v>EPA
United States
Environmental Protection
Agency
Office of Water
4503F
EPA841-B-97-003
November 1997
Volunteer Stream Monitoring
A Methods Manual
-------
-------
Volunteer Stream Monitoring:
A Methods Manual
-------
Acknowledgments
This draft manual was developed by the U.S. Environmental Protection Agency through
contract no. 68-C3-0303 with Tetra Tech, Inc. and through cooperative agreement no.
CT-901837-01-0 with the River Watch Network. The project manager was Alice Mayio,
USEPA Office of Wetlands Oceans and Watersheds. Principal authors include Eric
Dohner, Abby Markowitz, Michael Barbour, and Jonathan Simpson of Tetra Tech, Inc.;
Jack Byrne and Geoff Dates of River Watch Network; and Alice Mayio of USEPA.
Illustrations are by Emily Faalasli, Tetra Tech, Inc. In addition, a workgroup of volunteer
monitoring program coordinators contributed significantly to this product. The authors
wish to thank, in particular; Carl Weber of the University of Maryland and Save Our
Streams; Jay West and Karen Firehock of the Izaak Walton League of America; Anne
Lyon of the Tennessee Valley Authority; and the many reviewers who provided construc-
tive and insightful comments to early drafts of this document. This manual would not
have been possible without their invaluable advice and assistance.
NOTICE:
This document has been reviewed in accordance with U.S. Environmental Protection
Agency policy and approved for publication. Mention of trade names or commercial
products does not constitute endorsement or recommendation for use.
-------
Contents
Chapter 1—Introduction , 1
1.1 Manual Organization 3
Chapter 2—Elements of a Stream Study 5
2.1 Basic Concepts 6
2.2 Designing the Stream Study j , 12
2.3 Safety Considerations J 19
2.4 Basic Equipment 21
Chapter 3—Watershed Survey Methods 23
3.1 How to Conduct a Watershed Survey 25
3.2 The Visual Assessment .; 29
Chapter 4—Macroinvertebrates and Habitat 37
4.1 Stream Habitat Walk 43
4.2 Streamside Biosurvey 61
4.3 Intensive Stream Biosurvey 86
Chapter 5—Water Quality Conditions 125
5.1 Stream Flow > 134
5.2 Dissolved Oxygen and Biochemical Oxygen Demand 139
5.3 Temperature 148
5.4 pH 150
5.5 Turbidity 153
5.6 Phosphorus 158
5.7 Nitrates 165
5.8 Total Solids 171
5.9 Conductivity 173
5.10 Total Alkalinity J.: 176
5.11 Fecal Bacteria 180
Chapter 6—Managing and Presenting Monitoring Data 187
6.1 Managing Volunteer Data 189
6.2 Presenting the Data , 192
6.3 Producing Reports 198
Appendices 201
A. Glossary •. •' 202
B. Scientific Supply Houses < 205
C. Calculating Latitude and Longitude '. 207
-------
-------
INTRODUCTION
-jr*'%«£r'X£ ~&l£»"i ~' . ^f, J^?^>&£tJ~~"<*r
•* v i •i.s&f^t *" ***?*H « s*^ »T»\***f*«?_^,«e5, *J*^* ? ":"5 ^* -£-
_# *¥« js*r ii,r »-V->^>H-^?' ^rs^ ^7^,. "•" A
P^i»^?r§^^
&„«— '-" "lix. »i ""-jk^fc /s *' X ~i**3CS«^X
-------
INTRODUCTION
As part of its commitment to volun-
teer monitoring, the U.S. Environ-
mental Protection Agency (EPA)
has worked since 1990 to develop a series
of guidance manuals for volunteer pro-
grams. Volunteer Stream Monitoring: A
Methods Manual, the third in the series, is
designed as a companion document to
Volunteer Water Monitoring: A Guide for
State Managers. The guide describes the
role of volunteer monitoring in state
programs and discusses how managers can
best organize, implement, and maintain
volunteer programs. This document builds
on the concepts discussed in the Guide for
State Managers and applies them directly to
streams and rivers.
Streams and rivers are monitored by
more volunteer programs than any other
waterbody type. According to the fourth
edition of the National Directory of Volun-
teer Environmental Monitoring Programs
(January 1994), three-quarters of the more
than 500 programs listed conduct some sort
of stream assessment as part, or all, of their
monitoring project.
As the interest in monitoring streams
grows, so too does the desire of groups to
apply an integrated approach to the design
and implementation of programs. More and
more, volunteer monitors are interested in
taking a combination of physical, chemical,
and biological measurements and are
beginning to understand how land uses in a
watershed influence the health of its
waterways. This document includes sec-
tions on conducting in-stream physical,
chemical, and biological assessments as
well as land-use or watershed assessments.
The chemical and physical measure-
ments described in this document can be
applied to rivers or streams of any size.
However, the biological components
(macroinvertebrates and habitat) should be
applied only to "wadable" streams (i.e.,
where streams are small in width and
relatively shallow in depth, and where both
banks are clearly visible).
The purpose of this manual is not to
mandate new methods or override methods
currently being used by volunteer monitor-
ing groups. Instead, it is intended to serve
as a tool for program managers who want to
launch a new stream monitoring program or
enhance an existing program. Volunteer
Stream Monitoring presents methods that
have been adapted from those used success-
fully by existing volunteer programs.
Further, it would be impossible to
provide monitoring methods that are
uniformly applicable to all stream water-
sheds or all volunteer programs throughout
the Nation. Factors such as geographic
region, program goals and objectives, and
program resources will all influence the
specific methods used by each group. This
manual therefore urges volunteer program
coordinators to work hand-in-hand with
state and local water quality professionals
or other potential data users in developing
and implementing a volunteer monitoring
program. Through this partnership, volun-
teer programs gain improved credibility and
access to professional expertise and data;
agencies gain credible data that can be used
in water quality planning. Bridges between
citizens and water resource managers are
also the foundation for an active, educated,
articulate, and effective constituency of
environmental stewards. This foundation is
an essential component in the management
and preservation of our water resources.
EPA has developed two other methods
manuals in this series. Volunteer Lake
Monitoring: A Methods Manual was
published in December 1991. Volunteer
Estuary Monitoring: A Methods Manual
was published in December 1993. To
obtain any or all of these documents,
contact:
U.S. Environmental Protection Agency
Office of Wetlands, Oceans, and Watersheds
Volunteer Monitoring (4503F)
401 M Street, SW
Washington, DC 20460
-------
INTRODUCTION
1.1
Manual Organization
Volunteer Stream Monitoring: A
Methods Manual is organized into six
chapters. All chapters include references for
further reading.
Chapter One: Introduction
The first chapter introduces the manual
and outlines its organization.
Chapter Two: Elements of a
Stream Study
Chapter 2 introduces the concept of the
stream environment and presents informa-
tion on the leading sources of pollution
affecting streams in the United States. It
then discusses in some detail 10 questions
volunteer program coordinators must
answer in designing a stream study, from
knowing why monitoring is taking place to
determining how the program will ensure
the data collected are credible. The chapter
includes a highlight on training volunteer
monitors. The chapter concludes with
safety and equipment considerations.
Chapter Three: Watershed Survey
Methods
This chapter describes how to conduct
a watershed survey (also known as a
watershed inventory or visual survey),
which can serve as a useful first step in
developing a stream monitoring program. It
provides hints on conducting a background
investigation of a watershed and outlines
steps for visually assessing the stream and
its surrounding land uses.
Chapter Four:
Macroinvertebrates and Habitat
In this chapter, three increasingly
complex methods of monitoring the
biology of streams are presented. The first
is a simple stream survey that requires little
training or preparation; the second is a
widely used macroinvertebrate sampling
and stream survey approach that yields a
basic stream rating while monitors are still
at the stream; and the third is a macroinver-
tebrate sampling and advanced habitat
assessment approach that requires profes-
sional and laboratory support but can yield
data on comparatively subtle stream
impacts.
Chapter Five: Water Quality and
Physical Conditions
Chapter 5 summarizes techniques for
monitoring 10 different constituents of
water: dissolved oxygen/biochemical
oxygen demand, temperature, pH, turbid-
ity, phosphorus, nitrates, total solids,
conductivity, total alkalinity, and fecal
bacteria. The chapter begins with a discus-
sion on preparing sampling containers,
highlights basic steps for collecting
samples, and discusses taking stream flow
measurements. This chapter discusses why
each parameter is important, outlines
sampling and equipment considerations,
and provides instructions on sampling
techniques. , ,
Chapter Six: Managing and
Presenting Monitoring Data
Chapter 6 outlines basic principles of
data management, with an emphasis on
proper quality assurance/quality control
procedures. Spreadsheets, databases, and
mapping software are discussed, as are
basic approaches to presenting volunteer
data to different audiences. These ap-
proaches include simple graphs, summary
statistics, and maps. Lastly, the chapter
briefly discusses ideas for distributing
monitoring results to the public.
-------
INTRODUCTION
Appendices
• Appendix A provides a glossary of
terms used in this manual.
• Appendix B lists a number of
scientific supply houses where
monitoring and analytical equipment
can be purchased.
• Appendix C discusses how to
determine the latitude and longitude
of monitoring locations.
References and Further Reading
Ely, E. 1994. A Profile of Volunteer
Monitoring. Volunteer Monitor. 6(1):4.
Ely, E. 1994. The Wide World of Monitor-
ing: Beyond Water Quality Testing.
Volunteer Monitor. 6( 1 ) : 8 .
Lee, V. 1994. Volunteer Monitoring: A
Brief History. Volunteer Monitor.
USEPA. 1996. The Volunteer Monitor' s
Guide To Quality Assurance Project
Plans. EPA 841-B-96-003. September.
Office of Wetlands, Oceans, and Water-
sheds, 4503F, Washington, DC 20460.
USEPA. 1994. National Directory of
Volunteer Environmental Monitoring
Programs, fourth edition. EPA 841-B-
94-001. January. Office of Wetlands,
Oceans, and Watersheds, 4503F, Wash-
ington, DC 20460.
USEPA. 1993. Volunteer Estuary Monitor-
ing: A Methods Manual, EPA
842-B-93-004, December. Office of
Wetlands, Oceans, and Watersheds,
4503F, Washington, DC 20460.
USEPA. 1991. Volunteer Lake Monitoring:
A Methods Manual, EPA 440/4-91-002,
December. Office of Wetlands, Oceans,
and Watersheds, 4503F, Washington, DC
20460.
USEPA. 1990. Volunteer Water Monitor-
ing: A Guide for State Managers, EPA
440/4-90-010, August. Office of Wet-
lands, Oceans, and Watersheds, 4503F,
Washington, DC 20460.
-------
ELEMENTS OF A STREAM STUDY
-------
ELEMENTS OF A STREAM STUDY
This chapter is divided into three
sections. The first section provides
a review of basic concepts concern-
ing watersheds, the water cycle, stream
habitat, and water quality. This background
information is essential for designing a
stream monitoring program that provides
useful data.
Section 2.2 presents the 10 critical
questions that should be answered by
program planners. These include: Why is
monitoring taking place? Who will use the
monitoring data? and What parameters or
conditions will be monitored? The last
section discusses the importance of safety
in the field and laboratory.
2.1
Basic Concepts
Watersheds
A watershed is the area of land from
which runoff (from rain, snow, and springs)
drains to a stream, river, lake, or other body
of water (Fig. 2.1). Its boundaries can be
identified by locating the highest points of
lands around the waterbody. Streams and
rivers function as the "arteries" of the
watershed. They drain water from the land
Figure 2.1
Cross section
of a watershed
Volunteers
should get to
know the
watersheds of
their study
streams.
-------
ELEMENTS OF A STREAM STUDY
as they flow from higher to lower eleva-
tions.
A watershed can be as small or as large
as you care to define it. This is because
several watersheds of small streams usually
exist within the watershed of a larger river.
The watershed of the Mississippi River, for
example, is about 1.2 million square miles
and contains thousands of smaller water-
sheds, each defined by a tributary stream
that eventually drains into a larger river like
the Ohio River or Missouri River and to the
Mississippi itself.
The River System
As streams flow downhill and meet
other streams in the watershed, a branching
network is formed (Fig. 2.2). When ob-
served from the air this network resembles
a tree. The trunk of the tree is represented
by the largest river that flows into the ocean
or large lake. The "tip-most" branches are
the headwater streams. This network of
flowing water from the headwater streams
to the mouth of the largest river is called
the river system.
Water resource professionals have
developed a simple method of categorizing
the streams in the river system. Streams that
have no tributaries flowing into them are
called first-order streams. Streams that .
receive only first-order streams are called
second-order streams. When two second-
order streams meet, the combined flow
becomes a third-order stream, and so on.
The Water Cycle
The water cycle is the movement of
water through the environment (Fig. 2.3). It
is through this movement that water in the
river system is replenished. When precipita-
tion falls to earth in a natural (undeveloped)
watershed in the mid-Atlantic states, for
example, about 40 percent will be returned
to the atmosphere by evaporation or
transpiration (loss of water vapor by
plants). About 50 percent will percolate
into the soil. The remaining 10 percent of
the precipitation moves across the land as
runoff and drains into streams, wetlands,
and other bodies of water (Fig. 2.4, left
panel).
The water that soaks into the ground is
important for maintaining streamflow in
the river network during dry weather.
Percolating water slowly moves downward
through the soil until it drains into an area
where all the pores and cracks in the rock
are saturated with water. The top of this
zone is known as the water table.
Water in this saturated zone moves
laterally, following the laws of gravity and/
or water pressure from above. If the path of
this moving ground water intercepts a
Figure 2.2
A representa-
tion of a river
network with
stream order
marked
-------
ELEMENTS OF A STREAM STUDY
TKAHSPlRfCT/QfJ _C~
/><
{>>j,'f
-0_
'^^^^^^T^mm^^
Figure 2.3
The water
cycle
Water moving
through the
water cycle
replenishes
streams In the
watershed.
Figure 2,4
The fate of
precipitation in
undeveloped
vs. developed
watersheds
Surface runoff
increases and
ground water
recharge
decreases as
watersheds
become devel-
oped.
stream channel, the ground water is dis-
charged into the stream as a spring. The
combination of ground water discharges to
a stream is defined as its baseflow. At times
when there is no surface runoff, the entire
flow of a stream might actually be baseflow
from ground water (Fig. 2.5).
Some streams, on the other hand,
constantly lose water to the ground water.
This occurs when the water table is below
the bottom of the stream channel. Stream
water percolates down through the soil until
it reaches the zone of saturation. Other
streams alternate between losing and
gaining water as the water table moves up
and down according to the seasonal condi-
tions or pumpage by area wells.
The interactions between the water-
shed, soils, and water cycle define the
natural water flow (hydrology) of any
particular stream. Most significant is the
fact that developed land is more impervious
than natural land. Instead of percolating
into the ground, rain hits the hard surfaces
of buildings, pavement, and compacted
ground and runs off into a storm drain or
other artificial structure designed to move
water quickly away from developed areas
and into a natural watercourse. These
conditions typically change the fate of
precipitation in the water cycle (See Fig.
2.4, right panel). For example:
• Less precipitation is evaporated back
to the atmosphere. (Water is trans-
ported rapidly away via storm drains
and is not allowed to stand in pools.)
• Less precipitation is transpired back
to the atmosphere from plants.
(Natural vegetation is replaced by
buildings, pavement, etc.)
• Less precipitation percolates through
the soil to become ground water.
(This can result in a lower water
table and can affect baseflow.)
• More surface runoff is generated and
transported to streams. (Streamflow
becomes more intense during and
immediately after storms.)
Chapter 3, Watershed Survey Methods,
is designed to help volunteers learn about
their watershed. Using the watershed
survey approach, they will become familiar
with their watershed's boundaries, its
hydrologic features, and the human uses of
land and water that might be affecting the
quality of the streams within it.
The Living Stream Environment
A healthy stream is a busy place.
Wildlife and birds find shelter and food
near and in its waters. Vegetation grows
-------
ELEMENTS OF A STREAM STUDY
along its banks, shading the stream, slowing
its flow in rainstorms, filtering pollutants
before they enter the stream, and sheltering
animals. Within the stream itself are fish
and a myriad of insects and other tiny
creatures with very particular needs. For
example, stream dwellers need dissolved
oxygen to breathe; rocks, overhanging tree
limbs, logs, and roots for shelter; vegetation
and other tiny animals to eat; and special
places to breed and hatch their young. For
many of these activities, they might also
need water of specific velocity, depth, and
temperature.
• Human activities shape and alter many
of these stream characteristics. We dam up,
straighten, divert, dredge, dewater, and
discharge to streams. We build roads,
parking lots, homes, offices, golf courses,
and factories in the watershed. We farm,
mine, cut down trees, and graze our live-
stock in and along stream edges. We also
swim, fish, and canoe in the streams
themselves.
These activities can dramatically affect
the many components of the living stream
environment (Fig. 2.6). These components
include:
1. The adjacent watershed includes the
higher ground that captures runoff
and drains to the stream. For pur-
poses of this manual, the adjacent
watershed is defined as land extend-
ing from the riparian zone to 1/4 mile
from the stream.
2. Thefloodplain is the low area of land
that surrounds a stream and holds the
overflow of water during a flood.
3. The riparian zone is the area of
natural vegetation extending outward
from the edge of the stream bank.
The riparian zone is a buffer to
pollutants entering a stream from
runoff, controls erosion, and provides
stream habitat and nutrient input into
the stream. A healthy stream system
Losing Stream
Figure 2.5
L-f—r—-^ Water table
• • • ' ' '
Gaining Stream i watertabie
4.
5.
generally has a healthy riparian
zone. Reductions and impairment of
riparian zones occur when roads,
parking lots, fields, lawns, and other
artificially cultivated areas, bare soil,
rocks, or buildings are 'near the
stream bank.
The stream bank includes both an
upper bank and a lower bank. The
lower bank normally begins at the
normal water line and runs to the
bottom of the stream. The upper
bank extends from the break in the
normal slope of the surrounding land
to the normal high water line.
The streamside cover includes any
overhanging vegetation that offers
protection and shading for the stream
and its aquatic inhabitants.
6. Stream vegetation includes emer-
gent, submergent, and floating
plants. Emergent plants include
plants with true stems, roots, and
leaves with most of their vegetative
parts above the water. Submergent
plants also include some of the same
types of plants, but they are com-
pletely immersed in water. Floating
Streams losing
and gaining
water
The position of
the water table
sometimes plays
a role in determin-
ing the amount of
streamflow.
-------
ELEMENTS OF A STREAM STUDY
Components of
the stream
system
Volunteers
should be aware
that the sur-
rounding land
affects stream
habitat.
7.
8.
SUBSTRATE
(Stream
Bottom)
plants (e.g., duckweed, algae mats)
are detached from any substrate and
are therefore drifting in the water.
The channel of the streambed is the
zone of the stream cross section that
is usually submerged and totally
aquatic.
Pools are distinct habitats within the
stream where the velocity of the
water is reduced and the depth of the
water is greater than that of most
other stream areas. A pool usually an
has soft bottom sediments.
9.
Riffles are shallow, turbulent, but
swiftly flowing stretches of water
that flow over partially or totally
submerged rocks.
10. Runs or glides are sections of the
stream with a relatively low velocity
that flow gently and smoothly with
little or no turbulence at the surface
of the water.
11. The substrate is the material that .
makes up the streambed, such as
clay, cobbles, or boulders.
Whether streams are active, fast-
moving, shady, cold, and clear or deep,
slow-moving, muddy, and warm —or
something in between—they are shaped by
the land they flow through and by what we
do to that land. For example, vegetation in
the stream's riparian zone protects and
serves as a buffer for the stream's stream-
side cover, which in turn shades and
enriches (by dropping leaves and other
organic material) the water in the stream
channel.
Furthermore, the riparian zone helps
maintain the stability of the stream bank by
binding soils through root systems and
helps control erosion and prevent excessive
siltation of the stream's substrate. If human
activities begin to degrade the stream's
riparian zone, each of these stream compo-
nents—and the aquatic insects, fish, and
plants that inhabit them—also begins to
degrade.
Chapter 4 includes methods that
volunteers can use to assess the stream's
living environment—specifically, the
insects that live in the stream and the
physical components of the stream (the
habitats) that support them.
-------
ELEMENTS OF A STREAM STUDY
Water Quality
The water in a stream is always moving
and mixing, both from top to bottom and
from one side of the stream to the other.
Pollutants that enter the stream travel some
distance before they are thoroughly mixed
throughout the flow. For example, water
upstream of a pipe discharging wastewater
might be clean. At the discharge site and
immediately downstream, the water might
be extremely degraded. Further down-
stream, in the recovery zone, overall quality
might improve as pollutants are diluted
with more water. Far downstream the
stream as a whole might be relatively clean
again. Unfortunately, most streams with
one source of pollution often are affected
by many others as well.
Pollution is broadly divided into two
classes according to its source. Point source
pollution comes from a clearly identifiable
point such as a pipe which discharges
directly into a waterbody. Examples of
point sources include factories, wastewater
treatment plants, and illegal straight pipes
from homes and boats.
Nonpoint source pollution comes from
surface water runoff. It originates from a
broad area and thus can be difficult to
identify. Examples of nonpoint sources
include agricultural runoff, mine drainage,
construction site runoff, and runoff from
city streets and parking lots.
