to carbon dioxide by simultaneous exposure to persulfate ions 

and UV (ultraviolet) radiation. The carbon dioxide produced is then measured by an NDIR 

(nondispersive infrared) detector.

Section I: Standards

All glassware for preparing DOC standards should be Class A volumetric laboratory glassware 

which has been cleaned as follows:

1. Wash with laboratory detergent (Liquinox) in hot tap water

2. Rinse with hot tap water

3. Fill with (or soak in) 2N HCl overnight

4. Rinse 10 times with DIW

5. Use the glassware immediately (If this is not possible, store the glassware filled with DIW

and preferably use within 24 hours.)
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Preparation of DOC Calibration Standards:

DOC Calibration Standards Working Stock: 50 mg/L DOC

(Throughout this procedure, use only NIST-traceable stock solutions when using purchased stock 

solutions.)

Use a purchased 50 mg/L TOC/DOC standard or prepare as follows from a purchased 1000 mg/L 

TOC/DOC standard:

Using a volumetric pipette, add 25 mL of 1000 mg/L TOC/DOC standard to a 500 mL volumetric 

flask partially filled with DIW. Acidify with 2.5 mL H3PO4 to bring to pH < 2. Dilute to the line 

with DIW.

DOC Calibration Standards:

All standards must be made at the same time with the same source of DIW.

Prepare standards using volumetric glassware according to the following table:

Volume of stock Concentration (mg/L) Total Volume Vol. H3PO4 added

50 mL 10.0 250 mL 1.25 mL

50 mL 5.0 500 mL 2.5 mL

25 mL 2.5 500 mL 2.5 mL

10 mL 1.0 500 mL 2.5 mL

5 mL 0.5 500 mL 2.5 mL

5 mL 0.25 1000 mL 5 mL

0 mL 0.0 500 mL 2.5 mL

Add H3PO4 to standards after bringing to final volume.

Store at room temperature in volumetric flasks away from light. May be stored for 30 days.

Preparation of DOC Quality Control Standards:

The Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance 

Project Plan of this SOP gives the following Method Detection Limit Objective: 0.1 mg/L DOC

DOC Quality Control Standards Working Stock: 100 mg/L DOC

Use a purchased 100 mg/L TOC/DOC standard or prepare as follows from a purchased 1000 

mg/L TOC/DOC standard:

Using a volumetric pipette, add 50 mL of 1000 mg/L TOC/DOC standard to a 500 mL volumetric 

flask partially filled with DIW. Acidify with 2.5 mL H3PO4. Dilute to the line with DIW.

DOC MDL (method detection limit) Standard Solution:

0.25 mg/L DOC:

Pipette 5 mL of the 100 mg/L DOC Quality Control Standards Working Stock into a 2000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix. Add 10 mL 

H3PO4 and mix well.

DOC Detection Limit Quality Control Check Sample: 

0.50 mg/L DOC:
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Pipette 5 mL of the 100 mg/L DOC Quality Control Standards Working Stock into a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix. Add 5 mL H3PO4

and mix well.

DOC Calibration Quality Control Low Check Sample:

1.5 mg/L DOC:

Pipette 15 mL of the 100 mg/L DOC Quality Control Standards Working Stock into a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix. Add 5 mL H3PO4

and mix well.

DOC Calibration Quality Control High Check Sample:

3.0 mg/L DOC:

Pipette 30 mL of the 100 mg/L DOC Quality Control Standards Working Stock into a 1000 mL 

volumetric flask partially filled with DIW. Dilute to the line with DIW and mix. Add 5 mL H3PO4

and mix well.