Nationally, the pollutants most often
found in the stream environment are not
toxic substances like lead, mercury, or oil
and grease. More impacts are caused by
sediments and silt from eroded land and
nutrients such as the nitrogen and phospho-
rus found in fertilizers, detergents, and
sewage treatment plant discharges. Other
leading pollutants include pathogens such
as bacteria, pesticides, and organic enrich-
ment that leads to low levels of dissolved
oxygen. Common sources of pollution to
streams include:
• Agricultural activities such as crop
production, cattle grazing, and
maintaining livestock in holding
areas or feedlots. These contribute
pollutants such as sediments, nutri-
ents, pesticides, herbicides,
pathogens, and organic enrichment.
• Municipal dischargers such as
sewage treatment plants which
. contribute nutrients, pathogens,
organic enrichment, and toxicants.
• Urban runoff from city streets,
parking lots, sidewalks, storm
sewers, lawns, golf courses, and
building sites. Common pollutants
include sediments, nutrients,
oxygen-demanding substances, road
salts, heavy metals, petroleum
products, and pathogens.
Other commonly reported sources of
pollutants are mining, industrial discharg-
ers (factories), forestry activities, and
modifications to stream habitat and hydrol-
ogy.
Chapter 5 describes methods volun-
teers can use to monitor water quality and
detect pollutants from these sources.
-------
12 I ELEMENTS OF A STREAM STUDY
2.2
Designing the
Stream Study
Before beginning a stream monitoring
study, volunteer program officials should
develop a design or plan that answers the
10 basic questions listed below. Without
answers to these questions, the monitoring
program might well end up collecting data
that do not meet anyone's needs.
Answering these 10 questions is not
easy. A planning committee composed of
the program coordinator, key volunteers,
scientific advisors, program supporters, and
data users should resolve these questions
well before the project gets under way.
Naturally, the committee should also
address other planning questions less
directly related to monitoring design, such
as how to recruit volunteers and how to
secure funding for the project. Answers will
likely change as the program matures. For
example, program coordinators might find
that a method is not producing data of high
enough quality, data collection is too labor-
intensive or expensive, or additional
parameters need to be monitored.
1. Why is the monitoring taking
place?
Typical reasons for initiating a volun-
teer monitoring project include:
• Developing baseline characterization
data
• Documenting water quality changes
over time
• Screening for potential water quality
problems
• Determining whether waters are safe
for swimming
• Providing a scientific basis for
' making decisions on the management
of a stream or watershed
• Determining the impact of a munici-
pal sewage treatment facility,
industrial facility, or land use activity
such as forestry or farming
• Educating the local community or
stream users to encourage pollution
prevention and environmental
stewardship
• Showing public officials that local
citizens care about the condition and
management of their water resources
Of course, an individual program might
be monitoring for a number of reasons.
However, it is important to identify one or
two top reasons and develop the program
based on those objectives.
2. Who will use the monitoring
data?
Knowing your data users is essential to
the program development process. Potential
data users might include:
• State, county, or local water quality
analysts
• The volunteers themselves
• Fisheries biologists
• Universities
• Schoolteachers
• Environmental organizations
• Parks and recreation staff
• Local planning and zoning agencies
• State environmental agencies
• State and local health departments
• Soil and water conservation districts
• Federal agencies such as the U.S.
Geological Survey or U.S. Environ-
mental Protection Agency
Each of these users will have different
data requirements. Some users, such as
-------
ELEMENTS OF A STREAM STUDY
Type
Physical
condition
Biological
condition
Chemical
condition
Approach Applications'
Watershed
survey
Habitat
assessment
Macroinvertebrate
sampling
Water quality
sampling
Determine land use patterns; determine presence of current and
historical pollution sources; identify gross pollution problems;
identify water uses, users, diversions, and stream obstructions
Determine and isolate impacts of pollution sources,
particularly land use activities; interpret biological data; screen
for impairments
Screen for impairment; identify impacts of pollution and
pollution control activities; determine the severity of the pollution
problem and rank stream sites; identify water quality trends;
determine support of designated aquatic life uses
Screen for impairment; identify specific pollutants of;
concern; identify water quality trends; determine support of
designated contact recreation uses;-identify potentialpollution
sources
* Beyond education and promoting stewardship
Table 2.1
Some types of
monitoring
approaches and
their application
government analysts and planning/zoning
agencies, will have more stringent require-
ments than others and will require higher
levels of quality assurance. As the volun-
teer monitoring project is being designed,
program coordinators should contact as
many potential information users as pos-
sible to determine their data needs. It is
important to have at least one user commit-
ted to receiving and using the data. In some
cases that user might be the monitoring
group itself.
3. How will the data be used?
The range of uses of volunteer data is
limited only by the imagination. Volunteer
data could be used, for example, to influ-
ence local planning decisions about where
to site a sewage treatment facility or to
publicize a water quality problem and seek
community solutions. Collected data could
also be used to educate primary school
children about the importance of water
resources. Other data uses include the
support of:
• Local zoning requirements
• A stream protection study
• State preparation of water quality
assessments '
• Screening waters for potential
problems
• The setting of statewide priorities for
pollution control
Each data use potentially has different
data requirements. Knowing the ultimate -
uses of the collected volunteer data will
help determine the right kind of data to
collect and the level of effort required to
collect, analyze, store, and report them.
4. What parameters or conditions
will be monitored?
Determining what to monitor will
depend on the needs of the data users, the
intended use of the data, and the resources
of the volunteer program. If the program's
goal is to determine whether a creek is
suitable for swimming, for example, a
human-health-related parameter such as
-------
ELEMENTS OF A STREAM STUDY
fecal coliform bacteria should be moni-
tored. If the objective is to characterize the
ability of a stream to support sport fish,
volunteers should examine stream habitat
characteristics, the aquatic insect commu-
nity, and water quality parameters such as
dissolved oxygen and temperature. Alterna-
tively, if a program seeks to provide
baseline data useful to state water quality or
natural resource agencies, program design-
ers should consult those agencies to deter-
mine which parameters they consider of
greatest value.
Money for test kits or meters, available
laboratory facilities, help from state or
university advisors, and the abilities and
desires of volunteers will also clearly have
an impact on the choice of parameters to be
monitored. For characterization studies,
EPA usually recommends an approach that
integrates physical, chemical, and biologi-
cal parameters.
5. How good does the monitoring
data need to be?
Some uses require high-quality data.
For example, high-quality data are usually
needed to prove compliance with environ-
mental regulations, assess pollution im-
pacts, or make land use planning decisions.
In other cases the quality of the data is
secondary to the actual process of collect-
ing it. This is often the case for monitoring
programs that focus on the overall educa-
tional aspects of stream monitoring.
Data quality is measured in five
ways—accuracy, precision, completeness,
representativeness, and comparability (see
box—Data Quality Terms).
6. What methods should be
used?
The methods adopted by a volunteer
program depend primarily on how the data
will be used and what kind of data quality
is needed. There are, of course, many
sampling considerations including:
• How samples will be collected (e.g.,
using grab samples or measuring
directly with a meter)
• What sampling equipment will be
used (e.g., disposable Whirl-pak
bags, glass bottles, 500-micron mesh
size kick net, etc.)
• What equipment preparation meth-
ods are necessary (such as container
sterilization or meter calibration)
• What protocols will be followed
(such as the Winkler method for
dissolved oxygen, intensive stream
bioassessment approach for habitat
and benthic macroinvertebrates, etc.)
Analytical questions must also be addressed
such as:
• Will volunteers return to a lab for
macroinvertebrate identification or
dissolved oxygen titration procedures
or conduct them in the field?
• Will a color wheel provide nitrate
data of needed quality, or is a more
sophisticated approach needed?
• Should visual observation and habitat
assessment approaches be combined
with turbidity measures to best
determine the impact of construction
sites?
While sophisticated methods usually
yield more accurate and precise data (if
properly carried out), they are also more
costly and time-consuming. This extra
effort and expense might be worthwhile if
the goal of the program is to produce high-
quality data. Programs with an educational
focus, however, can often use less sensitive
equipment and less sophisticated methods
to meet their goals.
7. Where are the monitoring
sites?
Sites might be chosen for any number
of reasons such as accessibility, proximity
to volunteers' homes, value to potential
-------
ELEMENTS OF A STREAM STUDY
Data Quality Terms
Accuracy Is the degree of agreement between the sampling result and
the true value of the parameter or condition being measured. Accuracy is
most affected by the equipment and the procedure used to measure the
parameter.
Precision, on the other hand, refers to how well you are able to
reproduce the result on the same sample, regardless of accuracy.
Human error in sampling techniques plays an important role in estimat-
ing precision.
Representativeness is the degree to which collected data actually
represent the stream condition being monitored. It is most affected by
site location.
Completeness is a measure of the amount of valid data actually
obtained vs. the amount expected to be obtained as specified in the
original sampling design. It is usually expressed as a percentage. For
example, if 100 samples were scheduled but volunteers sampled only 90
times due to bad weather or broken equipment, the completeness record
would be 90 percent.
Comparability represents how well data from one stream or stream site
can be compared to data from another. Most managers will compare
sites as part of a statewide or regional report on the volunteer monitoring
program; therefore, sampling methods should be the same from site to
site.
Imprecise and inaccurate
Precise but inaccurate
Tni>
Vaui
Trui
Valix
Accurate but imprecise
Tiu«
Valui
Precise and accurate
True
Value
users such as state agencies, or location in
problem areas. If the volunteer program is
providing baseline data to characterize a
stream or screen for problems, it might
wish to monitor a number of sites repre-
senting a range of conditions in the stream
watershed (e.g., an upstream "pristine"
area, above and below towns and cities, in
agricultural areas and parks, etc.). For more
specific purposes, such as determining
whether a stream is safe to swim in, it
might only be necessary to sample selected
swimming areas. To determine whether a
particular land use activity or potential
source of pollution is, in fact, having an
impact, it might be best to monitor up-
stream and downstream of the area where
the source is suspected. To determine the
effectiveness of runoff control measures, a
paired watershed approach might be best
(e.g., sampling two similar small water-
sheds, one with controls in place and one
without controls).
A program manager might also select
one or more sites near professionally
monitored sites in order to compare the
quality of volunteer-generated data against
professional data. It might also be helpful
to locate some sites near U.S. Geological
Survey gauging stations, which can provide
useful data on streamflow. Certainly, for
any volunteer program, safety and accessi-
bility (both legal and physical) will be
important in determining site location. No
matter how sampling sites are chosen, most
monitoring programs will need to maintain
the same sites over time and identify them
clearly in their monitoring program design.
When selecting monitoring sites, ask
the following questions. Based on the
answers, you may need to eliminate some
sites or select alternative locations that
meet your criteria:
• Are other groups (local, state, federal
agencies; other volunteer groups;
-------
ELEMENTS OF A STREAM STUDY
schools or colleges) already monitor-
ing this site?
• Can you identify the site on a map
and on the ground?
• Is the site representative of the
watershed?
• Does the site have water in it during
the times of year that monitoring will
take place?
• Is there safe, convenient access to the
site (including adequate parking) and
a way to safely sample a flowing
section of the stream? Is there access
all year long?
• Can you acquire landowner permis-
sion?
• Can you perform all the monitoring
activities and tests that are planned at
this site?
• Is the site far enough downstream of
drains or tributaries? Is the site near
tributary inflows, dams, bridges, or
other structures that may affect the
results?
• Have you selected enough sites for
the study you want to do?
Once you have selected the monitoring
sites, you should be able to identify them
by latitude and longitude. This location
information is critical if your data will
potentially be used in Geographical Infor-
mation Systems (GIS) or in sophisticated
data management systems (See Appendix
C).
8. When will monitoring occur?
A program should specify:
• What time of day is best for sam-
pling. (Temperature and dissolved
oxygen, for example, can fluctuate
naturally as the sun rises and aquatic
plants release oxygen.)
• What time of year is best for sam-
pling. (For example, there is no point
in sampling fecal coliform bacteria at
swimming beaches in the winter,
when no one is swimming, or
sampling intermittent streams at the
height of summer, when because of
dry conditions the streams hold little
water.)
• How frequently should monitoring
take place? (It is possible, for ex-
ample, to conduct too many
biological assessments of a stream
and thereby deplete the stream's
aquatic community. A program
designed to determine whether
polluted runoff is a problem would
do well to monitor after storms and
heavy rainfalls.)
In general, monthly chemical sampling
and twice-yearly biological sampling are
considered adequate to identify water
quality changes over time. Biological
sampling should be conducted at the same
time each year because natural variations in
aquatic insect population and streamside
vegetation occur as seasons change. Moni-
toring at the same time of day and at
regular intervals (e.g., at 2:00 p.m. every 30
days) helps ensure comparability of data
over time.
9. How will monitoring data be
managed and presented?
The volunteer program coordinator
should have a clear plan for dealing with
the data collected each year. Field and lab
data sheets should be checked for complete-
ness, data should be screened for outliers,
and a database should be developed or
adapted to store and manipulate the data.
The elements of such a database should be
clearly explained in order to allow users to
interpret the data accurately and with
confidence.
-------
ELEMENTS OF A STREAM STUDY
Training Volunteer Monitors
Training should be an essential component of any volunteer stream monitoring project. When volunteers are properly trained in the goals of the
volunteer project and its sampling and analytical methods, they:
• Produce higher quality, more credible data. • Save program manager time and effort by becoming
• Better understand their role in protecting water quality. better monitors who require less supervision.
• Are more motivated to continue monitoring. • Feel more like part of a dedicated team.
Some of the key elements to consider in developing a training program for volunteers include the following:
1. Plan ahead When you are in the early stages of developing your training program, decide ;who will do the training, when training will occur,
where it will be held, what equipment and handouts volunteers will receive, and what, in they end, they will learn. Plan on at least one initial
training session at the start of the sampling season and a quality control session somewhat into the season (to see if volunteers are using
the right methods, and to answer questions). If volunteers will be sampling many different chemical parameters or will be conducting
intensive biological monitoring, you should probably schedule two initial training sessions—one to introduce volunteers to the program, and
the other to cover sampling and analytical methods in detail. You might also want to plan a post-season session that encourages volunteers
to air problems, exchange information, and make suggestions for the coming year. Make sure the program planning committee agrees to
the training plan.
2. Put it in writing. Once you've made these decisions, write them all down. Note the training specifics in the program's quality assurance
project plan. It might also help to develop a "job description" for the volunteers that lists the tasks they will perform in the field and lab, and
that identifies the obligations to which they will be held and the schedule they will follow. Hand this out at the first training session. Volun-
teers should leave the session knowing what is expected of them. If they decide not to join after all because the tasks are too onerous, it is
better for you to find out after the first session than later in the sampling year.
3. Be prepared. Nothing will discourage volunteers more than an ill-planned, chaotic initial training session. The elements of a successful
initial training session include:
Enthusiastic, knowledgeable trainers
Short presentations that encourage audience participation and don't strain attention spans
A low ratio of trainers to trainees
Presentations that include why the monitoring is needed, what the program hopes to accomplish, and what will be done with the data
An agenda that is followed (especially start and finish times) |
Good acoustics, clear voices, and interesting audiovisual aids
Opportunities for all trainees to handle equipment, view demonstrations of sampling .protocols, and practice sampling
Instruction on safety considerations
Refreshments and opportunities for trainees to meet one another, socialize, and have fun
Time for questions and answers.
4. Conduct quality control checks. After your initial training session(s), schedule opportunities to "check up" on how your volunteers are
performing. The purpose of these quality control checks is to ensure that all volunteers are monitoring using proper and consistent
protocols, and to emphasize the importance of quality control measures. Some time into the sampling season, observe how volunteers are
sampling, analyzing their samples, identifying macroinvertebrates, and recording their results. Either observe volunteers in small groups at
their monitoring sites or bring them to a central location for an organized quality control session. If your program is involved in chemical
monitoring, you might want all volunteers to analyze the same water sample using their own equipment, or hold a lab exercise in which
volunteers read and record results from equipment and kits that have already been set up. For a biological monitoring program, have
trainers or seasoned volunteers observe sampling methods in the field and provide preserved samples of macroinvertebrates for volunteers
to identify. Reserve time to answer questions, talk about initial findings, and have some fun.
5. Review the effectiveness of your training program. At the end of each training session, encourage volunteers to fill out a training evaluation
form. This form should help you assess the effectiveness of individual trainers and their styles, the handouts and audiovisual aids, the
general atmosphere of the training session, and what the volunteers liked most and least about the session. Use the results of the evalua-
tion to revise training protocols as needed to best meet program and volunteer needs. ;
-------
ELEMENTS OF A STREAM STUDY
Put It in Writing
When you and the volunteer program planning committee have answered the ten project design questions to everyone's satisfaction,
your next critical step is to put it all in writing. The written plan, including sampling and analytical methods, sites, parameters, project goals,
and data quality considerations, is your bible. With a written plan you:
• Document the particulars of your program for your data users
• Educate newcomers to the program
• Ensure that newcomers will use the same methods as those who came before them
• Keep an historical record for future program leaders, volunteers, and data users
Your written plan may simply consist of a study design and standard operating procedures such as a monitoring and lab methods manual.
You may, however, prefer to develop a more comprehensive quality assurance project plan. The quality assurance project plan is a document
that outlines the procedures you will use to ensure high quality data when conducting sample collection and analysis in your program.
By law, any water quality monitoring program that receives EPA funding is required to have an EPA-approved quality assurance project
plan. Even if you don't receive EPA funding, you will find that preparing a written plan helps ensure that your data are used with confidence,
now and In the future. (See The Volunteer Monitor's Guide to Quality Assurance Project Plans (EPA 841 -B-96-003 September 1996) for more
Information.)
Program coordinators will also have to
decide how they want to present data
results, not only to the general public and to
specific data users, but also to the volun-
teers themselves. Different levels of
analysis might be needed for different
audiences. A volunteer group collecting
data for state or county use should consult
with the appropriate agency before invest-
ing in computerized data management
software because the agency could have
specific needs or recommendations based
on its own data management protocols.
10. How will the program ensure
that data are credible?
Developing specific answers to ques-
tions 1-9 is the first step in ensuring that
data are credible. Credible data meet
specific needs and can be used with confi-
dence for those needs. Other steps include:
• Properly training, testing, and
retraining volunteers
• Evaluating the program's success
after an initial pilot stage and making
any necessary adjustments
• Assigning specific quality assurance
tasks to qualified individuals in the
program
• Documenting in a written plan all
the steps taken to sample, analyze,
store, manage, and present data
A written plan, known as a quality
assurance project plan, can be elaborate or
simple depending on the volunteer
program's goals. Its essential feature,
however, is that it documents how the data
are to be generated. Without such knowl-
edge, the data cannot be used with confi-
dence. It is also important for educating
future volunteers and data users about the
program and the data. People might be
analyzing the data 5 or 10 or more years
later to study trends in stream quality.
(Note: EPA requires that any monitoring
program sponsored by EPA through grants,
contracts, or other formal agreement must
carry out a quality assurance/quality
control program and develop a quality
assurance project plan.)
-------
ELEMENTS OF A STREAM STUDY
2.3
Safety Considerations
One of the most critical considerations
for a volunteer monitoring program is the
safety of its volunteers. All volunteers
should be trained in safety procedures and
should carry with them a set of safety
instructions and the phone number of their
program coordinator or team leader. Safety
precautions can never be overemphasized.
The following are some basic common
sense safety rules. At the site:
• Always monitor with at least one
partner. Teams of three or four
people are best. Always let someone
else know where you are, when you
intend to return, and what to do if
you don't come back at the appointed
time.
• Develop a safety plan. Find out the
location and telephone number of the
nearest telephone and write it down.
Locate the nearest medical center
and write down directions on how to
get between the center and your
site(s) so that you can direct emer-
gency personnel. Have each member
of the sampling team complete a
medical form that includes emer-
gency contacts, insurance
information, and pertinent health
information such as allergies, diabe-
tes, epilepsy, etc.
• Have a first aid kit handy (see box
below). Know any important medical
conditions of team members (e.g.,
heart conditions or allergic reactions
to bee stings). It is best if at least one
team member has first aid/CPR
training.
• Listen to weather reports. Never go
sampling if severe weather is pre-
dicted or if a storm occurs while at
the site.
Never wade in swift or high water.
Do not monitor if the stream is at
flood stage.
If you drive, park in a safe location.
Be sure your car doesn't pose a
hazard to other drivers and that you
don't block traffic.
Put your wallet and keys in a safe
place, such as a watertight bag you
keep in a pouch strapped to your
waist. Without proper precautions,
wallet and keys might end up
downstream.
Never cross private property without
the permission of the landowner.
Better yet, sample only at public
access points such as bridge or road
crossings or public parks. Take
along a card identifying you as a
volunteer monitor.
First Aid Kit
The minimum first aid kit should contain the following items:
• Telephone numbers of emergency personnel such as the police and an
ambulance service. :
• Several band-aids for minor cuts.
• Antibacterial or alcohol wipes.
• First aid creme or ointment.
• Several gauze pads 3 or 4 inches square for deep wounds with excessive
bleeding.
• Acetaminophen for relieving pain and reducing fever.
• A needle for removing splinters.
• A first aid manual which outlines diagnosis and treatment procedures.
• A single-edged razor blade for minor surgery, cutting tape to size, and
shaving hairy spots before taping.
• A 2-inch roll of gauze bandage for large cuts.
• A triangular bandage for large wounds.
• A large compress bandage to hold dressings in place.
• A 3-inch wide elastic bandage for sprains and applying pressure to bleeding
wounds.
• If a participant is sensitive to bee stings, include their doctor-prescribed
antihistamine.
i
Be sure you have emergency telephone numbers and medical information with
you at the field site for everyone participating in field work (including the leader) in
case there is an emergency.
-------
ELEMENTS OF A STREAM STUDY
I Confirm that you are at the proper
site location by checking maps, site
descriptions, or directions.
I Watch for irate dogs, farm animals,
wildlife (particularly snakes), and
insects such as ticks, hornets, and
wasps. Know what to do if you get
bitten or stung.
I Watch for poison ivy, poison oak,
sumac, and other types of vegetation
in your area that can cause rashes
and irritation.
I Never drink the water in a stream.
Assume it is unsafe to drink, and
bring your own water from home.
After monitoring, wash your hands
with antibacterial soap.
Do not monitor if the stream is
posted as unsafe for body contact. If
the water appears to be severely
polluted, contact your program
coordinator.
Do not walk on unstable stream
banks. Disturbing these banks can
accelerate erosion and might prove
dangerous if a bank collapses.
Disturb streamside vegetation as
little as possible.
Be very careful when walking in the
stream itself. Rocky-bottom streams
can be very slippery and can contain
deep pools; muddy-bottom streams
might also prove treacherous in areas
where mud, silt, or sand have accu-
mulated in sink holes. If you must
cross the stream, use a walking stick
to steady yourself and to probe for
deep water or muck. Your partner(s)
should wait on dry land ready to
assist you if you fall. Do not attempt
to cross streams that are swift and
above the knee in depth. Wear
waders and rubber gloves in streams
suspected of having significant
pollution problems.
• If you are sampling from a bridge, be
wary of passing traffic. Never lean
over bridge rails unless you are
firmly anchored to the ground or the
bridge with good hand/foot holds.
• If at any time you feel uncomfort-
able about the condition of the
stream or your surroundings, stop
monitoring and leave the site at
once. Your safety is more impor-
tant than the data!
When using chemicals:
• Know your equipment, sampling
instructions, and procedures before
going out into the field. Prepare
labels and clean equipment before
you get started.
• Keep all equipment and chemicals
away from small children. Many of
the chemicals used in monitoring are
poisonous. Tape the phone number
of the local poison control center to
your sampling kit.
• Avoid contact between chemical
reagents and skin, eye, nose, and
mouth. Never use your fingers to
stopper a sample bottle (e.g., when
you are shaking a solution). Wear
safety goggles when performing any
chemical test or handling preserva-
tives.
• Know chemical cleanup and disposal
procedures. Wipe up all spills when
they occur. Return all unused chemi-
cals to your program coordinator for
safe disposal. Close all containers
tightly after use. Do not switch caps.
• Know how to use and store chemi-
cals. Do not expose chemicals or
equipment to temperature extremes
or long-term direct sunshine.
-------
ELEMENTS OF A STREAM STUDY
2.4
Basic Equipment
Much of the equipment a volunteer will
need is easily obtained from either hard-
ware stores or scientific supply houses.
Other equipment can be found around the
house. In either case, the volunteer program
should clearly specify the equipment its
volunteers will need and where it should be
obtained.
Listed below is some basic equipment
appropriate for any volunteer field activity.
Much of this equipment is optional but will
enhance the volunteers' safety and effec-
tiveness.
• Boots or waders; life jackets if you
are sampling by boat
• Walking stick of known length for
balance, probing, and measuring
• Bright-colored snag- and thorn-
resistant clothes; long sleeves and
pants are best
• Rubber gloves to guard against
contamination
• Insect repellent/sunscreen
• Small first aid kit, flashlight, and
extra batteries
• Whistle to summon help in emergen-
cies
• Refreshments and drinking water
• Clipboard, preferably with plastic
cover
• Several pencils
• Tape measure
• Thermometer
• Field data sheet
• Information sheet with safety
instructions, site location informa-
tion, and numbers to call in
emergencies
• Camera and film, to document
particular conditions
Specific equipment lists for the chemi-
cal and biological monitoring procedures
included in the manual are provided in the
relevant chapters. ,
-------
ELEMENTS OF A STREAM STUDY
References and Further Reading
Dates, G. 1994. A Plan for Watershed-wide
Volunteer Monitoring. The Volunteer
Monitor. 6(2):8.
Ely, E. 1992. Building Credibility. The
Volunteer Monitor 4(2).
Ely, E. 1994. What Parameters Volunteer
Groups Test. The Volunteer Monitor.
Picotte, A. 1994. Citizen's Data Used to Set
Phosphorus Standards. The Volunteer
Monitor. 6(1):18.
Weber, P. and F. Dowman. 1994. The Web
of Water. The Volunteer Monitor.
6(2): 10.
USEPA. 1990. Volunteer Water Monitor-
ing: A Guide for State Managers. EPA
440/4-90-010. August. U.S. Environmen-
tal Protection Agency, Office of Water,
Washington, DC 20460.
USEPA. 1993. EPA Requirements for
Quality Assurance Project Plans for
Environmental Data Operations. EPA
QA/R-5. July. U.S. Environmental
Protection Agency, Quality Assurance
Management Staff, Washington, DC
20460.
USEPA. 1993. Integrating Quality Assur-
ance into Tribal Water Programs. U.S.
Environmental Protection Agency,
Region 8, 999 18th St., Suite 500,
Denver, CO 80202.
USEPA. 1996. The Volunteer Monitor's
Guide To Quality Assurance Project
Plans. EPA 841-B-96-003. September.
Office of Wetlands, Oceans, and Water-
sheds, 4503F, Washington, DC 20460.
-------
WATERSHED SURVEY METHODS I 23
-------
WATERSHED SURVEY METHODS
One of the most rewarding and least
costly stream monitoring activities
a volunteer program can conduct
is the watershed survey. Some programs
call it a windshield survey, a visual survey,
or a watershed inventory. It is, in essence, a
comprehensive survey of the geography,
land and water uses, potential and actual
pollution sources, and history of the stream
and its watershed.
The watershed survey may be divided
into two distinct parts:
• A one-time background investigation
of the stream and its watershed. (To
do this, volunteers research town and
county records, maps, photos, news
stories, industrial discharge records,
and oral histories.)
• A periodic visual assessment of the
stream and its watershed. (To do
this, volunteers walk along the
stream and drive through the water-
shed, noting key features.)
The watershed survey requires little in
the way of training or equipment. Its chief
uses include:
• Screening for pollution problems
• Identifying potential sources of
pollution
• Identifying sites for monitoring
• Helping interpret biological and
chemical information
• Giving volunteers and local residents
a sense of the value of the stream or
watershed
• Educating volunteers and the local
community about potential pollution
sources and the stressors affecting
the stream and its watershed
• Providing a blueprint for possible
community restoration efforts such
as cleanups and tree plantings
To actually determine whether those
stressors are, in fact, affecting the stream
requires additional monitoring of chemical,
physical, or biological conditions.
The watershed survey described in this
chapter was developed from survey ap-
proaches used by programs such as Rhode
Island Watershed Watch, Maryland Save
Our Streams, the Delaware Department of
Natural Resources and Environmental
Control, and Washington's Adopt-A-
Stream Foundation. References are pro-
vided at the end of this chapter for further
information on watershed surveys.
-------
WATERSHED SURVEY METHODS
3.1
How to Conduct a
Watershed Survey
The Background Investigation
Researching the stream is generally a
one-time activity that should yield valuable
information about the cultural and natural
history of the stream and the uses of the
land surrounding it. This information will
prove helpful in orienting new volunteers to
the purpose of the monitoring program, in
building a sense of the importance of the
stream and its role in the watershed, and in
identifying land use activities in the water-
shed with a potential to affect the quality of
the stream. The program might choose to
monitor these areas and activities more
intensively in the future.
The background investigation is
essentially a "detective investigation" for
information on the stream and includes the
following steps:
Taskl
J Determine what you want to
know about your stream
Before beginning the background
investigation, establish what it is you want
to know about the stream you are survey-
ing. Types of information include:
• Location of the stream's headwaters,
its length, where it flows, and where
it empties
• Name and boundaries of the water-
shed it occupies, the population in
the watershed, and the communities
through which it flows
• Roles of various jurisdictions in
managing the stream and watershed
• Percentage of watershed land area in
each town or jurisdiction
• Land uses in the stream's watershed
• Industries and others that discharge
to the stream
• Current uses of the stream (such as
fishing, swimming, drinking water
supply, irrigation)
• Historical land uses
• History of the stream
Any or all of these types of information
should prove valuable to the monitoring
program. You might also uncover other
important information in the process. At a
minimum, the investigation should yield
information on the size of the stream,
watershed boundaries, and general land use
in the area. By establishing categories of
information to investigate, program coordi-
nators can assign volunteers to specific
activities and end up with a complete
picture of the stream that answers many
questions of value to the program.
Task 2 I Determine the tools you will
~™ need
Offered below are some of the tools
you will need to find answers in your
background investigation of the stream.
Stream headwaters, length, tributaries,
final stream destination, and watershed
boundaries are best determined through
maps. Of greatest value are U.S. Geologi-
cal Survey 7 1/2- minute topographic maps
(on a 1:24,000 scale where 1 inch = 2,000
feet). At varying degrees of resolution, they
depict landforms, major roads and political
boundaries, developments, streams, tribu-
taries, lakes, and other land features.
Sporting goods stores and bookstores often
carry these maps, especially for recre-
ational areas that are likely to be hiked or
camped. The maps can also be ordered
through the U.S. Geological Survey (see
box-Obtaining USGS Topographic Maps).
Road, state, and county maps might
also prove helpful in identifying some of
-------
WATERSHED SURVEY METHODS
Obtaining USGS Topographic Maps
The U.S. Geological Survey's Earth Science Information Centers can
provide you with a catalog of available USGS topographic maps, a
brochure on how to use topographic maps, and general information on
ESIC services. Contact the main ESIC office at:
USGS Earth Science Information Center
507 National Center
12201 Sunrise Valley Drive
Reston, VA 22092
1-800-USA-MAPS
You can obtain a free USGS Indexing Catalog to help you identify the
map{s) you need by calling 1-800-435-7627. If you know the coordinates
of the map you need, you can order it directly from:
USGS
Branch of Information Services
Box 25286
Denver, CO 80225
Place your order in writing and include a check for $4.00 per map plus
$3.50 for shipping and handling. The ESIC can also refer you to commer-
cial map distributors that can get you the topographic maps sooner, for a
higher fee. USGS topographic maps might also be available from sporting
goods stores in your area.
these stream and watershed features.
Hydrologic unit maps, also available from
the U.S. Geological Survey but at a
1:100,000 scale of resolution (less detail
than the 7 1/2-minute maps cited above),
might also help you determine hydrologic
watershed boundaries. Atlases and other
reference materials at libraries can prove
helpful in determining facts about popula-
tion in the watershed.
Land uses in the stream watershed
might also be depicted on maps such as
those discussed above. You will verify this
information in the second half of the
watershed survey, when you are actually in
the field observing land around the stream.
Information from maps is particularly
useful in developing a broad statement
about general land use in the stream water-
shed (e.g., land use in the hypothetical
Volunteer Creek watershed is 60 percent
residential, 20 percent parkland/recre-
ational, and 20 percent light industrial).
Other sources of information include:
• Land use plans from local planning
offices, which include information
not only for current land uses but for
potential uses for which the area is
zoned
• Conservation district offices or
offices of the agricultural extension
service or Natural Resources Conser-
vation Service (Formerly the Soil
Conservation Service, these offices
might be able to provide information
on agricultural land in rural areas)
• Local offices of the U.S. Geological
Survey, which might provide a
variety of publications, special
studies, maps, and photos on land
uses and landforms in the area
• Aerial photographs, which might
provide current and historical views
of land uses
Industries and others that discharge to
the stream might be identified at the state,
city, or county environmental protection or
water quality office. (The name of the
agency will vary by locality.) At these
offices, you may ask to see records of
industries with permits to discharge treated
effluent to streams. These records are
maintained through the National Pollutant
Discharge Elimination System (NPDES).
All industrial and municipal dischargers are
required to have permits that specify where,
when, and what they are allowed to dis-
charge to waters of the United States.
Especially in older metropolitan areas,
combined sewers are also potential dis-
charges. Combined sewers are pipes in
which sanitary sewer waste overflow and
storm water are combined in times of heavy
rain. These combined sewers are designed
to discharge directly into harbors and rivers
during storms when the volume of flow in
the sewers exceeds the capacity of the
sewer system. The discharge might include
raw sanitary sewage waste. Combined
-------
WATERSHED SURVEY METHODS
sewers do not flow in dry weather. Maps of
sewer systems can be obtained from your
local water utility.
The state or local environmental agency
should also be able to provide location
information on other potential pollution
sources such as landfills, wastewater
treatment plants, and stormwater detention
ponds.
Current uses of the stream are estab-
lished in state water quality standards,
which specify what the uses of all state
waters should be. These uses can include,
for example, cold water fisheries, primary
contact recreation (swimming) and irriga-
tion. The state also establishes criteria or
limits on pollutants in the waters necessary
to maintain sufficient water quality to
support those uses, as well as a narrative
statement that prohibits degradation of
waters below their designated uses.
Section 305(b) of the Clean Water Act
requires states to report to the U.S. Envi-
ronmental Protection Agency on the
designated uses of their waters, the extent
of the impairment of those uses, and the
causes and sources of impairment. This
information is kept on file at the state water
quality agency. While state reports cannot
specify water uses and degree of impair-
ment in all individual streams in the state,
they are a good starting point. Write to the
state water quality agency for its biennial
water quality (section 305(b)) assessment.
You might also be able to obtain a copy
of your state's water quality standards or
establish contact with a water quality
specialist who can give you information on
standards for your stream. Again, informa-
tion on actual water uses will be verified
and detailed once you walk the stream
during the visual assessment portion of.
your watershed survey.
Historical land uses and the history of
the stream might take some legwork to
uncover. Local historical societies, librar-
ies, and newspaper archives are good places
to start. Look for historical photos of the
area and stories about fishing contests, fish
kills, spills, floods, and other major events
affecting the stream and its watershed.
County or town planning offices might be
able to provide information on when
residential developments were built and
when streams were channelized or di-
verted. State and local transportation
agencies might have records on when
highways and bridges were built. State
environmental regulatory agencies have
records of past or current applications to
modify stream hydrology through dredg-
ing, channelization, and stream bank
stabilization.
Long-time residents are another
invaluable source of information on the
history of your stream. People who fished
or swam in your stream in their youth
might have witnessed how the stream has
changed. They might remember industries
or land use activities of the past—such as
Getting to Know the
Boundaries of Your Watershed
Once you've obtained topographic maps of your area, follow these
steps to draw your watershed boundaries:
1.
2.
3.
4.
Locate and mark the downstream outlet of the watershed. For
rivers and streams, this is the farthest downstream point in which
you are interested.
Locate all water features such as streams, wetlands, lakes, and
reservoirs that eventually flow to the outlet. Start with major
tributaries, then include smaller creeks and drainage channels. To
determine whether a stream is flowing to or from a lake or river,
compare the elevation of land features to that of the waterbody.
Use arrows to mark the direction of stream or wetland flow.
Find and mark the high points (hills, ridges, saddles) on the map.
Then connect these points, following ridges and crossing slopes at
right angles to contour lines. This line forms the watershed
boundary.
If you don't need to know exact watershed boundaries, simply look at
the pattern of streamflow and draw lines dividing different stream systems.
This will give you an idea of the shape of your watershed and those that
border it. Also, once you've identified watershed boundaries, water
features, and flow direction, you might want to transfer this information to
a road map for easier use. '
From: Eleanor Ely, Delineating a Watershed,
The Volunteer Monitor 6(2), Fall 1994.
-------
WATERSHED SURVEY METHODS
Figure 3.1
A topographic
map with a
delineated
watershed.
Volunteers
should learn to
read a topo map
to learn about
the natural and
cultural features
of their study
stream's
watershed
mines or farms—that could have affected
the stream. They might have tales to tell
about fish they once caught or floods that
led to channelization and dams. Assembling
such oral histories is a particularly good
activity for school-age volunteers.
Tasks
Conduct the background
investigation
It is best to conduct your background
investigation of the stream in the early
stages of the volunteer program and use the
information it uncovers to help design the
program's monitoring plan, future activi-
ties, and projects.
The investigation might emphasize
those aspects which are most important to
the volunteers or the watershed, or it might
include all the resources and tools listed
above. In any case, rely on the interests of
the volunteers in designing and conducting
the background investigation, and divide
duties among different volunteers.
-------
WATERSHED SURVEY METHODS
Once the investigation has been con-
ducted, either the program coordinator or
an interested volunteer should compile the
information collected and present it to other
volunteers in written form or at a
program-wide meeting. At a minimum, key
information on land uses, water uses,
watershed boundaries, and dischargers
should be maintained in written form for
program use and for volunteers who might
join the program at a later date. Maps,
photographs, and other information on
previous water quality studies in the
watershed will be of particular value to the
program over time.
Obtaining Aerial Photographs
Historic and current aerial photographs can
be obtained from local, state, and federal
governments, as well as private firms. Try
planning offices, highway departments, soil and
water conservation districts, state departments
of transportation, and universities.
Federal sources of aerial photographs
include:
• USGS Earth Science Information Center
507 National Center
12201 Sunrise Valley Drive
Reston, VA 22092
1-800-USA-MAPS
• USDA Consolidated Farm Service Agencies
Aerial Photography Field Office
222 West 2300 South
P.O. Box 30010
Salt Lake City, UT 84103-0010
801-524-5856
• Cartographic and Architectural Branch
National Archives and Records Administra-
tion
8601 Adelphi Road
College Park, MD 20740-6001
301-713-7040
3.2
The Visual
Assessment
To conduct the visual stream assess-
ment portion of the watershed survey,
volunteers regularly walk, drive, and/or
canoe along a defined stretch of stream
observing water and land conditions, land
and water uses, and changes over time.
These observations are recorded on maps
and on visual assessment data sheets and
passed to the volunteer coordinator, who
can decide whether additional action is
needed. Volunteers might themselves
follow up by reporting on problems such as
fish kills, sloppy construction practices, or
spills they have identified during the visual
assessment.
The basic steps to follow are:
Taskl
Determine the area to be
assessed
The visual assessment will have most
value if the same stream or segment of
stream is assessed each time. In this way,
you will grow familiar with baseline stream
conditions and land and water uses, and
will be better able to identify changes over
time. You should choose the largest area
you feel comfortable assessing and ensure
that it has easy, safe, and legal access. The
area should have recognizable boundaries
that can be marked or identified on road
maps or U.S. Geological Survey topo-
graphic maps. This will help future volun-
teers continue the visual assessment in later
years and help the program coordinator
easily locate any problems that have been
identified.
Once you have identified the area to be
assessed, define it clearly in words (for
example, "Volunteer Creek from Bridge
over Highway One to confluence of Happy
-------
WATERSHED SURVEY METHODS
Creek at entrance to State Park"). Then,
either draw the outline and significant
features of the stream and its surroundings
on a blank sheet of paper or obtain a more
detailed map of the area, such as a plat,
road, or neighborhood map. This will serve
as the base map you will use to mark
stream obstructions, pollution sources, land
uses, litter, spills, or other problems identi-
fied during your visual assessment.
walk the stream (or the stream's problem
sites) at other times (see Tasks 4 and 5).
Task 2 I Determine when to survey
Because land and water uses can
change rapidly and because the natural
condition of the stream might change with
the seasons, it is best to visually assess the
stream or stream segment at least three
times a year. In areas with seasonal
changes, the best times to survey are:
• Early spring, before trees and shrubs
are in full leaf and when water levels
are generally high
• Late summer, when trees and shrubs
are in full leaf and when water levels
are generally low
• Late fall, when trees and shrubs have
dropped their leaves but before the
onset of freezing weather
In addition, you may wish to
spot-check potential problem areas more
frequently. These include construction sites,
combined sewer overflow discharges,
animal feedlots, or bridge/highway cross-
ings. If polluted runoff or failing septic
systems are suspected, schedule a survey
during or after heavy rainfall. If a stream is
diverted for irrigation purposes, surveys
during the summer season will identify
whether water withdrawals are affecting the
stream.
Again, it is important to survey the
stream at approximately the same time each
season to account for seasonal variations.
You might find it productive to drive
through the watershed once a year and to
Task 3 | Gather necessary equipment
In addition to the general and safety
equipment listed in Chapter 2, the follow-
ing equipment should be gathered before
beginning the visual assessment:
• Reference map such as road map or
USGS topographic map, to locate the
stream and the area to be assessed
• Base map to record land uses, land
characteristics, stream obstructions,
sources of pollution, and landmarks
• Field data sheet
• Additional blank paper, to draw
maps or take notes if needed
• Relevant information from back-
ground investigation (e.g., location
of NPDES outfalls, farms, aban-
doned mines, etc.)