Section II: Operation

Reagents:

• 21% Phosphoric Acid Reagent: Stable for one month

1. Rinse a clean (does not have to be acid-washed) laboratory flask multiple times 

with DI (deionized) water

2. Add 188 mL of DIW using a graduated cylinder which has been rinsed well

3. Carefully add 37 mL of 85% phosphoric acid (H3PO4) and mix well

• 10% Persulfate/5% Phosphoric Acid Reagent: Stable for one week

1. Rinse a clean (does not have to be acid-washed) laboratory flask multiple times with 

DIW

2. Add 25 g of 98+% sodium persulfate (Na2S2O8)

3. Add 213 mL DIW and mix well

4. Carefully add 9 mL of 85% phosphoric acid (H3PO4) and mix well

5. Wait 12 hours before using

Note: When analyzing more than 50 samples, double the amount of this reagent.

Start-up Procedure:

Note: If power to Phoenix has been turned off, allow 2 hours for NDIR detector to warm up 

before analysis.

1. Carefully remove the gas/liquid separator from the instrument and rinse well with DIW.

Add a few drops of 21% phosphoric acid reagent, fill to level of sidearm with DIW and 

then replace. (Perform at least weekly.)

2. Unplug the UV lamp, disconnect lines and carefully remove it from the instrument. Rinse 

well with DIW and then replace. (Perform at least weekly.)
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

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3. Carefully remove the IC sparger from the instrument. Rinse well with DIW and then 

replace. (Perform at least weekly.)

4. Remove plug from bottom of mist trap to drain excess water

5. Check the copper in the halogen scrubber. If it has become totally discolored, replace 

copper and tin in the halogen scrubber

6. Check screw at bottom of syringe and hand-tighten

7. Place appropriate reagent lines in Phosphoric Acid and Persulfate reagents

8. Rinse DIW reservoir a few times and fill with DIW

9. Place waste line into waste container

10. Log in to DOC computer

11. Open TOC Talk for Phoenix 8000 (version 3.6.385.2)

12. Type in User name

13. Type in Password

14. Open valve on nitrogen tank. (Make sure that the tank level is at 500 + psi.)

15. Choose System  Instrument and click to change system status from Standby to Ready

16. On the Instrument Setup/Status screen, check that the gas flow rates (From Detector and To 

UV Reactor) are 200 cc/min +/- 10% and within 15 psi of each other. Click OK to close 

window. (It is seldom necessary to adjust the nitrogen tank regulator, but never turn the 

tank input pressure above 35 psi. Low flow rates usually indicate a leak in the system.)

17. Exit the Instrument Setup/Status screen by choosing OK

Calibrating Instrument:

1. Choose Run and select Sample Setup

2. Choose File  Open

a. Select appropriate calibration curve file or create a new one as described below

b. Change Status field to Ready for the first line and then for all samples by choosing 

Edit  Reset Status for all rows

c. An example of a file for calibration going up to 10.0 mg/L is:

Pos Sample ID Sample Type Method ID Reps Status

1001 Prime Sample Prime System 1 Ready

1001 Clean Sample Cleaning Procedure 3 Ready

1001 Clean Sample Cleaning Procedure 5 Ready

1 0.0 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready

2 0.25 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready

3 0.5 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready

4 1.0 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready

5 2.5 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready
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Pos Sample ID Sample Type Method ID Reps Status

6 5.0 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready

7 10.0 mg/L TOC Standard Drinking H2O TOC 0.1-20ppm 3 Ready

3. Alternatively, to create a new file to use to calibrate: Choose File  New

a. Make sure that the Rack Style is 40 mL vial

b. Enter an 8-character Rack ID

c. Use the down arrow to add more lines

d. When you select TOC Standard, a pop-up menu will appear to select the appropriate 

range (usually 0.1 – 20 ppm)

e. Select the appropriate standard from the list and choose Exit

4. Load the appropriate standards in the autosampler according to the Autosampler list

5. Choose Save/Use

6. Choose System  Instrument and de-select the Auto Shutdown box if necessary

7. Choose Start (note that it will take several hours for the calibration to be completed)

Confirming Calibration:

1. After the standards are finished running, the Calibration Curve screen automatically 

appears. (To open it manually, select Results  Calibration.)

2. Select the calibration curve file created in step 2 above from the list, if it is not already 

displayed

3. In the Use column, select the standards that have the current date and de-select all others

4. Choose Recalc to display the new calibration curve

5. Choose Save as New Version

6. Choose Exit

Analyzing Samples:

1. Choose Run and select Sample Setup

2. Choose File  New

3. Build the Autosampler list:

a. Make sure that the Rack Style is 40 mL vial

b. Use the arrow key to add more lines

c. First 2 samples are:

Pos Sample ID Sample Type Method ID Reps Status

1001 Clean Sample Cleaning Procedure 6 Ready

1001 System Blank Blank TC Range 2 Blank TC Range 2 Drinking H2O 5 Ready

d. Throughout the run, include DOC Quality Control Standards, which must be run as 

sample type “Calibration Verification”

e. Enter sample data with the following format:

Pos = Sample position in autosampler rack
APPENDIX G. EXAMPLE STANDARD OPERATING INSTRUCTIONS FOR LAB ANALYSIS PROTOCOLS

311

Sample ID = Sample identification (usually site, date and time)

Sample Type = Sample

Method ID = Drinking H2O TOC 0.1 -20ppm (or range being used)

Reps = 2 (may run 3 from a 40 mL vial if desired)

Status = Ready

f. The last sample should be a DOC quality control check sample run as sample type 

“Calibration Verification”

g. Always finish the run with a cleaning procedure:

Pos Sample ID Sample Type Method ID Reps Status

1001 Clean Sample Cleaning Procedure 6 Ready

4. To begin the run immediately, choose Save/Use and enter a Rack ID, usually in the 

following format: YYYYMMDD (YYYY for 4-digit year, MM for 2-digit month and DD 

for 2-digit day)

a. If the analyzer is already running, use “Save as” to save the file to use later and then 

choose Exit

b. When ready to use the saved file, choose File  Open and select saved file

5. Choose System  Instrument and select the Auto Shutdown box

6. If the Calibration has been confirmed, choose Start to begin analysis

Shutdown Procedure:

1. After the run is complete, Exit the Sample Analysis screen

2. Choose System  Instrument and click to change system status from Ready to Standby and 

then choose OK

3. Choose System  Exit

4. If the analysis was completed less than 30 minutes ago, if asked, answer “Yes” to the 

question: “Would you like to leave the gas to the Perm Dryer ON?”

5. Turn off N2 tank. Wait 30 minutes if the analysis was completed less than 30 minutes for 

the gas to dry the Perm Dryer.

6. Cap waste container and dispose of waste properly

Long versus Short-time Shutdown:

Short-time Shutdown: Leave the Phoenix 8000 in Standby mode when not in use. Standby 

mode turns off all components except the detector.

Long-time Shutdown: If the Phoenix 8000 will not be run for over a month, the auto-sampler 

and analyzer should be turned off with the power switches in the back. When turning the analyzer 

back on, allow 2 hours for the NDIR detector to stabilize before beginning analyses.
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Section III: Quality Control

Sequence Set-up:

Described on pg. 70 in Figure 6-2 of Paulsen, 1997

1. Calibration standards

2. Laboratory blank (DIW)

3. Detection limit quality control check sample

4. Calibration quality control check sample

5. Samples (insert quality control check samples at regular intervals)

6. Calibration quality control check sample

Note: Analyze the MDL Standard Solution at least once during each sequence

Performance Criteria:

Laboratory blank: Value obtained should be less than the specified MDL objective given in the 

Measurement Data Quality Objectives Table in the Project Laboratory Quality Assurance Project 

Plan of this SOP

Detection limit quality control sample: Value obtained should be within specified limits: True 

value + MDL objective given in the Measurement Data Quality Objectives Table in the Project 

Laboratory Quality Assurance Project Plan of this SOP

Calibration quality control check sample: Value obtained should be within specified limits: True 

value + precision objective given in the Measurement Data Quality Objectives Table in the 