Task 4 | Drive (or walk) the watershed
The purpose of driving (or walking) the
watershed is to get an overall picture of the
land that is drained by your stream or
stream segment. It will help you understand
what problems to expect in your stream,
and it will help you know where to look for
those problems.
As with all other monitoring activities,
you should undertake your watershed drive
or walk with at least one partner. If you are
driving, one of you should navigate with a
road map and mark up the base map and
field sheet with relevant discoveries while
the other partner drives. You might want to
pull over to make detailed observations,
particularly near stream crossings. Remem-
ber never to enter private property without
permission (see Safety Considerations,
Chapter 2).
As you drive or walk the watershed,
look for the following:
-------
WATERSHED SURVEY METHODS I 31
The "lay" of the land—become
aware of hills, valleys, and flat
terrain. Does any of this area periodi-
cally flood?
Bridges, dams, and channels—look
for evidence of how the community
has dealt with the stream and its
flood potential over the years. Are
portions of it running through
concrete channels? Is it dammed,
diverted, culverted, or straightened?
Where the road crosses the stream, is
there evidence of erosion and pollu-
tion beneath bridges? Is streamflow
obstructed by debris hung up beneath
bridges?
Activities in the watershed—look for
land use activities that might affect
your stream. In particular, look for
construction sites, parking lots,
manicured lawns, farming, cattle
crossings, mining, industrial and
sewage treatment plant discharges,
open dumps, and landfills. Look for
the outfalls you identified in your
background investigation. Also look
for forested land, healthy riparian
zones, undisturbed wetlands, wild-
life, and the presence of recreational
users of the stream such as swimmers
or people fishing. (Note that heavy
recreational use or large flocks of
birds might adversely affect the
quality of streams, ponds, lakes, and
wetlands.)
TaskS
Walk the stream
Where you have safe public access or
permission to enter the stream, stop driving
or walking the watershed and go down to
the stream. Use all of your senses to
observe the general water quality condition.
Does the stream smell? Is it strewn with
debris or covered with an oily sheen or
foam? Does it flow quickly or sluggishly?
Is it clear or turbid? Are the banks eroded?
Is there any vegetation along the banks? If
you see evidence of water quality problems
at a particular site, you might want to
investigate them in more detail Drive or
walk upstream as far as you can, and try to
identify where the water quality problem
begins.
Use your field data sheet to record your
findings. Always be as specific1 as possible
when noting your location and the water
conditions you are observing. Draw new
maps or take pictures if that will help you
remember what you are observing. Don't
be afraid to take too many notes or draw
too many pictures. You can always sort
through them later.
Take note of the positive conditions
and activities you see as well as the nega-
tive ones. This, too, will help you charac-
terize the stream and its watershed. Look
for such things as people swimming or
fishing in the stream; stable, naturally
vegetated banks; fish and waterfowl; or
other signs that the stream is healthy.
For more information on what to look
for in and around the stream, consult
Chapter 4 and, in particular, the Stream
Habitat Walk.
Task 6 I Review your maps/field data
^^~^^~^ sheets
The last step of the watershed survey's
visual assessment is to review the maps,
drawings, photos, and field data sheets you
have assembled for your stream or stream
segment. What is this information telling
you about problem sites, general stream
condition, potential for future degradation,
and the need for additional action? In most
cases you will find that you have put
together an interesting picture of your
stream. This picture might prompt addi-
tional monitoring or community activity, or
could urge your program coordinator to
bring potential problems to the attention of
water quality or public health agencies in
your area.
-------
WATERSHED SURVEY METHODS
When reviewing your data, be sure
maps are legible and properly identified,
photos have identifiable references, and
field data sheets are filled out completely
and accurately. Your program coordinator
might ask for your field data sheets, maps,
and other material and can probably help
interpret the findings of your watershed
survey.
For More Information on Your Watershed
EPA's Surf Your Watershed internet web site is a service
designed to help citizens locate, share, and use information on
their watershed or community. While you are conducting your
watershed survey, you might find its features of value. Surf
provides:
• Access to a large listing of protection efforts and volunteer
opportunities by watershed.
• Information on water resources, drinking water sources, land
use. population, wastewater dischargers, and water quality
conditions.
• Capabilities to generate maps of your watershed and
determine the latitude and longitude of specific sites within
it.
• Opportunity to share your watershed information with other
on-line groups through links with other pages and data-
bases.
You can reach Surf Your Watershed on the web at
mvw.epa.gov/surf.
References and Further Reading
Delaware Nature Education Center. 1996.
Delaware Stream Watch Guide. July.
Ely, E. 1994. Delineating a Watershed. The
Volunteer Monitor. 6(2):3.
Ely, E. 1994. Land-Use Surveys. The
Volunteer Monitor. 6(2): 19.
Gordon, N.D., T.A. McMahon, et al. 1992.
Stream Hydrology: An Introduction for
Ecologists. John Wiley and Sons.
Kerr, M. and V. Lee. 1992. Volunteer
Monitoring: Pipe Detectives Manual,
March 1992. Rhode Island Sea Grant,
University of Rhode Island, Coastal
Resources Center.
Kerr, M. and V. Lee. 1992. Volunteer
Monitoring: Shoreline Mapping Manual.
March. Rhode Island Sea Grant, Univer-
sity of Rhode Island, Coastal Resources
Center.
Maryland Save Our Streams. Watershed
Survey, Stream Survey, and Construction
Site Inventory (packets). Maryland Save
Our Streams, 258 Scotts Manor Drive,
Glen Burnie, MD 21061.
Trautmann, N. and E. Barnaba. 1994.
Aerial Photographs - A Useful Monitor-
ing Tool. The Volunteer Monitor.
6(2):17.
University of Rhode Island. 1990. Rhode
Island Watershed Watch: Shoreline
Survey Manual for Lakes, Rivers, and
Streams. Draft. June.
Yates, S. 1988. Adopting a Stream: A ,
Northwest Handbook. Adopt-A-Stream
Foundation. University of Washington
Press.
-------
•g
C. SI
f** s
13
oaaaoa o ooaaaaaoa- aaaa aaaaaoaa
E E
"4- ro
c 55
oonaaa o aooaooaaa1 noon aaaooaaa
CO
CO
UJ
CO
CO
<
<
I)
CO
tn
03
•ie
g
!
D.
0
2
'CD
I
CO
aoaoaa
ga
if
-
naaoaaaaO;
!
-E c S
aooD aaaaaoao
§
£
s
LU
CO
O
UJ
I
CO
cc
UJ
to
% <8
o
•S-
(0
Q)
E
Watershed na
f^nuntv:
£
(0
OT
•s
s
-p-
•5 . .2
» £ 'S.
E 13 «o
x .5> «
2 to 2-
, a 5 2
» < .£ to
0)
E
H-
I
I
o
CM t:
*• 5*
in s
£
g
0
s_
1
(0
£.
E
1
a
c
'2
t
m
c
'CO
CC
a
'§
S.
2
o
CO
a
1
a
0)
6
a
c
D
«
(U
O
D
'
3
>. -p
"
i i
D
CO
O
> s
o o
D
-------
T3
O
I
I
to
gS Dona anon aoooa aoooaaoaoaa
> CO
I
J8
5S
= "0
aaao oaaa aoooa aaaaaooa
ODD
55 >9
I
"8
«
1 ^
O
5
OT
CD
i
¥O
T3
CO
CO
CD
OT
onaa aoaa aoaao ooaooooaooa
.i
c
o
\s
o
to
o
o
o
ts
to
>, §
- o
1
S'.S'g
CD CD *-
S > ir
U= "rr CD
•g D) cB o > > t:
111!
C OQ GC m O O) CO
5 a
CD
D)
•i g
E 1
.9- E
c/) SI
CD
C
E
•o
CD
1 £
CO CO
J2 3
< a
CD
f~
O
ecreation
oc
_co
1
CD
0
£1
CD
TJ
CO
Biking/Off-ro;
2
•*-•
c
T3
Horseback ri
Boat ramp
2
D)
C
:E
CO
Q.
D3
_C
D)
0
CO
.CD
Swimming ar
CO
cfl
D)
Ic
LE
CO
CO
.g
'c
o
h.
Golf course
CO
Q.
Jl
'2
^?
Campground
C35
Power boatin
CD
O
-------
WATERSHED SURVEY METHODS
O i-
a> «
N *•
•a c
ss
a> >
r O
q>£
•2-6
Si
'
_ o
fiTS
(O
o —-
S
S
D.
a.
CO
0)
D)
a>
a>
.D
u
«o
u
•a
w
u
(0
.2
o
D)
(D
Q.
CO
a.
D)
CD
CO =
LO
CD
IS
O
r^
CO
D)
CD
CO
o
'co
CO
= =
co co
o £ E
Z CO CO
•«
c £ c
o 2 o
1 8 I
o
800
1_ »— *—
£ H ~
CD CD
CO CO
o
£
03
£•
§
co
CO
0)
o
c
g
8
j<:
co
XI
CD
CO
CD
Q.
•c
c _•:
O CJ>
E «,
l-l
*
05 15
^1
co 5
-O (Q
12
CD CO
- o
o a a o a
JB^
.. o t:
c z co
o
8 a
lii
__ _g
2> E «
to • »
M- ^£ J*
I||
m C P
521
§ 2 «
•g °«
Hi
o < <
aao aoaaoa
-------
WATERSHED SURVEY METHODS
o
I
s
•e
a
•^ -
pe
eter of
e
X
CO
1 a, ri
I S | •§
1= i "• o
a a a a
§>
>,
CO
a
O
a
O
o
jgs/sewage Q Chemical
CD
1
cn
O
CD
O
a
o
**
&
CO
1
8.
'5.
5
S CD
x: J3
0 |
O
CD <=
•i 5
° E
-C tO
o 2
a w
p^
•8
->
CD
CD
o
0-
CD
1
2
S
O>
•SL
1
CD
i
IF
CO
o
E"
CO
i
d)
•Si-
(D
«
Q a)
o
•a
O
CO CO O
05 S c
'a. >*-—
£ £ f
> CD •
CO
CO
E co
CO "fl)
•< -a
o o
co o & :§
c _ co o
co c TJ .<2
" §.== "2
CD S; °.
O .2 CD
COS
.g
aoaaao
co .9 o
5 Sill
«• I '15
T- Zi co 2 £
J
o
a
co
CD
a
io
t
11
II
on applies to a
CD
O)
CO
'§
Q
a
CO
JD
I
£
1o
if JE
I a
81 ! £ I
on of
This
Q In stream
0
1
_o
CD
.Q
8
0
Q
I I
CO O
II
M
S. o
CD
O>
O D) C TJ
•§ g 'a -g
lill
i
^ 2 aooaoaaaaa
-------
MACROINVERTEBRATES AND HABITAT I 37
-------
MACROINVERTEBRATES AND HABITAT
Biological monitoring, the study of
biological organisms and their
responses, is used to determine
environmental conditions. One type of
biological monitoring, the biological survey
or biosurvey, is described in this chapter.
The biosurvey involves collecting, process-
ing, and analyzing aquatic organisms to
determine the health of the biological
community in a stream.
In wadable streams (streams that can be
easily walked across, with water no deeper
than about thigh-high), the three most
common biological organisms studied are
fish, algae, and macroinvertebrates. This
manual discusses macroinvertebrate
monitoring only.
Macroinvertebrates are organisms that
are large (macro) enough to be seen with
the naked eye and lack a backbone (inverte-
brate). They inhabit all types of running
waters, from fast-flowing mountain streams
to slow-moving muddy rivers. Examples of
aquatic macroinvertebrates include insects
in their larval or nymph form, crayfish,
clams, snails, and worms (Fig. 4.1): Most
live part or most of their life cycle attached
to submerged rocks, logs, and vegetation.
Aquatic macroinvertebrates are good
indicators of stream quality because:
• They are affected by the physical,
chemical, and biological conditions
of the stream.
• They can't escape pollution and
show the effects of short- and long-
term pollution events.
• They may show the cumulative
impacts of pollution.
• They may show the impacts from
habitat loss not detected by tradi-
tional water quality assessments.
• They are a critical part of the
stream's food web.
• Some are very intolerant of pollution.
• They are relatively easy to sample
and identify.
The basic principle behind the study of
macroinvertebrates is that some are more
sensitive to pollution than others. There-
fore, if a stream site is inhabited by organ-
isms that can tolerate pollution—and the
more pollution-sensitive organisms are
missing—a pollution problem is likely.
For example, stonefly nymphs—
aquatic insects that are very sensitive to
most pollutants—cannot survive if a
stream's dissolved oxygen falls below a
certain level. If a biosurvey shows that no
stoneflies are present in a stream that used
to support them, a hypothesis might be that
dissolved oxygen has fallen to a point that
keeps stoneflies from reproducing—or has
killed them outright.
This brings up both the advantage and
disadvantage of the biosurvey. The advan-
tage of the biosurvey is that it tells us very
clearly when the stream ecosystem is
impaired, or "sick," due to pollution or
habitat loss. It is not difficult to realize that
a stream full of many kinds of crawling and
swimming "critters" is healthier than one
without much life. The disadvantage of the
biosurvey, on the other hand, is that it
cannot definitively tell us why certain types
of creatures are present or absent.
In this case, the absence of stoneflies
might indeed be due to low dissolved
oxygen. But is the stream under-oxygenated
because it flows too sluggishly or because
pollutants in the stream are damaging water
quality by using up the oxygen? The
absence of stoneflies might also be due to
other pollutants discharged by factories or
running off farmland, water temperatures
that are too high, habitat degradation such
as excess sand or silt on the stream bottom
that has ruined stonefly sheltering areas, or
other conditions. Thus a biosurvey should
be accompanied by an assessment of
habitat and water quality conditions in
order to help explain biosurvey results.
Habitat, as it relates to the biosurvey, is
defined as the space occupied by living
organisms. In a stream, habitat for macroin-
-------
MACROINVERTEBRATES AND HABITAT
Insects
Stoneflies (Order: Plecoptera)
Mayflies (Order: Ephemeroptera)
Caddisflies (Order: Trichoptera)
Dragonflies & Damselflies
(Order: Odonata)
Flies & Midges (Order: Diptera)
Water Bugs {Order: Hemiptera)
Dobsonfly (Order: Megaloptera)
Beetles (Order: Coleoptera)
Crustaceans
Crayfish & Freshwater shrimp
(Order: Decapoda)
Scud (Order: Amphipoda)
Isopod (Order: Isopoda)
Snails
Mussels & Clams
Worms
Leeches
Figure 4.1
Types of
macroinverte-
brates found in
streams
Many biosurvey
programs
include the
identification of
various macro-
invertebrates.
(Organisms are
not drawn to
scale)
Drawings f rom A Field Guide to Kentucky Rivers and Streams
-------
MACROINVERTEBRATES AND HABITAT
vertebrates includes the rocks and sedi-
ments of the stream bottom, the plants in
and around the stream, leaf litter and other
decomposing organic material that falls into
the stream, and submerged logs, sticks, and
woody debris. Macroinvertebrates need the
shelter and food these habitats provide and
tend to congregate in areas that provide the
best shelter, the most food, and the most
dissolved oxygen. A habitat survey exam-
ines these aspects and rates the stream
according to their quality. This chapter
includes both simple and intensive habitat
surveys volunteers can conduct.
Monitoring for water quality conditions
such as low dissolved oxygen, temperature,
nutrients, and pH helps identify which
pollutants are responsible for impacts to a
stream. Water quality monitoring is dis-
cussed in Chapter 5.
Uses of the Biosurvey and
Habitat Assessment
The information provided by
biosurveys and habitat assessments can be
used for many purposes.
• To screen for impairment.
Biosurveys can be used to identify
problem sites along a stream. A
habitat assessment can help deter-
mine whether the problem is due, at
least in part, to a habitat limitation
such as poor bank conditions.
• To identify the impact of pollution
and of pollution control activities.
Because macroinvertebrates are
stationary and are sensitive to
different degrees of pollution,
changes in their abundance and
variety vividly illustrate the impact
pollution is having on the stream.
Loss of macroinvertebrates in the
stream, or of trees along the stream
bank, are environmental impacts that
a wide segment of society can relate
to. Similarly, when a pollution
control activity takes place—say, a
fence is built to keep cows out of the
stream—a biosurvey may show that
the sensitive macroinvertebrates
have returned and a habitat assess-
ment might find that the formerly
eroded stream banks have recovered.
To determine the severity of the
pollution problem and to rank
stream sites. To use biological data
properly, water resource analysts
generally compare the results from
the stream sites under study to those
of sites in ideal or nearly ideal
condition (called a reference condi-
tion). Individual stream sites can
then be ranked from best to worst,
and priorities can be set for their
improvement.
To determine support of aquatic life
uses. All states designate their
waters for certain specific uses, such
as swimming or as cold water
fishery. States establish specific
standards (limits on pollutants)
identifying what concentrations of
chemical pollutants are allowable if
designated stream uses are to be
maintained. Increasingly, states are
also developing biological criteria—
essentially, statements of what
biological conditions should be in
various types of streams throughout
the state. States are required by the
Clean Water Act to report on those
waters which do not support their
designated uses.
Biological surveys directly
examine the aquatic organisms in
streams and the stressors that affect
them. Therefore, these surveys are
ideal tools to use in determining
whether a stream's designated
aquatic life uses are supported.
To identify water quality trends. In
any given site, biological data can be
used to identify water quality trends
(increasing or decreasing) over
several years.
-------
MAGROINVERTEBRATES AND HABITAT
Designing a Biosurvey Program
In most cases, this manual recommends
that local aquatic biologists assist in the
development of volunteer biological
monitoring programs. This is because the
types of habitats and organisms in streams
vary widely with geography and climate.
Tools as basic as macroinvertebrate identi-
fication keys might need to be adapted to
local conditions.
Many volunteer monitoring programs
rely for assistance on aquatic biologists
working for state water-quality or natural
resource agencies. Others are assisted by
university personnel, hire their own expert
staff, or contract out for consulting services.
Whatever the source of expertise, profes-
sional guidance is essential for creating a
successful biosurvey program. This
manual strongly recommends a close
level of coordination with state or local
agencies that might use the data volun-
teers collect.
Monitoring approaches—and the level
of professional guidance and assistance
needed—clearly vary with the goals and
resources of individual volunteer groups.
Therefore, this manual presents three
different approaches or tiers to biological
monitoring.
• Stream Habitat Walk (detailed in
section 4.1) is for groups focused
primarily on educating volunteers
about their streams and for identify-
ing severe pollution problems.
Volunteers conduct simple visual
assessments of habitat to gain a
greater appreciation of local stream
ecology.
It is based on a protocol known
as Streamwalk developed by the
EPA Region 10 Office in Seattle,
Washington, and is widely used by
volunteers throughout the Pacific
Northwest.
• Streamside Biosurvey (detailed in
section 4.2) trains volunteers to
collect macroinvertebrates and
identify them to order level (stonefly,
mayfly, caddisfly, etc.) in the field.
Monitors evaluate the macroinverte-
brate community structure by sorting
specimens into three general sensitiv-
ity categories. In addition, volunteers
characterize habitat by conducting a
modified Stream Habitat Walk.
This tier is based on, a protocol
developed by the Ohio Department
of Natural Resources and adapted by
the Izaak Walton League of America.
It has been used by volunteer moni-
tors nationwide, including programs
in Ohio, Tennessee, Georgia, Vir-
ginia, Kentucky, Illinois, and West
Virginia.
Intensive Biosurvey (detailed in
section 4.3) requires that volunteers
work under the supervision of
professional aquatic biologists.
Volunteers undergo formal training
and conduct quality-controlled
sampling and analysis. Using micro-
scopes in a laboratory setting,
macroinvertebrates are identified to
Figure 4.2
^•••MBBBi
Taxonomic
classification
system
Depending on
the program,
volunteers might
be asked to
identify macroin-
vertebrates to
the order level
in the field or to
the family level
if using micro-
scopes in the
laboratory.
Taxonomic Classification
Scientists have developed a system for classifying all living
creatures based on shared characteristics (taxonomic classifica-
tion). It is a tiered system that begins on a large scale (i.e., Animal
Kingdom/Plant Kingdom) and works its way down to the level of
individual species. To illustrate, the burrowing mayfly is classified
as follows.
Kingdom: Animal
Phylum: Arthropoda
Class: Insecta
Order: Ephemeroptera :
Family: Ephemerida
Genus: Hexagenia
Species: limbata
-------
MACRO1NVERTEBRATES AND HABITAT
Table 4.1
Tiered frame-
work for
volunteer
biological
monitoring
programs
Program
designers might
choose simple
or complex
approaches
according to
program goals
and resources.
the family level (what types of
stoneflies, mayflies, caddisflies, etc.).
Analytical techniques are subse-
quently applied to the data to draw
conclusions about the biological
health of the sampled site. This
rigorous biosurvey approach results
in data that can yield information on
subtle stream impacts and trends.
Based primarily on EPA's Rapid
Bioassessment Protocols, this
approach has been adapted by Mary-
land Save Our Streams, the River
Watch Network and other groups.
We have modified the approaches used
by other groups to add to their capabilities
or to make them more generally applicable
to all U.S. streams. Individual programs
might choose to start with the simplest,
least resource-intensive approach and work
their way toward increasing complexity as
resources, expertise, and volunteer interest
allow. However, groups might decide to
begin with a more complex approach that
better suits their program goals. Table 4.1
illustrates some of the key differences in
the three biological monitoring approaches
discussed in this manual.