Project Laboratory Quality Assurance Project Plan of this SOP

REFERENCES

ASTM International. 2003. Standard D 6919-03: Test Method for Determination of Dissolved 

Alkali and Alkaline Earth Cations and Ammonium in Water and Wastewater by Ion 

Chromatography. ASTM International, West Conshohocken, PA. Available at: 

<www.astm.org> (last accessed in February 2012)

U.S. Environmental Protection Agency [EPA]. 1997. Methods for the Determination of Organic 

and Inorganic Compounds in Drinking Water, Volume 1. (EPA/815-R-00-014). Office of 

Ground Water and Drinking Water, Washington, DC. 470p.

U.S. Geological Survey [USGS]. Standard Operating Procedure for Laboratory Analysis of One 

Analyte (Ammonium) in Dilute Fresh Water – Flow Injection Analysis. Unpublished 

manuscript.

Webb, R.F., R., F.A. Deviney, and S.W. Maben. University of Virginia Standard Operating 

Procedure for Laboratory Analysis: Shenandoah Watershed Study/Virginia Trout Stream 

Sensitivity Study. Unpublished manuscript.
313

APPENDIX H. 

TRAINING CHECKLISTS 

Table H-1. Checklist of materials and supplies for stream sampling site visits.

Items: 

Standard Items

Collection permits and entry permits, if required.

Site Documentation Forms (for new sites)

Clipboard

Site Documentation Reports (compiled in folders for existing sites)

Stream sampling record forms

Insulated container with ice or frozen refrigerant (packed in sealed plastic bags or other containers)

Small insulated container (with ice) for hike-in sites

Watch for recording time

Digital field camera with free memory and extra charged battery

GPS unit with extra batteries

Compass

Field thermometer (with string attached)

Pre-processed sample bottle(s) with sample label attached. Include a second bottle if sampling at that site is to be 

replicated. Put each bottle in a clean plastic zipper-lock bag.

Plastic gloves in sealed plastic bag

60 mL plastic syringes (with Luer type tip) with completed sample labels attached. Plastic container with snap-on lid 

to hold filled syringes

Syringe valves (Mininert® with Luer type adapter, or equivalent, available from a chromatography supply company)

Water Chemistry labels (if not already filled out and attached to sample containers at base site)

Soft-lead pencils and write-in-rain-type pens for filling out field data forms and notebook entries

Fine-tipped indelible markers for filling out labels

Roll or box of tape strips

Field operations and methods documents

First aid kit

Backpack

Extra zipper-lock bags
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Items: 

Optional Items (may be required for specific studies):

60 mL glass bottles with septum caps and with completed sample labels attached

Calibrated multiparameter sonde, data logger, and cable, with extra batteries

Calibration standards, quality control check samples, DIW, rinse bottles, waste tray and container, calibration cup, 

and sensor guard for sonde (multiple sensors combined in a unit that is lowered into the water)

Sonde calibration and post-calibration record forms

Measuring tape

Waders or high-top waterproof boots for wading

Clear packaging tape to cover labels

Dissolved oxygen/temperature meter with probe

DO repair kit containing additional membranes and probe filling solution

Conductivity meter with probe

Table H-2. Checklist of material needed for use in field for site documentation.

Items 

A. Available site documentation records for previously established sites:

site location maps, topographic maps, and road maps

site descriptions and access notes

site tag numbers and tag tree descriptions (where applicable)

site coordinates

site photos

B. Preliminary site documentation for new sites:

site location maps, topographic maps, and road maps, indicating approximate site locations

general site descriptions and access notes

C. General material for site documentation

regional-scale topographic and road maps

Stream or Lake Sampling Site Documentation Forms on waterproof paper

clipboard or field notebook and pens for use with waterproof paper

GPS unit with replacement batteries

digital camera with charged battery and charged replacement battery

site tags, aluminum nails, and hammer (if applicable)

measuring tape

blaze orange material for flagging tag trees in photos (if applicable)

gate keys (if needed )

cell phone with numbers of project staff and management agency offices
APPENDIX H. TRAINING CHECKLISTS

315

Table H-3. Equipment and supply checklist for measuring stream discharge.

Quantity Item 

1 Surveyor’s telescoping leveling rod (7 m long, metric scale, round cross-section)

1 50-m fiberglass measuring tape and reel

1 Small bubble level to make sure the tape is level

1 Current velocity meter, probe, and operating manual

1-2 Extra batteries for velocity meter

1 Top-set wading rod (metric or English scale) for use with current velocity meter

1 Portable weir with 60° “V” notch (optional)

1 Plastic sheeting to use with weir (optional)

1 Plastic bucket (or similar container) with volume graduations

1 Stopwatch

1 Covered clipboard

Soft (#2) pencils

Stream Discharge forms (one per stream, plus extras if needed for timed filling procedure or 

additional velocity-area intervals)

1 copy Field operations and methods documents

1 set Laminated sheets of procedure tables and/or quick reference guides for stream discharge

Table H-4. Lake verification checklist.

Item 

Site information folder for lake to be sampled

Clipboard

Lake Sampling Site Documentation Form

Field notebook

Sampling permit (if needed)

GPS unit with manual, extra battery pack

50-m line to attach to rock anchor
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Table H-5. Checklist of equipment and supplies for benthic macroinvertebrates. (Source: Peck et al. 2006.)

Quantity Item 

1 Modified kick net (D-frame with 500 μm mesh) and 4-ft handle

Spare net(s) and/or spare bucket assembly for end of net

1 Watch with timer or a stopwatch

2 Buckets, plastic, 8- to 10-qt capacity (collapsible for back country)

1 Sieve with 500 μm mesh openings or sieve-bottomed bucket, 500 μm mesh openings

2 pr. Watchmakers’ forceps (straight and curved)

1 Wash bottle, 1-L capacity, labeled STREAM WATER

1 Small spatula, spoon, or scoop to transfer sample

1 Funnel, with large bore spout

4 to 6

Each

Sample jars, HDPE plastic with leakproof screw caps, 500-ml and/or 1-L capacity, suitable for 

use with ethanol

2 gal 95% ethanol, in a suitable container (smaller amounts can be carried in for back country work 

or ethanol can be added at the vehicle after returning from the field)

2 pr. Rubber gloves suitable for use with ethanol

1 Cooler (with suitable absorbent material) for transporting ethanol and samples in vehicle

2 Preprinted benthic sample labels with sample ID numbers

4 Preprinted benthic sample labels without sample ID numbers

6 Blank labels on waterproof paper for placing inside of jars

1 Sample Collection Form for site

Soft (#2) lead pencils

Fine-tip indelible markers

1 pkg. Clear tape strips

4 rolls Plastic electrical tape

1 Knife, pocket, with at least two blades

1 Scissors

1 Pocket-sized field notebook (optional)

1 pkg. Kim-wipes in small re-sealable plastic bag

1 copy Field operations and methods manual

1 set Laminated sheets of procedure tables and/or quick reference guides for benthic 

macroinvertebrates
APPENDIX H. TRAINING CHECKLISTS

317

Table H-6. Equipment and supplies for collecting zooplankton samples. (Source: U.S. EPA 2007.)

Quantity1 Item 

2 Wisconsin fine mesh (80 μm2) net with attached collection bucket

2 Wisconsin coarse mesh (243 μm2) net with attached collection bucket

2 Sample line, marked at 0.5 m increments

2 Secchi disk with cable

2/site+ 125 ml wide-mouth polyethelene sample jars (two per site, plus additional for replicates and other 

back-up sampling)

1 Squirt bottle with DIW

95% ethanol

2/site+ CO2 tablets

1/site+ 500 ml wide-mouth container

2 Two lids converted to form strainers (one with 80 μm, one with 243 μm mesh), made by drilling two 

holes in each lid and gluing a piece of the netting to the inside of the lid using silicone glue