Protocol Elements Stream Habitat Walk Streamside Biosurvey Intensive Biosurvey
Program Objectives
Complexity of
Approach
Resource Investment
Training
• Education/public awareness
• Gross problem identification/
screening
• Simple visual assessment of
habitat and physical charac-
teristics
• Basic observational biological
data recording general
abundance/variety of
macroinvertebrates and
presence or absence of
macrophytes, algae, and fish
• Scientific personnel assist in
project design, preparation of
documentation, and orienta-
tion of volunteers
• Minimal equipment (maps,
manuals, forms)
• Primarily self-instructional
using manuals/documentation
(some training is desirable)
• Education/public awareness
• Problem identification/
screening
• Preliminary ranking of sites
for further study
• Visual assessment of habitat
and physical characteristics
• In-stream biota collected and
evaluated at streamside for
relative sensitivity/tolerance
and identified to order/family
level
• Scientific personnel involved
in project design, preparation
of documentation, training,
and supervision of biosurveys
• Sampling gear, maps,
manuals, forms, references
• Periodic workshops and
streamside training sessions
• Education/public awareness
• Problem identification/
screening
• Assessing severity of
problems
• Ranking of sites for manage-
ment action
• Comprehensive habitat and
physical assessment
• Instream biota collected,
preserved, and identified in
lab to family level (multimetric
approach)
• Reference sites or conditions
identified
• Scientific personnel active in
all levels and mandatory for
assessment and data
interpretation
• Laboratory and storage
facilities in addition to other
equipment
• Voucher and reference
collections required
• Formal lab and field training
with experienced team
leaders before all assess-
ments
-------
MACROINVERTEBRATES AND HABITAT
4.1
Stream Habitat Walk
The Stream Habitat Walk is an easy-to-
use approach for identifying and assessing
the elements of a stream's habitat. It is
based on a simple protocol known as
Streamwalk, developed by EPA's Regional
Office in Seattle, Washington and consists
primarily of visual observation of stream
habitat characteristics, wildlife present, and
gross physical attributes. A simple in-
stream macroinvertebrate evaluation can
also be performed. This approach requires
little in the way of equipment and training.
The Stream Habitat Walk is most
useful as:
• A screening tool to identify severe
water quality problems
• A vehicle for learning about stream
ecosystems and environmental
stewardship
Because the Stream Habitat Walk is not
scientifically rigorous, data from this
approach are less likely to be used by state
and local water quality management
agencies than are data from other biological
monitoring approaches. However, the
Stream Habitat Walk's ease of use, adapt-
ability, and low cost make it a highly
attractive approach for many programs
whose primary focus is public awareness
and citizen involvement.
Step 1—Prepare for the Walk
TASK1 | Schedule your Habitat Walk
To provide data that accurately charac-
terize your stream and can be used to
document general trends in your area, you
should walk the same site at least three
times a year, during different seasons. It is
usually best to visit your site in early
spring, late summer, and fall if you live in a
part of the country that experiences sea-
sonal variations in leaf cover, vegetation
growth, and water flow. It is a good idea to
check with a local aquatic biologist for
assistance in determining the best times to
schedule monitoring. For purposes of
accuracy and consistency, it is'best to
monitor the same site from year to year and
at the same time of the year (e.g., in the
spring and, more specifically, in the same
month).
TASK 2
Obtain a U.S. Geological
Survey (USGS) topographic
map of your area
One of the most valuable tools for
conducting stream monitoring work is a
U.S. Geological Survey (USGS) topo-
graphic map. These "topo" maps display
many important features of the landscape
including elevations, waterways, roads, and
buildings. They are critical tools for
defining the watershed of your study
stream. (See Chapter 3 for a discussion of
topographic maps.)
TASKS
Select and mark the Habitat
Walk location(s)
Choosing the location for stream
monitoring is a task defined by the goals of
your individual program. Program manag-
ers may select sites themselves or in
collaboration with local or state water
quality personnel. Other programs allow
their volunteers to choose the site based on
their personal interests. (See Chapter 2 for
a discussion on choosing monitoring
locations.) If a Watershed Survey is
conducted (see Chapter 3), this information
should play a role in deciding which areas
are the best candidates for the Stream
Habitat Walk.
Once a monitoring site is chosen, it
should be marked on the topo map. This
will document the location and serve as a
record in case future volunteers or data
users need to find the site.
-------
MACROINVERTEBRATES AND HABITAT
TASK 4
Become familiar with safety
procedures
Volunteers must always keep safety in
mind while conducting any stream monitor-
ing activity. Provide all Stream Habitat
Walk participants with a list of safety do's
and don'ts and have them review this list
thoroughly. Chapter 3 covers several
important safety concerns that should be
incorporated into a stream monitoring
program. Remember, volunteer safety is
more important than the data. Some re-
minders include:
• Let someone know where you're
going and when you expect to return.
Make sure you have an "in case of
emergency" phone number with you
before leaving for the field.
• Do not cross streams in high flows.
• Never go into the field alone; always
work in teams of at least two people.
• If for any reason you feel unsafe, do
not attempt to monitor on that day.
TASKS
Gather equipment and tools
for the Habitat Walk
There is nothing more frustrating than
arriving at a field monitoring site and not
having all your equipment and supplies.
Providing volunteers with a checklist of
necessary items will help keep them
organized. In addition to the basic equip-
ment listed in Chapter 2, you will need the
following for the Stream Habitat Walk.
For locating the site
• U.S. Geological Survey (USGS)
topographic map of the stream area
(supplemented by regular street map
if needed)
For recording observations
• Stream Habitat Walk field data sheet
For marking-off the stream stretch of study
• Tape measure, string, or twine (25
yards)
For working in and around the stream
• Thermometer for measuring water
temperature (Scientific,supply
houses sell armored thermometers
that are best suited for this purpose,
although you can obtain a good
thermometer from an aquarium store.
Some thermometers need to be
calibrated before use. See Chapter 5
for instruction on calibrating and
using thermometers.)
• Watch with a second hand or a
stopwatch
For observing macroinvertebrates (op-
tional)
• A bucket
• A shallow white pan. (Alternatives:
white plastic plate or the bottom of a
white plastic detergent jug)
• Tweezers or soft brush
• Ice cube trays (for sorting macroin-
vertebrates)
• Magnifying glass
TASK 6
Become familiar with the
Stream Habitat Walk field
data sheet and the defini-
tions of its elements
It is important to become familiar with
the Stream Habitat Walk field data sheet
and its instructions before you begin your
Stream Habitat Walk. If you are unclear
about any instructions when you are
conducting your Walk, just leave that space
blank and keep going. You might wish to
contact your volunteer program coordinator
for further explanation after you have
completed your Walk.
At the end of this section is a sample
field data sheet. You might find it necessary
to modify this sheet slightly to better meet
the needs of your volunteers, your ecologi-
cal region, and your program. When you
fill out your field data sheet, base your
responses on your best judgment of condi-
-------
MACROINVERTEBRATES AND HABITAT
tions in a stretch of stream that includes
about 50 yards both upstream and down-
stream of the place where you are standing.
If you identify features and problems
beyond your chosen 100-yard length, feel
free to note them on your map and form.
You might want to conduct additional
Walks in the area where those features are
found.
Instructions on how to fill out the field
data sheet are included right on the form.
They are also covered in an expanded
format, with illustrations, in this text.
Although many of the required measures
are relatively self-explanatory, it might be a
good idea to make copies of these instruc-
tions for all volunteer teams to take into the
field as an additional training tool.
Step 2—Delineate and sketch
your site
TASK1
Delineate the site
Using your tape measure or 25 yards of
string or twine, measure off four 25-yard
lengths alongside the stream for a total of
100 yards. Start from a point of reference
such as a tree, large rock, or bend in the
stream.
TASK 2 I Sketch your site on the field
• data sheet
On the field data sheet, sketch the 100-
yard section of stream. (Fig, 4.3). Drawing
the map will familiarize you with the
terrain and stream features and provide you
and other volunteers with a visual record of
your habitat walk. You should walk the
100-yard length from at least one bank.
On your sketch, note features such as
riffles, runs, pools, ditches, wetlands, dams,
riprap, outfalls, tributaries, landscape
features, jogging paths, vegetation, and
roads. Use your topo map or a compass to
determine which direction is north and
mark it on your sketch. If you see important
features outside your 100-yard length of
stream, mark them on your sketch but note
that they are outside the stream reach.
Remember to use pencil or waterproof ink
when drawing your map or filling out the
field data sheets because regular ink will
run if wet.
Select a 25-yard section of the site.
You will be filling out your field data sheet
for this section only. Mark the section on
the sketch. If you want to conduct multiple
walks, choose another 25-yard section or
move to an entirely different location. Even
though you will only be completing the
data forms for the 25 yard reach, it is
important to sketch the full 100-yard
section so that you can document the
stream features surrounding the evaluated
reach.
TASKS
Complete the top portion of
your field data sheet
Include stream name, date, and county
(or appropriate local designation) of your
site, and describe its location as precisely
as possible. It is best to stand at or near a
permanent marker such as a bridge, abut-
ment, or road. Remember, you or another
volunteer will be coming back to the same
spot again and again, so be as specific as
you can. Some programs might ask you for
the latitude and longitude of your location;
others might ask for a map reference
number or other site identifier.
Latitude and longitude information is
critical for mapping and for many data
management programs. It is also required if
the data is to be entered in USEPA's
STOrage and RETrieval System
(STORET) or used in a Geographical
Information System (GIS).
An easy way to determine latitude and
longitude is to use a global positioning
system (GPS), a hand-held tool that looks
like a calculator. GPS units receive signals
form orbiting satellites and then use the
information from the satellites to calculate
the lat/long coordinates of the user. In
-------
Figure 4.3
Example of a
stream sketch
Volunteers
should note
important
stream features
on their sketch
including riffles
and pools.
general, these tools are accurate up to 15
meters. GPS units are relatively inexpen-
sive and can be purchased from scientific
supply houses and many camping or
outdoor stores. Many government agencies
are using GPS and might be able to loan a
system to your program. Latitude and
longitude can also be calculated manually
using a USGS topographical map and a
ruler (See Appendix C).
-------
MACROINVERTEBRATES AND HABITAT
Step 3—Conduct the Stream
Habitat Walk
Detailed instructions for performing the
Stream Habitat Walk begin on page 48 of
this section.
TASK1
Complete the habitat charac-
terization components of the
walk for the 25-yard section
of stream: the "In-Stream
Characteristics," "Stream
Bank and Channel Charac-
teristics," and "Local
Watershed Characteristics"
sections of the field data
sheet
These elements involve making obser-
vations about the stream itself as well as the
riparian zone and immediate watershed.
TASK 2
Complete the "Visual
Biological Survey" section of
the field data sheet
This involves simple visual observa-
tions of the presence or absence of wildlife
and obvious aquatic life in the stream,
including fish, aquatic plants, and algae.
TASKS
Complete the "Macroinverte-
brate Survey" section of the
field data sheet
This is optional and serves as an
introduction to the types of life that inhabit
some of the microhabitats of the stream—
the spaces under and on rocks and in and on
twigs and leaves. To conduct this survey,
you will need to select the method(s) that
best suits your stream. Use the rock-
rubbing method in streams with riffles, or
use the stick-picking method if your stream
does not have riffles. Clumps of submerged
leaves may be present in either type of
stream and are often an important micro-
habitat for macroinvertebrates. You may
choose to sort through these leaf packs in
addition to rock-rubbing or stick-picking.
You will also need some specific
equipment (a bucket, tweezers, picnic
plate, etc.). Be sure to dress appropriately
because you'll probably get wet.
Remember to return the organisms to
the stream when you finish the macroinver-
tebrate survey. Then, check to make sure
your field data sheet has been completed as
fully as possible.
Step 4—Check data forms for
completeness and return forms
to program coordinator
After completing the habitat character-
ization and biological survey, make sure
you have completed the field data sheet to
the extent possible and that the recorded
data are legible. If you are not able to
determine how to answer a question on the
field data sheet, just leave the space blank.
If you leave a space blank, indicate that it
is because you are not able to answer the
question (e.g., write "not able to answer" or
"does not apply" in the space).
Upon completion of the Stream Habitat
Walk, present a copy of the field data sheet
to your volunteer program coordinator.
You may want to keep a copy of the field
data sheet, and other appropriate data, for
your own records and to evaluate any
future discrepancies in the data. If you
have identified an urgent problem, such
as leaking drums of chemicals, foul
odors, or fish kills, contact your pro-
gram coordinator or the agency with
whom you are working as soon as
possible.
-------
MACROINVERTEBRATES AND HABITAT
Figure 4.4
Overview and
cross sections
of a pool, riffle,
and run
Varying flows
and depths
create a variety
of habitats for
macroinverte-
brates.
Instructions for
completing the
Stream Habitat Walk
data sheet
For ease of use, the following num-
bered instructions correspond to the num-
bers on the field data sheet.
In-stream Characteristics
1. Pools, riffles, and runs. A mixture of
flows and depths creates a variety of
habitats to support fish and inverte-
brate life. Pools are deep with slow
water. Riffles are shallow with fast,
turbulent water running over rocks.
Runs are deep with fast water and
little or no turbulence.
2. Stream bottom (substrate) is the
material on the stream bottom.
Identify what substrate types are
present. Substrate types include:
• Silt/clay/mud. This substrate has a
sticky, cohesive feeling. The
particles are fine. The spaces
between the particles hold a lot of
water, making the sediments
behave like ooze.
• Sand (up to 0.1 inch). A sandy
bottom is made up of tiny, gritty
particles of rock that are smaller
than gravel but coarser than silt
(gritty, up to pea size).
• Gravel (0.1-2 inches). A gravel
bottom is made up of stones
vs
POOL
-------
MACROINVERTEBRATES AND HABITAT
ranging from tiny quarter-inch
pebbles to rocks of about 2 inches
(fine gravel - pea size to marble
size; coarse gravel - marble to
tennis ball size).
• Cobbles (2-10 inches). Most
rocks on this type of stream
bottom are between 2 and 10
inches (between a tennis ball and
a basketball).
• Boulders (greater than 10
inches). Most of the rocks on the
bottom are greater than 10 inches
(between a basketball and a car in
size).
• Bedrock. This kind of stream
bottom is solid rock (or rocks
bigger than a car).
3. Embeddedness is the extent to which
rocks (gravel, cobbles, and boulders)
are sunken into the silt, sand, or mud
of the stream bottom (Fig. 4.5).
Generally, the more rocks are
embedded, the less rock surface or
space between rocks is available as
habitat for aquatic macroinverte-
brates and for fish spawning.
Excessive silty runoff from erosion
can increase a stream's embedded-
ness. To estimate embeddedness,
observe the amount of silt or finer
sediments overlying, in between, and
surrounding the rocks.
4. Presence of logs or woody debris
(not twigs and leaves) in stream can
slow or divert water to provide
important fish habitat such as pools
and hiding places. Mark the box that
describes the general amount of
woody debris in the stream.
5. Naturally occurring organic material
in stream. This material includes
leaves and twigs. Mark the box that
describes the general amount of
organic matter in the stream.
Figure 4.5
6. Water appearance can be a physical
indicator of water pollution.
• Clear - colorless, transparent
• Milky - cloudy-white or grey, not
transparent; might be natural or
due to pollution
• Foamy - might be natural or due
to pollution, generally detergents
or nutrients (foam that is several
inches high and does not brush
apart easily is generally due to
some sort of pollution)
• Turbid - cloudy brown due to
suspended silt or organic material
• Dark brown - might indicate that
acids are being released into the
stream due to decaying plants
• Oily sheen - multicolored reflec-
tion might indicate oil floating in
the stream, although some sheens
are natural
A representa-
tion of a rocky-
bottom stream
becoming
embedded with
sand and silt
As silt settles on
the streambed,
spaces between
the rocks are
filled in and the
stream be-
comes more
embedded.
-------
• Orange - might indicate acid
drainage
• Green - might indicate excess
nutrients being released into the
stream
7. Water odor can be a physical indica-
tor of water pollution
• No smell or a natural odor
• Sewage - might indicate the
release of human waste material
• Chlorine - might indicate over-
chlorinated sewage treatment/
water treatment plant or swim-
ming pool discharges
• Fishy - might indicate the pres-
ence of excessive algal growth or
dead fish
• Rotten eggs - might indicate
sewage pollution (the presence of
methane from anaerobic condi-
tions)
8. Water temperature can be particu-
larly important for determining the
suitability of the stream as aquatic
habitat for some species of fish and
macroinvertebrates that have distinct
temperature requirements. Tempera-
ture also has a direct effect on the
amount of dissolved oxygen avail-
able to the aquatic organisms.
Measure temperature by submerging
a thermometer for at least 2 minutes
in a typical stream run. Repeat once
and average the results.
Stream Bank and Channel
Characteristics
9. Depth of runs and pools should be
determined by estimating the vertical
distance from the surface to the
stream bottom at a representative
depth at each of the two habitats.
10. The width of the stream channel can
be determined by estimating the
width of the streambed that is
covered by water from bank to bank.
If it varies widely, estimate an
average width.
11. Stream velocity can have a direct
influence on the health, variety, and
abundance of aquatic communities.
If water flows too quickly, organisms
might be unable to maintain their
hold on rocks and vegetation and be
washed downstream; if water flows
too slowly, it might provide insuffi-
cient aeration for species needing
high levels of dissolved oxygen.
Stream velocity can be affected by
dams, channelization, terrain, runoff,
and other factors. To measure stream
velocity, mark off a 20-foot section
of stream run and measure the time it
takes a stick, leaf, or other floating
biodegradable object to float the 10
feet. Repeat at least three times and
pick the average time. Divide the
distance (20 feet) by the average time
(seconds) to determine the velocity
in feet per second. (See Chapter 5,
Section 5.1 on flow for a more in-
depth discussion of using a float to
estimate velocity.)
12. The shape of the stream bank, the
extent of artificial modifications, and
the shape of the stream channel are, -
determined by standing at the
downstream end of the 25-yard
section and looking upstream.
(a) The shape of the stream bank
(Fig. 4.6) may include.
• Vertical or undercut bank - a
bank that rises vertically or
overhangs the stream. This
type of bank generally pro-
vides good cover for
macroinvertebrates and fish
and is resistant to erosion. If
seriously undercut, it might be
vulnerable to collapse.
• Steeply sloping - a bank that
slopes at more than a 30
-------
MACROINVERTEBRATES AND HABITAT
degree angle. This type of
bank is very vulnerable to
erosion.
• Gradual sloping - a bank that
has a slope of 30 degrees or
less. Although this type of
stream bank is highly resistant
to erosion, it does not provide
much streamside cover.
(b) Artificial bank modifications
include all structural changes to
the stream bank such as riprap
(broken rock, cobbles, or boulders
placed on earth surfaces such as
the face of a dam or the bank of a
stream, for protection against the
action of the water) and bulk-
heads. Determine the approximate
percentage of each bank (both the
left and right) that is artificially
covered by the placement of
rocks, wood, or concrete.
(c) The shape of the stream channel
can be described as narrow (less
than 6 feet wide from bank to
bank), wide (more than 6 feet
from bank to bank), shallow (less
than 3 feet deep from the stream
substrate to the top of the banks)
or deep (more than 3 feet from the
stream substrate to the top of the
banks). Choose the category that
best describes the channel.
• Narrow, deep
• Narrow, shallow
• Wide, deep
• Wide, shallow
13. Streamside cover information helps
determine the quality and extent of
the stream's riparian zone. This
information is important at the
stream bank itself and for a distance
away from the stream bank. For
example, trees, bushes, and tall grass
can contribute shade and cover for
fish and wildlife and can provide the
stream with needed organic material
such as leaves and twigs. Lawns
indicate that the stream's riparian
zone has been altered, that pesticides
and grass clippings are a possible
problem, and that little habitat and
shading are available. Bare soil and
pavement might indicate problems
with erosion and runoff. Looking
upstream, provide this information
for the left and right banks of the
stream.
• Evergreen trees (conifers) - cone-
bearing trees that do not lose their
leaves in winter.
• Hardwood trees (deciduous) - in
general, trees that shed their
leaves at the end of the growing
season. :
Figure 4.6
^^MMMHM
Types of
streambank
shapes
Undercut banks
provide good
cover for fish
and macroinver-
teb rates.
-------
MACROINVERTEBRATES AND HABITAT
• Bushes, shrubs - conifers or
deciduous bushes less than 15 feet
high.
• Tall grass, ferns, etc. - includes
tall natural grasses, ferns, vines,
and m losses.
• Lawn - cultivated and maintained
short grass.
• Boulders - rocks larger than 10
inches.
• Gravel/cobbles/sand - rocks
smaller than 10 inches; sand.
• Bare soil
• Pavement, structure - any struc-
tures or paved areas, including
paths, roads, bridges, houses, etc.
14. Stream shading is a measurement of
the extent to which the stream itself
is overhung and shaded by the cover
identified in 13 above. This shade (or
overhead canopy) provides several
important functions in the stream
habitat. The canopy cools the water;
offers habitat, protection, and refuge
for aquatic organisms; and provides a
direct source of beneficial organic
matter and insects to the stream.