2/site+ Zipper lock-type plastic bag

Clear tape for covering labels

Electrical tape

1/site+ Zooplankton Sample Data Form

Pencils and permanent markers

Mild (10%) bleach solution for cleaning net and strainer lids between lakes; backwash net with a 

garden hose after use

1 It is advisable to include some extras, beyond what is needed for the number of sites to be sampled. 2 These two mesh sizes (80 and 243 μm) are general guidelines. Other sizes could be used.
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Table H-7. Checklist of equipment and supplies for sampling lake water chemistry and Secchi depth.

Quantity Item 

Standard Items:

1 Field thermometer

1-2 Sample bottle(s) with completed sample label attached (in clean plastic bag). Include a second bottle if 

sampling at that site is to be replicated

2-4 60 mL plastic syringes (with Luer type tip) or glass bottles with septum caps with completed sample labels 

attached

1 Plastic container with snap-on lid to hold filled syringes

2-4 Syringe valves (Mininert® with Luer type adapter, or equivalent, available from a chromatography supply 

company)

1 Cooler with 4 to 6 plastic bags (1-gal) of ice or a medium or large opaque garbage bag to store the water 

sample at shoreline

1 Lake Sampling Record Form

1 set Water Chemistry labels (if not already filled out and attached at base site)

2-4 Soft-lead pencils and write-in-rain pens for filling out field forms and notebook entries

2-4 Fine-tipped indelible waterproof markers for filling out labels

1 copy Field operations and methods documents

2-4 Plastic gloves stored in a secure plastic bag

1 Survey grade global positioning system and compass

1 Digital camera with extra memory cards and batteries

1 Backpack with waterproof cover (if site is not accessible by vehicle)

1 Van Dorn sampler with messenger and cable

1 Raft or float tube with pump for inflating

1 First aid kit

1 Locally determined safety equipment

1 Secchi disk and line (with depth increments)

1 Measuring tape

Optional Items:

roll/box Clear packaging tape to cover labels (tape strips)

1 Dissolved oxygen/temperature meter with probe

1 DO repair kit containing additional membranes and probe filling solution

1 Conductivity meter with probe

1 250-mL or 500-mL plastic bottle of conductivity QCCS labeled RINSE (in plastic bag)

1 250 mL or 500-mL plastic bottle of conductivity QCCS labeled TEST (in plastic bag)
319

APPENDIX I. 

JOB HAZARD ANALYSIS The following pages show an example Job Hazard Analysis (JHA). This JHA may be used but 

should be revised to include any safety concerns specific to your area.
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U.S. Department of Agriculture 1. WORK PROJECT/ACTIVITY 2. LOCATION 3. UNIT

Forest Service

JOB HAZARD ANALYSIS (JHA) 4. NAME OF ANALYST 5. JOB TITLE 6. DATE PREPARED

References-FSH 6709.11 and -12

(Instructions Following Form)

7. Tasks/Procedures 8. HAZARDS 9. ABATEMENT ACTIONS

Engineering Controls * Substitution * Administrative Controls * PPE

Back Country Travel Overdue Plan ahead. Leave an itinerary of planned trip and follow carefully. Establish regular check 

in times. Never travel or work alone in remote areas. Each person should carry: radio with 

extra batteries or well charged batteries (know proper channels and tones to use), first-aid 

kit, compass and map, waterproof matches, fire starter, pocket knife, survival kit, flashlight, 

extra food, extra clothing, rain gear, signal mirror, extra water.

Disorientation Find sheltered spot and prepare camp. If unable to orient yourself use signal mirror, flares 

or fire to attract attention.

Environmental Hazards Watch weather conditions. Choose safe travel routes, avoid snag or rock slide areas.

Maintain secure footing and working positions. Be on guard against injury from falling 

trees, snags, limbs, rolling logs, or rocks. Maintain safe distance between people (10 feet).

Be sure others know where you are. Wear eye protection.

Contaminated Water/Giardia Carry extra water; filter water; use water tablets or boil water before drinking to avoid 

contamination by giardia. 

Horse Travel Riding horses is dangerous. Death or serious injury 

may occur. Horses can spook, trip, and fall. Some 

hazards to be aware of while riding:

Rough terrain

Rolling debris on trails, dust, brush and limbs

Fast-moving water crossings

Snow banks 

Back packers

Foot getting stuck in stirrup while getting off

Falling from horse

Personal protective equipment (PPE) will be riding boots, long sleeve shirt and pants. 

Riding helmets recommended for inexperienced riders.

Carry a radio at all times and check in and out with the local forest.

Be alert, talk to the animals while working with them to make them aware of your 

presence. Watch for falling debris and wildlife that may spook your animals. When in rough 

terrain, keep only the balls of your feet in the stirrups, or get off and walk. Check water 

crossing before entering water. Take extra time on switchbacks to give the pack animals 

time to round the corners. Snow banks contain air pockets underneath; consider firmness 

of snow and possible entrapment of stock in snow while crossing. If it is warm, do not 

cross over deep snow. Talk to backpackers and ask them to step to the side of the trail 

and stand quietly while you pass.

Vehicle Travel/Driving Vehicle accidents and associated injury Always wear safety belts and make sure everyone is buckled up! Drive carefully on heavily 

travelled roadways. Driving defensively means anticipating the other drivers actions before 

they happen. Back your vehicle in when parking and use a ground guide when available. 

Drive carefully in snow and mud, chain up BEFORE you get stuck. Don't attempt 

accessing remote areas in poor conditions. Roads are narrow, drive defensively, giving 

yourself enough time/space to react to other drivers. Maintain stopping distance of half the 

distance you can see. Drive with headlights on. Stop and take a break if you feel sleepy 

while driving, or let someone else drive (work–rest-ratio: Drive no more than 10 hours in 

one day and take a 10 minute break every two hours). If possible, remove hazards from 

roadbed rather than try to drive over or around them. 
321

In the Field Slips and falls/Balance/Crossing Snow Use traction devices on shoes and waders. Move slowly, take your time. Use a 

walking staff to provide a three point support. Snow banks contain air pockets 

underneath; consider firmness of snow before crossing or avoid crossing snow 

patches when possible.

Crossing streams with high flow velocity, drowning Evaluate a stream before entering. Follow the "rule of 10" 1) if stream is 1 foot deep 

and flowing @10 ft/sec, it is too hazardous to wade 2) if stream is 2 feet deep and 

flowing at 5 ft/sec, it is too hazardous to wade. If you do enter a stream and discover it 

is too dangerous to wade, back out using your wading pole for balance. Secure packs 

with quick release straps and be ready to discard if an emergency arises. Be prepared 

to administer CPR.

Mid-lake water sampling : Float tube Wear a life vest. Return to shore if you become extremely tired. Have other crew 

members on shore watch for hazards in the water.

Mid-lake water sampling: Van dorn samplers Be cautious when operating sampler, it can trap fingers.

Exposure

Cold weather conditions

Hot weather conditions

Sunburn

Severe weather

Hypothermia: Work in teams of two. Have warming devices available. Wear proper 

equipment that is in good condition. Be aware of signs of hypothermia, its prevention, 

detection, and its treatment. Stay in tune to current weather and extended forecasts.

Heat exhaustion: maintain adequate water intake by drinking water periodically 

throughout the day to avoid dehydration. Wear loose-fitting light colored clothing to 

prevent overheating.

Sunburn: Wear protective clothing, hat, sunglasses, and sunscreen (apply frequently).

Severe weather: During lightning storms or when a storm is approaching, avoid high 

points, move to low ground, and stay low, not under a tree.

Hazard trees/Blow-down/heavy debris Be aware of your surroundings, including hazard trees with hanging or leaning debris 

that may be dislodged and fall, especially during high winds. Do not rest or camp 

under hazard trees or hanging/leaning debris in trees.