Determine the extent to which vege-
tation shades the stream at your site.
15. General conditions of the stream
bank and stream channel, and other
conditions that might be affecting the
stream are determined by standing at
the downstream end of the 25-yard
site and looking upstream. Provide
observations for the right and left
banks of the stream.
(a) Stream bank conditions that
might be affecting the stream.
• Natural plant cover degraded.
Note whether streamside
vegetation is trampled or
missing or has been replaced
by landscaping, cultivation, or
pavement. (These conditions
could lead to erosion.)
• Banks collapsed/eroded. Note
whether banks or parts of
banks have been washed away
or worn down. (These condi-
tions could limit habitats in the
area.)
• Garbage/junk adjacent to the
stream. Note the presence of
litter, tires, appliances, car
bodies, shopping carts, and
garbage dumps.
• Foam or sheen on bank. Note
whether there is foam or an
oily sheen on the stream bank.
Sheen may indicate an oil spill
or leak, and foam may indicate
the presence of detergent.
(b) Stream channel conditions that
might be affecting the stream.
• Mud/silt/sand on bottom/
entering stream. Excessive
mud or silt can interfere with
the ability of fish to sight
potential prey. It can clog fish
gills and smother fish eggs in
spawning areas in the stream
bottom. It can be an indication
of poor construction practices,
urban area runoff, silviculture
(forestry-related activities), or
agriculture in the watershed. It
can also be a normal condition
in slow- moving, muddy-
bottom streams.
• Garbage or junk in stream.
Note the presence of litter,
tires, appliances, car bodies,
shopping carts, and garbage.
(c) Other general conditions that
might be affecting the stream.
• Yard waste (e.g., grass
clippings). Is there evidence
that grass clippings, cut
branches, and other types of
yard waste have been dumped
into the stream?
-------
MACROINVERTEBRATES AND HABITAT
• Livestock in or with unre-
stricted access to stream. Are
livestock present, or is there
an obvious path that livestock
use to get to the water from
adjacent fields? Is there
streamside degradation caused
by livestock?
• Actively discharging pipes.
. Are there pipes with visible
openings discharging fluids or
water into the stream? Note
such pipes even though you
may not be able to tell where
they come from or what they
are discharging.
• Other pipes. Are there pipes
near or entering the stream?
Note such pipes even if you
cannot find an opening or see
matter being discharged.
• Ditches. Are there ditches
draining the surrounding land
and leading into the stream?
Local Watershed Characteristics
16. Adjacent land uses can potentially
have a great impact on the quality
and state of the stream and riparian
areas. Determine the land uses, based
on your own judgment of the activi-
ties in the watershed surrounding
your site within a quarter of a mile.
Enter a "1" if a land use is present
and a "2" if it is clearly having a
negative impact on the stream.
Visual Biological Survey
17. Wildlife in or around the stream
might indicate that the stream and its
adjacent area are of sufficient quality
to support animals with food, water,
and habitat. Look for signs of frogs,
turtles, snakes, ducks, deer, beaver,
etc.
18. Are. fish present in the stream? Fish
can indicate that the stream is of
sufficient quality for other organ-
isms. Indicate the average size and
note any visible barriers to the
movement of fish—obstructions that
would keep fish from moving freely
upstream or downstream.
19. Aquatic plants provide food and
cover for aquatic organisms. They
also might provide very general
indications of stream quality. For
example, streams that are overgrown
with plants could be over-enriched
by nutrients. Streams devoid of
plants could be affected by extreme
acidity or toxic pollutants. Aquatic
plants may also be an indicator of
stream velocity because plants
cannot take root in fast-flowing
streams.
20. Algae are simple plants that do not
grow true roots, stems, or leaves and
that mainly live in water, providing
food for the food chain. Algae may
grow on rocks, twigs, or other
submerged materials, or float on the
surface of the water. It naturally
occurs in green and brown colors.
Excessive algal growth may indicate
excessive nutrients (organic matter
or a pollutant such as fertilizer) in
the stream.
Macroinvertebrate Survey
(optional)
21. Macroinvertebrates are organisms
such as clams, mussels, snails,
worms, crayfish, and larval insects
that lack a backbone and can be seen
with the naked eye. To locate
macroinvertebrates in the stream, use
one or more of the following meth-
ods.
(a) Rock-rubbing method. (Use this
method in streams with riffle
areas and rocky bottoms.)
-------
MACROINVERTEBRATES AND HABITAT
• Remove several rocks from
within a riffle area of your
stream site (e.g., randomly
pick one rock from each side
of the stream, one rock from
the middle, and one rock from
in between). Try to choose
rocks that are submerged
during normal flow condi-
tions. Each rock should be
about 4-6 inches in diameter
and should be easily moved
(not embedded).
• Either inspect the rock's
surface for any living organ-
isms or place the rock in a
light-colored bucket or
shallow pan, add some stream
water, and brush the rock with
a soft brush or your hands. Try
to dislodge the foreign par-
ticles from the rock's surface.
Also look for clumps of gravel
or leaves stuck to the rock.
These clumps may be
caddisfly houses and should
be dislodged as well.
(b) Stick-picking method. (Use this
method in streams without riffles
or without a rock bottom.)
• Collect several sticks (ap-
proximately one inch in
diameter and relatively short)
from inside the stream site,
and place them in a bucket
filled with stream water.
Select partially decomposed
objects that have soft, pulpy
wood and a lot of crevices and
are found in the flowing water,
not buried in the bottom.
• Fill the shallow pan with water
from the stream and remove
one of the sticks from the
bucket. While examining the
stick, make sure you hold it
over the pan so no organisms
are lost. Remember that the
organisms will have sought
shelter, and they could be
hiding in loose bark or crev-
ices. After examining the
sticks, break up the bark and
woody material. Examine each
stick carefully. Using tweezers
or a soft brush, carefully
remove anything that re-
sembles a living organism and
place it in the pan. Also
examine the bucket contents in
case anything has fallen off
the sticks.
(c) Leaf-pack sorting method. (This
method can be used in streams
with or without a rock bottom.)
• Remove several handfuls of
submerged leaves from the
stream and place them into a
bucket. Remove the leaves
one at a time and look closely
for the presence of insects.
Using tweezers or a soft brush,
carefully remove anything that
resembles a living organism
and place it in a pan contain-
ing stream water. Also
examine the bucket contents to
see if anything has fallen off
the leaves.
22. Note whether you have found any
macroinvertebrates using one of the
above methods.
23. After collecting macroinvertebrates
using any of the above methods,
examine the types of organisms by
gross morphological features (e.g.,
snails or worm-like). Use a magnify-
ing glass to observe the organisms in
water so you can clearly see the legs,
gills, and tails. Note the relative
abundance of each type on the field
data sheet.
Many types of macroinverte-
brates can be found in a healthy
-------
MACROINVERTEBRATES AND HABITAT
stream. Because different species can
tolerate different levels of pollution,
observing the variety and abundance
of macroinvertebrates can give you a
sense of the stream's health. For
example, if pollution-tolerant organ-
isms are plentiful and pollution-
intolerant ones are found only
occasionally, this might indicate a
problem in the stream. Types of
organisms you find may include:
• Worm-like organisms (like worms
and leeches) either adhere to
rocks or sticks or move slowly.
They are generally tolerant of
pollution.
• Crayfish look like lobsters or
shrimp. They are generally
somewhat tolerant of pollution.
• Snail-like organisms include
snails and clam-like organisms.
They range from somewhat
tolerant of pollution to somewhat
intolerant.
• Insects include a wide variety of
organisms that generally have
distinct legs, head, bodies, and
tails and often move quickly over
rocks or sticks. They come in
many sizes and shapes as well as
a wide range of pollution-toler-
ance levels.
When finished, return all the organ-
isms to the stream.
-------
MACROINVERTEBRATES AND HABITAT
STREAM HABITAT WALK
Stream Name:
County:
Investigators:
Site (description):
Latitude:.
Site or Map Number:
Date:
State:
Longitude:
Time:
Weather in past 24 hours:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
Weather now:
Q Storm (heavy rain)
P Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
-------
MACROINVERTEBRATES AND HABITAT I 57
Sketch of site
On your sketch, note features that affect stream habitat, such as: riffles, runs, pools, ditches, wetlands, dams, riprap,
outfalls, tributaries, landscape features, logging paths, vegetation, and roads.
-------
MACROINVERTEBRATES AND HABITAT
CO
.0
•2
0
m
CD
TO
CO
Q.
o
in
0)
O)
CO
a.
o
in
CO
D>
CO
Q.
o
in
a
O)
CO
a.
5
CD
D)
CO
§.
11
o
c *-M r\s n n n t~
^
i n n n n
fi\ -f-' t ,.. *™ CO c ~P ^ ^ ' — ' ' — ' ' — * t~~f *""* ' — ' ' — ' l~~'
"o CM oa
ftj A A
c5 o a
T5
S
.213 £
"S. to SC
a
•C £ CC
o ~
1 1 11
1 *«•&*
S IS Is
"2 o o
li £ f
. Q. D.
•g -s -8
S
O T- (Or
.«
g E v E v
s 1° 1°
£ a a
33 S S
f
s
5 e
o>
•
•
M
i
a!
TI
v
V
ft.
-i
I
CO
^r
II
GL
to
•^
i^? *^
In-Stream Characterls
Check which stream habitats are presen
"t/T
c
=>
CO
CD
en
£
CO
CO
(You can check more than 1 habitat)
Q Pool(s) Q Riffle(s) Q R
Nature of particles in the stream bottom
.!£
dth of stream channe
'i
&
I
Q.
O
W
O
CD
0
CO
CD CD ^
Q measured Q i
"S
>£
a
a
o
CD
.&
"o
a>
to
g>
CO
T—
T-
cT^*
im (100 yds.), pickth
pe of the stream ban
Vertical/undercut
Steeply sloping (> 30
2 •
s ^
w o E
a 2 as
= £ E <5 a
raw S
|S i5
a
Jl S
CM
£
a .°>a
rr
— . w
0^ C
e o
Gradual/no slope (< o
f icial bank modificati
Bank 0-25% covered
•c
CO
1^
o tt
o •£ « a
s
$
g
_ o
a
aa
l_L. ^
S ^ *
•o "S
- -
S ;;a:r-aaaaaaaa
"c
N i o
j- J<: CD
•^ "^ JJ 0) w ii
a a '5 |g g co s
S Q. CO ^ ^
«•§•§ ^o™ HCQI--
o S S" ¥ w *
CD C C
C" ^- *~
•?? irt m
*. z z
CO
£
Boulders/rocks
Gravel/sand
Bare soil
Pavement, structu
2 g "is cMaaaoaaaa
w o co ^
O
a a a « a a
CO
£
CO
••^
o
^^>
a. CD 3 j
O>'5> £?
E E o
i* CO 3
0 ID **
_i tn -2-
CO
a a a a a
a a a
en
!>
£
s?
O^1 "U
LO Q)
T3 fl)
5
CD
i
CO
2
0)
C
H
CD
0.
a
Boulders (over 1 0" in diam.) Q
Bedrock (solid) Q
Pick the category that best describes the
which gravel, cobbles, and boulders on 1
bottom are embedded (sunk) in silt, sane
Q Somewhat/not embedded (0-25%) Q f
Q Halfway embedded (50%) Q (
Presence of logs or large woody debris i
Q None Q Occasional
"i"
CD
81
CL
"5
•••
&
CO
'1
0-
a
Presence of naturally-occurring organic
(i.e., leaves and twigs, etc.) in stream:
Q None Q Occasional
O>
i
c?
0 C
in ir
CD a
D) C
CO CC
Q. a.
j=
ill II
Water appearance:
0
a
O O Z O
a a a a
Q Clear Q Light brown
Q Milky Q Dark brown
Q Foamy Q Oily sheen
Q Turbid
Water odor:
Q Sewage Q Fishy
Q Chlorine Q Rotten eggs
Water temperature:
n
LL
o
V,
0
o
-------
MACROINVERTEBRATES AND HABITAT
I
.8
§•
S
o
8
= CM
(0 s
a*
a
CD
CO
OS
g 2
'a
,-„
-D
cc eo
o
CCCL
Z)
o
§
l
l
ooc
. E — ,
1 S'.i'i
tllf
-S > CD §
SSloSoScBg
t
CD
o
Left
2
Q Q
Q Q
2
H_,
aa
aa
aa
oo
o
o
« 'w
.
cow
a |
CM a1 'a a a CMOO. waaon, o
8|
fi^
Si S
""S a
o CA
a> > ~' a.
il s I?
a S O ° r
= s
I?1
5tf
11
T-oaoo T-
1
I
S
35
1 a S*
3 c -e
a £
O)
c
£
o
51-
55
J- -e .
o
ca
2
^ to
. CO
8)8
.
s | t » «
f*
.
^ S 1 t £• 8
s -s g .1 ® •§
T-aaoa ^DOt-aaooa
-------
MACROINVERTEBRATES AND HABITAT
s
1
EC-
CO
J£
u
_<0
I- *>
is co
C CO
l!
CO
"•g
CO
co
CD
CO
CO
ec
81
II
Is
2 0.
•§*
I!
Is"
" o
as q =
S o ;
I
•5-g
2 o
18 -
5 :£
co "g
•a co
CD L-
o
o
E o
•S 2
CO CO
T3
3
XI
CO
co
CD
a
CD
25 S s' g i
u. J= a. I
.5 c
•8 2
§ 8
i
« cc
a a
£
CD
.E
o
CO
J£
^
a.
.ti
CO
CD
D
CM
CD
CO
£
o.
S
2
^
S o
| z
I °
u
m
CD
CM
a 5-
I
a>
.Q
*w
o
y
<0
•a
e.
-
g
o
8
a
ic
Q. ?
CO
CM
CO 5 CO
- "8 ?,
CD S?
CO £
£ O
tl
ap
w O
CO
•i "p ••§
§• .1 §
• CO !?:
I S o>
S 8 £
CD
.n
_c
I |
1 ig
s °!
•^ o 3
I w g, -CO
& .SPTH ?
a:
a
g,oa
«
CD
F
Pools
'
i
Q.
0
ana
-o I
CD c*
JZ C
O <0
CO 0)
o a
g. a a a
<
05
CO
£
t5
fl> w ro
^ "o JT
^ o *-
c S5 5
8 |1
» •§ §
CO CO o
•fe CD C
CM
g»
ws
CO
XCD
=
1 S
£
T _
D)CO
f
W
'c
O
s> aa
_
CO
c
.2
w
§
0
a
•o
CD O-
s: CD
II
CD b
•o 3
0
a
-------
MACROINVERTEBRATES AND HABITAT
4.2
Streamside Biosurvey
The Streamside Biosurvey is based on
the simple macroinvertebrate sampling
approach developed and used by the Ohio
Department of Natural Resources and the
Izaak Walton League of America's Save
Our Streams program and adapted by many
volunteer monitoring programs throughout
the United States.
This assessment approach has two basic
components. The first is a biosurvey of
aquatic organisms that involves collecting
and identifying macroinvertebrates in the
field and calculating an index of stream
quality. The second is the habitat character-
ization method known as the Streamside
Biosurvey Habitat Walk.
Two methods of macroinvertebrate
sampling are detailed in this section—one
for rocky-bottom streams (using a kick net)
and one for muddy-bottom streams (using a
dip net). Figure 4.7 illustrates and describes
the nets used for these assessments. Both of
these aquatic organism collection proce-
dures have been widely tested and used
successfully by many groups. You should
consult with a local aquatic scientist to
determine which method is appropriate for
streams in your area.
Like the Stream Habitat Walk de-
scribed in Section 4.1, the Streamside
Biosurvey is useful as a screening tool to
identify water quality problems and as an
educational tool to teach volunteers about
pollution and stream ecology. But instead
of randomly picking up rocks or sticks and
brushing off macroinvertebrates for simple
observation purposes, Streamside Bio-
survey volunteers are trained to use special
nets and standardized sampling protocols to
collect organisms from a measured area of
stream habitat. Volunteers identify col-
lected organisms, usually to the order level,
and sort them into taxonomic groups based
Note
The Streamside Biosurvey is based on protocols developed
and widely used by programs such as the Ohio Department of
Natural Resources, the Izaak Walton League of America, and
others. This manual recommends some modifications to their
established protocols. These include:
• A finer mesh size for the kick and dip nets used to sample
for macroinvertebrates
• In rocky-bottom streams, compositing three samples into
one before identifying macroinvertebrates rather than
identifying macroinvertebrates in three separate samples
and choosing the best result. Compositing generally
provides a more representative sample of the macroinverte-
brate community than a discrete sample taken from one part
of the riffle. Riffle areas have what is known as a patchy
distribution of organisms, meaning that different types of
organisms are naturally found in different parts of the riffle.
In order to more accurately assess the macroinvertebrate
community in a rocky-bottom stream site, it is important to
take a representative sample that includes organisms found
in different microhabitats—such as in different parts of the
riffle or in riffles of various if lows and depths.
• A new method for calculating the stream quality rating. This
modification incorporates a weighting factor to take into
account the abundance of organisms in each pollution
tolerance category (pollution-sensitive, somewhat tolerant,
and tolerant).
• In muddy-bottom streams, varying how much each habitat
type is sampled depending on its abundance at the sam-
pling site.
on their ability to tolerate pollution. Using
this information, volunteers can> then
calculate a simple stream quality rating of
good, fair, or poor.
Because the Streamside Biosurvey
involves a standardized sampling protocol,
a basic level of training, professional
assistance, and a simple stream rating
based on macroinvertebrate diversity and
abundance, this approach is more effective
than the Stream Habitat Walk in character-
izing stream health and determining
general water quality trends over several
years. However, this method is not gener-
ally suited to determining subtle pollution
impacts due, in part, to its uncomplicated
level of macroinvertebrate identification
-------
MACROINVERTEBRATES AND HABITAT
and analysis. This, of course, is also one of
the Streamside Biosurvey's greatest
strengths, since volunteers can be easily
trained in its methods.
Key features of the Streamside
Biosurvey are as follows:
• It includes the Streamside Biosurvey
Habitat Walk as its physical habitat
characterization and visual biological
characterization components. This
protocol is a somewhat more detailed
version of the Stream Habitat Walk
described in Section 4.1.
• It centers around a macroinvertebrate
survey in which organisms are
collected according to specific
protocols, identified in the field
(generally to taxonomic order), and
are then released back into the
stream.
• For the identification process,
volunteers group macroinvertebrates
into three categories based on their
pollution tolerance or sensitivity.
Volunteers then calculate a water
quality index by counting the speci-
mens in each sensitivity category and
determining whether they are rare,
common, or dominant; multiplying
the number of taxa in each category
by a weighting factor; adding all the
scores; and comparing results to a
water quality rating scale that has
been determined by a locally knowl-
edgeable biologist/ecologist.
• The Streamside Biosurvey requires
some equipment and training.
Training can be conducted at the
stream site, although some advance
preparation is required. For example,
a biologist with regional experience
should assist in developing the
macroinvertebrate key and the
tolerance category groupings on the
field data sheets. A reference collec-
tion is recommended to help
volunteers identify macroinverte-
brates.
Step 1—Prepare for the
Streamside Biosurvey field work
Much of the preparation work for this
approach is similar to that of the Stream
Habitat Walk (section 4.1). Refer back to
that section for relevant information on the
following tasks:
• Scheduling the biosurvey
• Obtaining a USGS topographical
map
• Selecting and marking monitoring
locations
• Becoming familiar with safety
procedures
TASK 1 I Gather tools and equipment
for the Streamside Biosurvey
In addition to the basic equipment
listed in Section 2.4, you should collect the
following equipment needed for the macro-
invertebrate collection of the Streamside
Biosurvey:
• Vial with tight cap filled about one-
half full with 70 percent ethyl
alcohol
• Buckets (2)
• Hand lens, magnifying glass, or field
microscope
• Tweezers, eyedropper, or spoon
• Plastic bag
• Large, shallow, white pans, such as
dishpans (2)
• Spray water bottle
• Plastic ice cube tray
• Taxonomic key to aquatic organisms
• Calculator
• For rocky-bottom streams—Kick
net, a fine mesh (500 urn) nylon net
approximately 3x3 feet with a 3-foot
long supporting pole on each side is
recommended—Fig.4.7).
-------
MACROINVERTEBRATES AND HABITAT
Nets recommended in this manual
Kick net
For rocky-bottom stream sampling, a kick net
of 590 urn (a #30 mesh size) or 500 urn (#35
mesh size) is recommended. (Mesh size is
usually measured in microns, urn. The higher
the number, the coarser the mesh.)
D-framenet
For muddy-bottom stream sampling, a long-
handled D-frame or dip net is recommended
for reaching into vegetation that grows along
stream banks or is attached to the stream
bottom, and for sweeping up macroinverte-
brates dislodged from woody debris. D-frame
nets also come in different mesh sizes.
This manual recommends that volunteer programs purchase their macroinvertebrate
sampling nets from scientific supply houses to ensure a standard degree of net quality and known
mesh size. Some supply houses might sell the components of the net separately. Volunteer
programs then buy the net material commercially, supply their own handles, and build the nets
using volunteer labor.