Scrapes and punctures Wear proper clothing, long sleeved shirts and pants. Use first aid as necessary. 

Eye injuries Travel with care through heavy brush. Use eye protection in brushy areas.

Insect bites/stings Avoid wearing heavy fragrances. Carry first-aid and sting relief kits. Make sure all 

crew members are informed about others who are allergic and what to do if they need 

assistance. Carry necessary emergency medication.

Animal encounters Bears: store food away from sleeping area. If you encounter one make lots of noise 

and back away. Rodents: (haunta virus) avoid feces and dust. Store food tightly and 

dispose if contaminated. Rattle snakes: watch your step on the trail and step over 

logs/rocks and other obstacles.

10. LINE OFFICER SIGNATURE 11. TITLE 12. DATE

(over)
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JHA Instructions (References-FSH 6709.11 and .12) 

The JHA shall identify the location of the work project or activity, the name of employee(s) writing the JHA, the 

date(s) of development,and the name of the appropriate line officer approving it. The supervisor acknowledges 

that employees have read and understand the contents, have received the required training, and are qualified to 

perform the work project or activity. 

Blocks 1, 2, 3, 4, 5, and 6: Self-explanatory.

Block 7: Identify all tasks and procedures associated with the work project or activity that have potential to 

cause injury or illness to personnel and damage to property or material. Include emergency 

evacuation procedures (EEP).

Block 8: Identify all known or suspect hazards associated with each respective task/procedure listed in block 7.

For example:

a. Research past accidents/incidents

b. Research the Health and Safety Code, FSH 6709.11 or other appropriate literature.

c. Discuss the work project/activity with participants

d. Observe the work project/activity

e. A combination of the above

Emergency Evacuation Instructions (Reference FSH 6709.11)

Work supervisors and crew members are responsible for developing and discussing field emergency evacuation 

procedures (EEP) and alternatives in the event a person(s) becomes seriously ill or injured at the worksite.

Be prepared to provide the following information:

a. Nature of the accident or injury (avoid using victim's name).

b. Type of assistance needed, if any (ground, air, or water evacuation).

c. Location of accident or injury, best access route into the worksite (road name/number), identifiable 

ground/air landmarks.

d. Radio frequency(s).

e. Contact person. 

f. Local hazards to ground vehicles or aviation.

g. Weather conditions (wind speed & direction, visibility, temp).

h. Topography. 

i. Number of person(s) to be transported.

j. Estimated weight of passengers for air/water evacuation. 

The items listed above serve only as guidelines for the development of emergency evacuation procedures. 

JHA and Emergency Evacuation Procedures Acknowledgement

Block 9: Identify appropriate actions to reduce or eliminate the hazards identified in block 8. Abatement 

measures listed below are in the order of the preferred abatement method:

We, the undersigned work leader and crew members, acknowledge participation in the development of this JHA 

(as applicable) and accompanying emergency evacuation procedures. We have thoroughly discussed and 

understand the provisions of each of these documents:

a. Engineering Controls (the most desireable method of abatement).

For example, ergonomically designed tools, equipment, and SIGNATURE and DATE SIGNATURE and DATE

furniture.

b. Substitution. For example, switching to high flash point, non-toxic solvents.

Work Leader

c. Administrative Controls. For example, limiting exposure by reducting the work schedule; 

establishing appropriate procedures and practices.

d. PPE (least desirable method of abatement). For example, using hearing protection when 

working with or close to portable machines 

(chain saws, rock drills portable water pumps).

e. A combination of the above.

Block 10: The JHA must be reviewed and approved by a line officer. Attach a 

copy of the JHA as justification for purchase orders when procuring 

PPE. 

Blocks 11 and 12: Self-explanatory.
Editing, design, and layout by Forest Service Publishing Arts, a division of the Washington Office National 

Forest System Business Administration Support Services staff.