Many programs use coarser mesh than is recommended in this manual. Coarser mesh is
generally less expensive. However, smaller organisms can be lost through the mesh during
sampling. If you are in doubt as to what mesh size to use, consult your technical advisor. If
possible—and especially if you want your volunteer data to be used by state and local water
managers—it is best to use nets of the same type and size as those which water quality profes-
sionals use in your state.
Other types of commonly used nets
Metal frame net
Used by the River Watch Network for
sampling both rocky-bottom and muddy-
bottom streams.
Surber sampler
Used by professional monitoring programs,
this sampler delineates an exact stream
bottom area to be disturbed.
Figure 4.7
Examples of
macroinverte-
brate sampling
nets
Nets used by
professionals
and volunteers
vary in overall
size, design,
and mesh size.
-------
MACROINVERTEBRATES AND HABITAT
• For muddy-bottom streams—D-
frame net (a dip net with a frame 12
inches wide with a fine nylon mesh,
usually about 500 [im, attached to
the frame).
Step 2—Collect and Sort
Macroinvertebrates
The method you use to collect macroin-
vertebrates using this approach depends on
the type of stream you are sampling.
Rocky-bottom streams are defined as those
with bottoms made up of gravel, cobbles,
and boulders in any combination and
usually have definite riffle areas. Riffle
areas are fairly well oxygenated and,
therefore, are prime habitats for benthic
macroinvertebrates. In these streams, use
the rocky-bottom sampling method.
Muddy-bottom streams have muddy,
silty, or sandy bottoms and lack riffles.
Generally, these are slow moving, low-
gradient streams (i.e., streams that flow
along relatively flat terrain). In such
streams, macroinvertebrates generally
attach themselves to overhanging plants,
roots, logs, submerged vegetation, and
stream substrate where organic particles are
trapped. In these streams, use the muddy-
bottom sampling method.
Both methods are detailed below.
Regardless of which collection method is
used, the process for counting, identifying,
and analyzing the macroinvertebrate sample
for the Streamside Biosurvey is the same.
Rocky-Bottom Sampling Method
Use the following method of macroin-
vertebrate sampling in streams that have
riffles and gravel/cobble substrates. You
will collect three samples at each site and
composite (combine) them to obtain one
large total sample.
TASK1
Identify the sampling
location
You should have already located your
site on a map along with its latitude and
longitude (see Task 3, page 45).
1. You are going to sample in three
different spots within a 100-yard
stream reach. These spots may be
three separate riffles; one large riffle
with different current velocities; or,
if no riffles are present, three run
areas with gravel or cobble substrate.
Combinations are also possible (if,
for example, your site has only one
small riffle and several run areas).
Mark off your 100-yard stream
reach. If possible, it should begin at
least 50 yards upstream of any
human-made modification of the
channel, such as a bridge, dam, or
pipeline crossing, Avoid walking in
the stream, since this might dislodge
macroinvertebrates and alter your
sampling results.
2. Sketch the 100-yard sampling area.
Indicate the location of your three
sampling spots on the sketch. Mark
the most downstream site as Site 1,
the middle site as Site 2, and the
upstream site as Site 3. (See Fig.
4.8.)
TASK 2 | Get into place
1. Always approach your sampling
locations from the downstream end
and sample the site farthest down-
stream first (Site 1) (see Fig. 4.9,
Panel #1). This minimizes the
possibility of biasing your second
and third collections with dislodged
sediment or macroinvertebrates.
Always use a clean kick net,
relatively free of mud and debris
from previous uses. Fill a bucket
about one third full with stream
water and fill your spray bottle.
-------
MACROINVERTEBRATES AND HABITAT I 65
2. Select a 3-foot by 3-foot riffle area
for sampling at Site 1. One member
of the team, the net holder, should
position the net at the downstream
end of this sampling area. Hold the
net handles at a 45 degree angle to
the water's surface (see Fig. 4.9,
Panel #2). Be sure that the bottom of
the net fits tightly against the stream-
1 bed so no macroinvertebrates escape
under the net. You may use rocks
from the sampling area to anchor the
net against the stream bottom. Don't
allow any water to flow over the net.
TASK 3
1
Dislodge the macroinverte-
brates
Pick up any large rocks in the 3-foot
by 3-foot sampling area and rub them
thoroughly over the partially-filled
bucket so that any macroinverte-
brates clinging to the rocks will be
dislodged into the bucket (see Fig.
4.9, Panel #3). Then place each
cleaned rock outside of the sampling
area. After sampling is completed,
rocks can be returned to the stretch
of stream they came from.
2. The member of the team designated
as the "kicker" should thoroughly stir
up the sampling area with their feet,
starting at the upstream edge of the
3-foot by 3-foot sampling area and
working downstream, moving toward
the net. All dislodged organisms will
be carried by the stream flow into the
net (see Fig. 4.9, Panel #4). Be sure
to disturb the first few inches of
stream sediment to dislodge burrow-
ing organisms. As a guide, disturb
the sampling area for about 3 min-
utes, or until the area is thoroughly
worked over.
3. Any large rocks used to anchor the
net should be thoroughly rubbed into
the bucket as above.
Sampling sites
TASK 4
Remove the net
1
Next, remove the net without
allowing any of the organisms it
contains to wash away. While the net
holder grabs the top of the net
handles, the kicker grabs the bottom
of the net handles and the net's
bottom edge. Remove the net from
the stream with a forward scooping
motion (see Fig. 4.9, Panel #5).
2. Roll the kick net into a cylinder
shape and place it vertically in the
partially filled bucket. Pour or spray
water down the net to flush its
contents into the bucket (see Fig.
4.9, Panel #6). If necessary, pick
debris and organisms from the net by
hand. Release back into the stream
any fish, amphibians, or reptiles
caught in the net.
TASK 5 \ Collect the second and third
^^^™1™^ samples
Once you have removed all the organ-
isms from the net repeat these tasks at Sites
2 and 3. Put the samples from all three
sites into the same bucket. Combining the
debris and organisms from all three sites
into the same bucket is called compositing.
Figure 4.8
••^•••^•i
Location of
sample sites in
a rocky-bottom
stream with
riffles
Within a 100
yard reach
volunteers begin
their sampling at
the most
downstream site
and then work
their way
upstream.
-------
Figure 4.9
•^••••••i
Procedures for
collecting a
macrofnverte-
brate sample in
a rocky-bottom
stream
Volunteers must
follow set
protocol to
collect an
unbiased
sample.
1. Approach the sample site from the
downstream end.
2. Position the net at a 45° angle with
the bottom tight against the sub-
strate.
3. Dislodge macroinvertebrates by
rubbing rocks thoroughly.
4. Disturb the substrate thoroughly
with your feet.
5. Remove the net with a forward
scooping motion.
6. Flush out the net with clean
stream water.
-------
MACROINVERTEBRATES AND HABITAT
Hint: If your bucket is nearly full of water
after you have washed the net clean, let the
debris and organisms settle to the bottom
of the bucket. Then cup the net over the
bucket and pour the water through the net
into a second bucket. Inspect the water in
the second bucket to be sure no organisms
came through.
TASK 6
Sort macroin vertebrates
Pour the contents of the bucket into a
large, shallow, white pan. Add some stream
water to the pan, and fill the ice cube tray
with stream water. Using tweezers, eye
dropper, or spoon, pick through the leaf
litter and organic material looking for
anything that swims, crawls, or seems to be
hiding in a shell, like a snail. Look care-
fully; many of these creatures are quite
small and fast-swimming. Sort similar
organisms into the ice cube tray.
Note: Instructions for counting, identifying,
and analyzing the macroinvertebrate
sample follow the muddy-bottom sampling
method. (See page 70, Step 3)
Muddy-Bottom Sampling Method
In muddy-bottom streams, as in rocky-
bottom streams, the goal is to sample the
most productive habitats available and look
for the widest variety of organisms. The
most productive habitats are the ones that
harbor a diverse population of pollution
sensitive-macroinvertebrates. Volunteers
should sample by using a D-frame net to
jab at the habitat and scoop up the organ-
isms that are dislodged. The objective is to
collect a combined sample from 20 jabs
taken from a variety of habitats.
Picking Bugs
Some monitoring programs find it easier to collect organisms
from the net by hand-picking them rather than washing the sample
into a pan and then trying to pick through the floating debris. The
advantage to placing the organisms in a pan is that they are more
likely to survive while in the pan and their characteristic move-
ments will help in organism identification.
If you prefer to pick bugs directly off the net, a white back-
ground, such as a white plastic trash bag under the net, will help
you see the bugs more clearly, lln addition, periodically wetting the
net with a water bottle will help keep the bugs alive and moving.
Identification can be made easier if you sort the organisms into
groups based on physical similarities and place them together in
sections of an ice cube fray as you pick them from the pan or net.
TASK 1 | Determine which habitats are
™^"™""""""^ present
Muddy-bottom streams usually have
four habitats (Fig. 4.10). It is generally best
to concentrate sampling efforts on the most
productive habitat available, yet to sample
other principal habitats if they are present.
This ensures that you will secure as wide a
variety of organisms as possible. Not all
habitats are present in all streams or present
in significant amounts. If your sampling
areas have not been preselected, try to
determine which of the following habitats
are present. (Avoid standing in the stream
while making your habitat determinations.)
• Vegetated bank margins. This
habitat consists of overhanging bank
vegetation and submerged root mats
attached to banks. The bank margins
may also contain submerged,
decomposing leaf packs trapped in
root wads or lining the streambanks.
This is generally a highly productive
habitat in a muddy-bottom stream,
and it is often the most abundant
type of habitat.
• Snags and logs. This habitat consists
of submerged wood, primarily dead
trees, logs, branches, roots, cypress
knees and leaf packs lodged between
rocks or logs. This is also a very
productive muddy-bottom stream
habitat.
-------
I Aquatic vegetation beds and decay-
ing organic matter. This habitat
consists of beds of submerged, green/
leafy plants that are attached to the
stream bottom. This habitat can be as
productive as vegetated bank mar-
gins, and snags and logs.
Silt/sand/gravel substrate. This
habitat includes sandy, silty, or
muddy stream bottoms; rocks along
the stream bottom; and/or wetted
gravel bars. This habitat may also
contains algae-covered rocks (some-
times called Aufwuchs). This is the
least productive of the four muddy-
bottom stream habitats, and it is
always present in one form or
another (e.g., silt, sand, mud, or
gravel might predominate).
TASK 2
Determine how many times
^~~~^^~1 to jab in each habitat type
Your goal is to jab a total of 20 times.
The D-frame net is 1 foot wide, and a jab
should be approximately 1 foot in length.
Thus, 20 jabs equals 20 square feet of
combined habitat.
Figure 4.10
Four habitats
found in
muddy-bottom
streams
Volunteers will
likely find the
most macroin-
vertebrates in
vegetated
habitats and
snags and logs.
• If all four habitats are present in
plentiful amounts, jab the vegetated
banks 10 times and divide the
remaining 10 jabs among the remain-
ing 3 habitats.
• If three habitats are present in
plentiful amounts and one is absent,
jab the silt/sand/gravel substrate—
the least productive habitat—5 times
and divide the remaining 15 jabs
among the other two more produc-
tive habitats.
• If only two habitats are present in
plentiful amounts, the silt/sand/
gravel substrate will most likely be
one of those habitats. Jab the silt/
sand/gravel substrate 5 times and the
more productive habitat 15 times.
• If some habitats are plentiful and
others are sparse, sample the sparse
habitats to the extent possible, even
if you can take only one or two jabs.
Take the remaining jabs from the
plentiful habitat(s). This rule also
applies if you cannot reach a habitat
because of unsafe stream conditions.
Jab a total of 20 times.
Because you might need to make an
educated guess to decide how many jabs to
take in each habitat type, it is critical that
you note, on the field data sheet, how many
jabs you took in each habitat. This informa-
tion can be used to help characterize your
findings.
TASK 3 I Get into place
Outside and downstream of your first
sampling location (1st habitat), rinse the dip
net and check to make sure it does not
contain any macroinvertebrates or debris
from the last time it was used. Fill a bucket
approximately one-third full with clean
stream water. Also, fill the spray bottle with
clean stream water. This bottle will be used
to wash down the net between jabs and
after sampling is completed.
-------
MACROINVERTEBRATES AND HABITAT
This method of sampling requires only
one person to disturb the stream habitats.
While one person is sampling, a second
person should stand outside the sampling
area, holding the bucket and spray bottle.
After every few jabs, the sampler should
hand the net to the second person, who then
can rinse the contents of the net into the
bucket.
TASK 4
Dislodge the macroinverte-
brates
Approach the first sample site from
downstream, and sample as you walk
upstream. Here is how to sample in the four
habitat types:
• Sample vegetated bank margins by
jabbing vigorously, with an upward
motion, brushing the net against
vegetation and roots along the bank.
The entire jab motion should occur
underwater.
• To sample snags and logs, hold the
net with one hand under the section
of submerged wood you are sam-
pling: With the other hand (which
should be gloved), rub about 1
square foot of area on the snag or
log. Scoop organisms, bark, twigs, or
other organic matter you dislodge
into your net. Each combination of
log rubbing and net scooping is one
jab (Fig. 4.11).
• To sample aquatic vegetation beds,
jab vigorously, with an upward
motion, against or through the plant
bed. The entire jab motion should
occur underwater.
• To sample a silt/sand/gravel sub-
strate, place the net with one edge
against the stream bottom and push
it forward about a foot (in an up-
stream direction) to dislodge the first
few inches of silt, sand, gravel, or
rocks. To avoid gathering a netful of
mud, periodically sweep the mesh
bottom of the net back and forth in
the water, making sure that water
does riot run over the top of the net.
This will allow fine silt; to rinse out
of the net.
When you have completed all 20 jabs,
rinse the net thoroughly into the bucket. If
necessary,-pick any clinging organisms
from the net by hand and put them in the
bucket.
Figure 4.11, ,
Collecting a
sample from a
log
Volunteer rubs
the log with one
hand and
catches dis-
lodged organ-
isms and other
material in the
net.
-------
MACROINVERTEBRATES AND HABITAT
TASKS
Sort the macroinvertebrates
Pour the contents of the bucket (water,
organisms, and organic material) into a
large, shallow, white pan and fill the ice
cube tray with clean stream water. Using
tweezers, eye dropper, or spoon, pick
through the leaf litter and organic material
looking for anything that swims, crawls, or
seems to be hiding in a shell (like a snail).
Look carefully; many of these creatures are
quite small and fast-swimming. Sort similar
organisms into the plastic ice cube tray.
Step 3—Identify Macroinverte-
brates and Calculate Stream
Rating
The following methods are used for
both the rocky- and muddy-bottom assess-
ments.
Task 1 I Identify Macroinvertebrates
1. Identify the collected macroinverte-
brates. Using the hand lens or
magnifying glass and the aquatic
organism identification key, carefully
observe the collected macroinverte-
brates. Refine your initial sort so that
like individuals are placed in the
same section(s) of the ice cube tray.
If you cannot identify an organism,
place one or two specimens in the
alcohol-filled vial and forward it to
your program coordinator for identi-
fication.
2. On your field data sheet, note the
number of individuals of each type of
organism you have identified (Sec-
tion 3 of the field data sheet—See
Fig. 4.12.).
Note: When you feel that you have
identified all the organisms to the best of
your ability, return the macroinvertebrates
to the stream.
3. Assign one of the following abun-
dance codes to each type of
organism. Record the code next to
the actual count on the field data
sheet.
R (rare)
C (common)
D (dominant)
if 1-9 organisms are
found in the sample
if 10-99 organisms are
found in the sample
if 100 or more organ-
isms are found in the
sample
Your field data sheet should be orga-
nized to help you sort macroinvertebrates
into three groups based on their ability to
tolerate pollution. A local authority (such
as a state biologist or entomologist)
should determine which organisms
belong in each pollution tolerance cat-
egory for your region.
Generally, the three tolerance groups
are as follows:
• Group I (sensitive organisms)
includes pollution- sensitive
organisms such as mayflies,
stoneflies, and non net-spinning
caddisflies, which are typically
found in good-quality water.
• Group II (somewhat sensitive
organisms) includes somewhat
pollution-tolerant organisms such
as net-spinning caddisflies,
crayfish, sowbugs, and clams,
found in fair-quality water.
• Group III (tolerant organisms)
includes pollution-tolerant
organisms such as worms,
leeches, and midges, found in
poor-quality water.
TASK 2 j Calculate the stream quality
""""^^"'""^ rating
The stream water quality rating takes
into account the pollution sensitivity of the
organisms and their relative abundance.
This is accomplished through use of a
weighting system.
-------
MACROINVERTEBRATES AND HABITAT I 71
MACROINVERTEBRATE COUNT
i
Identify the macroinvertebrates in your sample and assign them letter codes based
on their abundance: R (rare) = 1-9 organisms; C (common) = 10-99 organisms; and
D (dominant) = 100 plus organisms. ;
Group I
Sensitive
Group II Group III
Somewhat-Sensitive Tolerant
C(50) Water penny larvae
R@) Hellgrammites
Mayfly nymphs
Gilled snails
Riffle beetle adult
C(25) stonefly nymphs
Non net-spinning
caddisfly larvae
. Beetle larvae
. Clams
. Crane fly larvae
. Crayfish
Damselfly nymphs
R(5) Aquatic worms
Blackfly larvae
Leeches
. Midge larvae
C(50) Snails
D(100) Scuds
£>(750)sowbugs
R(8) Fishfly larvae
Alderfly larvae
C(27) Net-spinning
caddisfly larvae
The weighting system acknowledges
the most desirable combinations of pollu-
tion sensitivity and abundance by assigning
these extra weights within a 5, 3, and 1
point scale. Pollution-sensitive organisms
receive a weighting factor based on a 5-
ppint scale. Somewhat sensitive organisms
are weighted on a 3-point scale, and
tolerant organisms are weighted on a 1-
point scale. As can be seen in Table 4.2, a
sample's ideal combination of organisms
would be "sensitive" and "somewhat
sensitive" organisms in common abundance
(10-99 organisms), and pollution "tolerant"
organisms in rare abundance (less than 10
organisms). This is because it is never ideal
for any given type of organism to dominate
a sample, and because it is best to have a
wide variety of organisms including a few
pollution-tolerant individuals.
1. Add the number of R's, C's and D's
in each of the 3 pollution tolerance
groupings. Then, for each grouping,
multiply the total number of R's, C's
and D's by the relevant weighting
factor. Table 4.3 illustrates sample
calculations for determining the
water quality rating for (hypotheti-
cal) Volunteer Creek.
Note: The tolerance category groupings
shown on the Biosurvey Data Sheet were
developed for streams in the mid-Atlantic
(Maryland, Virginia, West Virginia, District
of Columbia, Pennsylvania). These
groupings may not totally apply in other
regions of the United States. It Js impor-
tant that a local aquatic biologist take a
look at these categories and make any
changes necessary for your region.
In addition, depending on the level of
taxonomic training volunteers receive, you
might consider separating out some other
families of organisms. For instance, the
tolerance groupings given here separate
caddisflies into net-spinning and non net-
spinning families. Mayflies might also be
separated into different tolerance group-
ings. It is not recommended here, however,
because of the difficulty in distinguishing
mayfly families in the field without a
microscope.
Some volunteer programs, like the one
coordinated by the Audubon Naturalist
Society in Maryland, conduct intensive field
identification training workshops and teach
volunteers to distinguish several families in
the field. Creating more specific tolerance
groupings may be an option for your
program if you have the resources and
expertise to conduct more intensive
taxonomic field training.
Figure 4.12
Sample macro-
invertebrate
count for
(hypothetical)
Volunteer
Creek
-------
MACROINVERTEBRATES AND HABITAT
Table 4.2
Weighting
factors used in
calculating
stream water
quality ratings
Abundance Weighting Factor
Rare (R)
Common (C)
Dominant (D)
Group 1
Sensitive
5.0
5.6
5.3
Group II
Somewhat Sensitive
3.2
3.4
3.0
Group III
Tolerant
1.2
1.1
1.0
Table 4.3
Sample calcu-
lations of index
values for
Volunteer
Creek
Table 4.4
Tentative
rating scale for
streams in
Maryland
Group I
Sensitive
1 (No. of R's) x 5.0 = 5.0
2 (No. ofC's)x5.6=11.2
Index Value for Group I = 16.2
Group II
Somewhat Sensitive
3 (No. of R's) x 3.2 = 9.6
1 (No. of C's) x 3.4 = 3.4
2 (No. of D's) x 3.0 = 6.0
Index Value for Group II = 19.0
Group HI
Tolerant
1 (No. of R's) x 1.2 = 1.2
1 (No. of C's) x 1.1 =1.1
Index Value for Group III = 2.3
Score Rating
>40
20-40
<20
Good
Fair
Poor
2. To obtain a water quality rating for
the site, total the values for each
group and add them together. The
total score for the sample stream site
is: 16.2 (Group I) + 19.0 (Group II) +
2.3 (Group IH) = 37.5.
3. The final step is to compare the score
to water quality ratings (good to
poor) established by a trained
biologist familiar with local stream
fauna. Table 4.4 presents a tentative
rating scale for streams in Maryland.
Assuming Volunteer Creek is located
in Maryland, the stream would
receive a rating of "Fair."
Note: In addition to adjusting the rating
scale according to regional location, it
might also need to be adjusted for muddy-
bottom vs. rocky-bottom streams. An
experienced stream biologist can calculate
the best rating system for your area's
streams by examining data from several
streams.
In a healthy stream, the sensitive
(Group I) organisms will be well repre-
sented in a sample. It is important to
remember that macroinvertebrate popula-
tions can fluctuate seasonally and that these
natural fluctuations can affect your results.
Therefore, it is best to compare the results
by season from year to year. (Compare your
spring sampling results to each other, not to
fall results.)
-------
MACROINVERTEBRATES AND HABITAT I 73
Step 4—Conduct the Streamside
Biosurvey: Habitat Walk
You will conduct a habitat assessment
(which will include measuring general
characteristics and local land use) in a 100-
yard section of stream that includes the
riffles from which organisms were col-
lected.
TASK 1 [ Delineate the habitat assess-
ment boundaries
1. Begin by identifying the most
downstream riffle that was sampled
for macroinvertebrates. Using your
tape measure or twine, mark off a
100-yard section extending 25 yards
below the downstream riffle and
about 75 yards upstream.
2. Complete the identifying information
on your field data sheet for your
habitat assessment site. On your
stream sketch, be as detailed as
possible, and be sure to note which
riffles were sampled.
TASK 2
Complete the Physical
Characteristics, Local
Watershed Characteristics,
and Visual Biological Survey
sections of the field sheet
For safety reasons as well as to protect
the stream habitat, it is best to estimate
these characteristics rather than actually
wading into the stream to measure them.
In-stream Characteristics
1. Pools, riffles, and runs create a
mixture of flows and depths and
provide a variety of habitats to
support fish and invertebrate life.
Pools are deep with slow water.
Riffles are shallow with fast, turbu-
lent water running over rocks. Runs
are deep with fast water and little or
no turbulence.
2. Stream bottom (substrate) is the
material on the stream bottom.
Identify what substrate types are
present. Substrate types include:
• Silt/clay/mud—This substrate has
a sticky, cohesive feeling. The
particles are fine. The spaces
between the particles hold a lot of
water, making the sediments
behave like ooze.
• Sand (up to 0.1 inch)—A sandy
bottom is made up of tiny, gritty
particles of rock that are smaller
than gravel but coarser than silt
(gritty, up to pea size).
• Gravel (0.1-2 inches)—A gravel
bottom is made up of stones
ranging from tiny quarter-inch
pebbles to rocks of about 2 inches
(fine gravel - pea size to marble
size; coarse gravel - marble to
tennis ball size).
• Cobbles (2-10 inches)—Most
rocks on this type of stream
bottom are between 2 and 10
inches (between a tennis ball and
a basketball). [
Figure 4.13
Overview and
cross sections
* of a pool, riffle,
and run .
Varying flows
and depths
create a variety
of habitats for
macroinverte-
brates.
-------
MACROINVERTEBRATES AND HABITAT
• Boulders (greater than 10
inches)—Most of the rocks on the
bottom are greater than 10 inches
(between a basketball and a car in
size).
• Bedrock—is solid rock (or rocks
bigger than a car).
Estimate the percentage of substrate
types at your site.
3. Embeddedness is the extent to which
rocks (gravel, cobbles, and boulders)
are sunken into the silt, sand, or mud
of the stream bottom (Fig. 4.14).
Generally, the more rocks are
embedded, the less rock surface or
space between rocks is available as
habitat for aquatic macroinverte-
brates and for fish spawning.
Excessive silty runoff from erosion
can increase the embeddedness in a
stream. To estimate the
embeddedness, observe the amount
of silt or finer sediments overlying,
in between, and surrounding the
rocks.
4. Streambed stability can provide
additional clues to the amount of
siltation in a stream. When you walk
in the stream, note whether your feet
sink significantly into sand or mud.
5. Presence of logs or woody debris
(not twigs and leaves) in stream can
slow or divert water to provide
important fish habitat such as pools
and hiding places. Mark the box that
describes the general amount of
woody debris in the stream.
6. Naturally occurring organic material
in stream. This material includes
leaves and twigs. Mark the box that
describes the general amount of
organic matter in the stream.
7. Water appearance can be a physical
indicator of water pollution.
• Clear - colorless, transparent
• Milky - cloudy-white or grey, not
transparent; might be natural or
due to pollution
• Foamy - might be natural or due
to pollution, generally detergents
or nutrients (foam that is several
inches high and does not brush
apart easily is generally due to
some sort of pollution)
• Turbid - cloudy brown due to
suspended silt or organic material
• Dark brown - might indicate that
acids are being released into the
stream due to decaying plants
• Oily sheen - multicolored reflec-
tion might indicate oil floating in
the stream, although some sheens
are natural
• Orange - might indicate acid
drainage
• Green - might indicate excess
nutrients being released into the
stream
8. Water odor can be a physical indica-
tor of water pollution
• No smell or a natural odor
• Sewage - might indicate the
release of human waste material
• Chlorine - might indicate over-
chlorinated sewage treatment/
water treatment plant or swim-
ming pool discharges
• Fishy - might indicate the pres-
ence of excessive algal growth of
dead fish
• Rotten eggs - might indicate
sewage pollution (the presence of
methane from anaerobic condi-
tions)
9. Water temperature can be particu-
larly important for determining the
suitability of the stream as aquatic
habitat for some species of fish and
macroinvertebrates that have distinct
-------
MACROINVERTEBRATES AND HABITAT
temperature requirements. Tempera-
ture also has a direct effect on the
amount of dissolved oxygen avail-
able to the aquatic organisms.
Measure temperature by submerging
a thermometer for at least 2 minutes
in a typical stream run. Repeat once
and average the results.
Stream Bank and Channel
Characteristics
10. Depth of runs and pools should be
determined by estimating the vertical
distance from the surface to the
stream bottom at a representative
depth at each of the two habitats.
11. The width of the stream channel can
be determined by estimating the
width of the streambed that is
covered by water from bank to bank.
If it varies widely, estimate an
average width.
12. Stream velocity can have a direct
influence on the health, variety, and
abundance of aquatic communities.
If water flows too quickly, insects
might be unable to maintain their
hold on rocks and vegetation and be
washed downstream; if water flows
too slowly, it might provide insuffi-
cient aeration for species needing
high levels of dissolved oxygen.
Stream velocity can be affected by
dams, channelization, terrain, runoff,
and other factors. To measure stream
velocity, mark off a 20-foot section
of stream run and measure the time it
takes a stick, leaf, or other floating
biodegradable object to float the 20
feet. Repeat 5 times and pick the
average time. Divide the distance
(20 feet) by the average time (sec-
onds) to determine the velocity in
feet per second. (See Chapter 5,
Section 1 on flow for a more in-
depth discussion on using floats to
estimate velocity.)
13. The shape of the stream bank, the
extent of artificial modifications, and
the shape of the stream channel are
determined by standing at the
downstream end of the 25-yard
section and looking upstream.
(a) The shape of the stream bank (Fig.
4.15) may include.
• Vertical or undercut bank - a
bank that rises vertically or
overhangs the stream. This type
of bank generally provides good
cover for macroinvertebrates and
fish and is resistant to erosion.
However, if seriously undercut, it
might be vulnerable to collapse.
• Steeply sloping - a bank that
slopes at more than a 30 degree
angle. This type of bank is very
vulnerable to erosion.
Figure 4.14
A representa-
tion of a rocky-
bottom stream
becoming
embedded with
sand and silt
As silt settles on
the streambed,
spaces between
the rocks are
filled in and the
stream be-
comes more
embedded.
-------
MACROINVERTEBRATES AND HABITAT
Figure 4.15
Types of
streambank
shapes
Undercut banks
provide good
cover for fish
and macroinver-
tebrates.
*<*•'/: /i &&££=smm^'
• Gradual sloping - a bank that has
a slope of 30 degrees or less.
Although this type of stream bank
is highly resistant to erosion, it
does not provide much streamside
cover.
(b) Artificial bank modifications include
all structural changes to the stream
bank such as riprap (broken rock,
cobbles, or boulders placed on earth
surfaces such as the face of a dam or
the bank of a stream, for protection
against the action of the water) and
bulkheads. Determine the approxi-
mate percentage of each bank (both
the left and right) that is artificially
covered by the placement of rocks,
wood, or concrete.
(c) The shape of the stream channel can
be described as narrow (less than 6
feet wide from bank to bank), wide
(more than 6 feet from bank to bank),
shallow (less than 3 feet deep from
the stream substrate to the top of the
banks) or deep (more than 3 feet
from the stream substrate to the top
of the banks). Choose the category
that best describes the channel.
• Narrow, deep
• Narrow, shallow
• Wide, deep
• Wide, shallow
14. Streamside cover information helps
determine the quality and extent of
the stream's riparian zone. This
information is important at the
stream bank itself and for a distance
away from the stream bank. For
example, trees, bushes, and tall grass
can contribute shade and cover for
fish and wildlife and can provide the
stream with needed organic material
such as leaves and twigs. Lawns
indicate that the stream's riparian
zone has been altered, that pesticides
and grass clippings are a possible
problem, and that little habitat and
shading are available. Bare soil and
pavement might indicate problems
with erosion and runoff. Looking
upstream, provide an estimate of the
percentage of the stream bank (left
and right stream banks) covered by
the following:
• Trees
• Bushes, shrubs - conifers or
deciduous bushes less than 15 feet
high
• Tall grass, ferns, etc. - includes
tall natural grasses, ferns, vines,
and mosses
• Lawn - cultivated and maintained
short grass
• Boulders - rocks larger than 10
inches •
• Gravel/cobbles/sand - rocks
smaller than 10 inches; sand
-------
MACROINVERTEBRATES AND HABITAT
• Bare soil
• Pavement, structure - any man-
made structures or paved areas,
including paths, roads, bridges,
houses, etc.
15. Stream shading is a measurement of
the extent to which the stream itself
is overhung and shaded by the cover
identified in 14 above. This shade (or
overhead canopy) provides several
important functions in the stream
habitat. It cools the water; offers
habitat, protection, and refuge for
aquatic organisms; and provides a
direct source of beneficial organic
matter and insects to the stream.
Determine the extent that vegetation
shades the stream at the site.
16. General conditions of the stream
bank and stream channel, and other
conditions that might be affecting the
stream are determined by standing at
the downstream end of the 25-yard
site and looking upstream. Provide
observations for the right and left
banks of the stream.
(a) Stream bank conditions that might be
affecting the stream.
• Natural plant cover degraded—
note whether streamside
vegetation is trampled or missing
or has been replaced by landscap-
ing, cultivation, or pavement.
(These conditions could lead to
erosion.)
• Banks collapsed/eroded—note
whether banks or parts of banks
have been washed away or worn
down. (These conditions could
limit habitats in the area.)
• Garbage/junk adjacent to the
stream—note the presence of
litter, tires, appliances, car bodies,
shopping carts, and garbage
dumps.
• Foam or sheen on bank—note
whether there is foam or an oily
sheen on the stream bank. Sheen
may indicate an oil spill or leak,
and foam may indicate the
presence of detergent.
(b) Stream channel conditions that
might be affecting the stream.
• Mud/silt/sand on bottom/entering
stream—can interfere with the
ability of fish to sight potential
prey. It can clog fish gills and
smother fish eggs in spawning
areas in the stream bottom. It can
be an indication of poor construc-
tion practices, urban area runoff,
silviculture (forestry-related
activities), or agriculture in the
watershed. It can also be a
normal condition, especially in a
slow-moving, muddy-bottom
stream.
• Garbage or junk in stream—note
the presence of litter, tires,
appliances, car bodies, shopping
carts, and garbage.
(c) Other general conditions that might
be affecting the stream.
• Yard waste (e.g., grass clip-
pings)—ris there evidence that
grass clippings, cut branches, and
other types of yard waste have
been dumped into the stream?
• Livestock in or with unrestricted
access to stream—are livestock
present, or is there an! obvious
path that livestock use to get to
the water from adjacent fields? Is
there streamside degradation
caused by livestock?
• Actively discharging pipes—are
there pipes with visible openings
discharging fluids or water into
the stream? Note such pipes even
though you may not be able to
tell where they come from or
what they are discharging.
-------
MACROINVERTEBRATES AND HABITAT
• Other pipes—are there pipes near
or entering the stream? Note such
pipes even if you cannot find an
opening or see matter being
discharged.
• Ditches—are there ditches,
draining the surrounding land and
leading into the stream?
Local watershed characteristics
17. Adjacent land uses can potentially
have a great impact on the quality
and state of the stream and riparian
areas. Determine the land uses, based
on your own judgment of the activi-
ties in the watershed surrounding
your site within a quarter of a mile.
Enter a "1" if a land use is present
and a "2" if it is clearly having a
negative impact on the stream.
Visual biological survey
18. Are fish present in the stream? Fish
can indicate that the stream is of
sufficient quality for other organ-
isms.
19. Barriers to the movement offish in
the stream are obstructions that
would keep fish from moving freely
upstream or downstream.
20. Aquatic plants provide food and
cover for aquatic organisms. Plants
also might provide very general
indications of stream quality. For
example, streams that are overgrown
with plants could be over enriched by
nutrients. Streams devoid of plants
could be affected by extreme acidity
or toxic pollutants. Aquatic plants
may also be an indicator of stream
velocity because plants cannot take
root in fast-flowing streams.
21. Algae are simple plants that do not
grow true roots, stems, or leaves and
that mainly live in water, providing
food for the food chain. Algae may
grow on rocks, twigs, or other
submerged materials, or float on the
surface of the water. It naturally
occurs in green and brown colors.
Excessive algal growth may indicate
excessive nutrients (organic matter or
a pollutant such as fertilizer) in the
stream.
Step 4—Complete all the field
data sheets
After you have completed macroin-
vertebrate sampling, analysis of findings,
and the habitat characterization, make sure
you have completed the field data sheet to
the extent possible and that the recorded
data are legible. If you are not able to
determine how to answer a question on the
field data sheet, just leave the space blank.
Return all completed forms to your pro-
gram coordinator.
-------
MACROINVERTEBRATES AND HABITAT
STREAMSIDE BIOSURVEY: MACROINVERTEBRATES
Stream Name:
County:
Investigators:
Site (description):
Latitude:
State:
Longitude:
Site or Map Number:
Date:
Time:
Weather in past 24 hours:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
Weather now:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
-------
OJ
o s
o —
to D.
11
0
"*-
o
.«•
Q
II
o> S> =
£ 5 0.
*- CO 3
° £ 2
E So
§ -a t.
co £0
.2 t3
8
CM
CO
X
33"
DC
Tf
co'
x
b
q
co
X
33"
Q
'-a
x
v
SJft
•"(03
•5 > s
ill *
-^
o£
£ CO
&I
2 D-
T3 ®
T3 CO
CO JB
S3
CO jfi
•SI'S
s
CO
3
D-
CD
CD
T3
= tr r:
££ »
Group
#ofC's)x5.B
CO
in
X
33"
Q
•fc
3fc
• s-
£ 5 Q.
•-(03
•R > O
° M *-
|i2
co££
a.
^1
co ra
-1
i_ CO
CD (D
log
||§
3 g .•&
•i *!
o-S §•
g .s^
.2JS
-j
cr
ffl
I
D)
C
he
£ a5
§|
CO t
•= CO
£ CD
2? to
§.=
If
"T O
i CVJ
O V
CM
° .ir O
<
•^1
!-S
anc
and
CO
0
.Q
CD
I
'5
=
I
O
•iii
Q CO i~
CO CO ii
E J=> J!
5l§
^ ^ E
^•8 E
1 S 8
CD J3 —
2 ^> o
>
*-•
'1
d)
2
(0
If
o E
t- o
O (0
Group I
Sensitive
S
I
as
f
1
0
CD
CQ
CO
CD
^^ WJ IU
->• CD —
CD
-S5
s 5 ii
0)
i
CO
jo 2 co
O O O
1
ca
CO
a.
>,
c
co
D>
3
CD
CO
>,
q=
8 -S
to o o .2
Q CO CO U.
(D
i
_co
_>,
CD
73
<
J
,S«
i^
•Q.CO
"S
ST3
co
ZO
11
i I
a? £
2
CO
t s
I g
>,
CO
CD
0)
-0
J2 l?a>
a. 'E S?
1 tl
= .
2 -2> 5, 0 ^
« "33 to s •£
5 X 2 O DC
CO
§1
Z 0
-------
MACROINVERTEBRATES AND HABITAT
STREAMSIDE BIOSURVEY: HABITAT WALK
Stream Name:
County:
Investigators:
Site (description):
Latitude:.
State:
Longitude:
Site or Map Number:
Date:
Time:
Weather in past 24 hours:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
Weather now:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
-------
Sketch of site
On your sketch, note features that affect stream habitat, such as: riffles, runs, pools, ditches, wetlands, dams, riprap,
outfalls, tributaries, landscape features, logging paths, vegetation, and roads.
-------
MACROINVERTEBRATES AND HABITAT
X
1
t
0) <
O L
O
^,
1
£
R
CD
c?
Q_
'•"'
R
CD
§
a.
)
R
o
c?
D-
i
1
;
IS
CD
C?
Q-
•5 to! ?
•g si . B
£
OJ CM
Streambank and Channel Charac
(a) Approximate depth of run(s):
Q
(b) Approximate depth of pool(s):
Q <1ft Q 1-2 ft Q >
Approximate width of stream channel:
feet Q measured Q estimated
O T-
jj
a
I a
^Bi
P
••;
B?
fe (A
^••B V^
)
In-Stream Characteris
Check which stream habitats are presen
(You can check more than 1 habitat)
Q Pool(s) Q Riffle(s) Q R
I
^
0)
D)
w
£.
B
'co
to
Nature of particles in the stream bottom
CN
i
£
I
'o
_<2
a
£
(0
Cs
Silt/Clay/Mud _
Sand (up to 0.1 " in diam.)
i« s £ ^1
Looking upstream (100 yds.), pick the descript
best fits the shape of the stream bank and the
^~*
cc
'•a
CM
1
(5
CO
*~*
• E
_ra
b
T—
1
CM
- co
£
i
CO
£
Left Rig!
Q Vertical/undercut Q
5?
Boulders (over 10" in diam.)
Bedrock (solid)
TOTAL 1
Q Steeply sloping (> 30°) Q
Q Gradual/no slope (< 30°) Q
(b) Extent of artificial bank modifications:
Left Rig
Q Bank 0-25% covered Q
^__^
i^
0>
D>
ffl
a.
o i
*Z I
Z?
o
c£ T3
in Q
— ' •§
T3 CD
I E
CD ®
5"2 E 75
S E ^ "o.
Q Bank 25-50% covered Q
Q Bank 50-75% covered Q
Q Bank 75-1 00% covered Q
(c) Shape of the channel:
- - Q Narrow, deep • — - Q Wide, dee[
Q Narrow, shallow Q Wide, shal
^_^
i^
CD
D)
CO
Q.
"S«0 » E
u 2 _ o o
m *5 TT ^> O
Pick the category that best describes th<
which gravel, cobbles, and boulders on
bottom are embedded (sunk) in silt, sani
Q Somewhat/not embedded (0-25%) Q 1
Q Halfway embedded (50%) Q i
CO
Streambank sinks beneath your feet in:
*
%
Q.
CO
>s
CO
a
i
CO
1
a
03
0
a
,
r^
8.
co
a.
E
c
:
•
i
!
J5
'•p*
CD
0- '
a
Presence of logs or large woody debris i
Q None Q Occasional
in
___
i»^
0)
a>
CO
a.
Looking upstream (100 yds.), describe the
m
T"
streamside cover
(a) Along water's edge and stream bank only:
__
f^.
CD
O)
CO
0-
1
_co
CD
*
i
CD
0.
a
Presence of naturally-occurring organic
(i.e., leaves and twigs, etc.) in stream:
Q None Q Occasional
CO
£
D>
ir
-§
TO
of
CO «
CD •£
CD 2>
H CO
|
f—
CD .22
C
D) D)
CO CB
D- QL
CD Q)
C j^ n
z5
D a
Water odor:
Q Sewage Q Fishy
Q Chlorine Q Rotten eggs
Water temperature:
°C or
CO O>
-------
MACROINVERTEBRATES AND HABITAT
*•
rs
to
s
CO
1
"•
CO
,* p-
3 i.
3
3
>^
>»
rt
•a c £&
V) tO ^ o
2; *i = Ss H
•;
•
§ &
n> .(D
jcal Watershed Chan
ut 1/4 mile of the site; adj
-J i
cv
>
12 S
£ *
>i
VI
J-
the local watershed can pot
a stream. Check "1" if presen
act on the stream.
e c
: Q.
Residential
Single-family housing
Multifamily housing
Lawns
Commercial/institutional
Roads, etc.
Paved roads or bridges
Unpaved roads
£
,*~ ^
.. CO D) CO
c Q- c "O
0 £ =6 ^ D>
«" S o ^ § '55
5i -_i c ts — C o
C c fc o ca ca c
fill ! i ft?
^cuo i-^ „, -i "CD =
||^S_-a°|c.f | w | '% ^
Illf
oooo o>2o,2co Sooiog^ £ .S P> -a — £5 S
OIODC
"E EC
§>
^\
v?
o
re top of the streambank out
eft (Percent)
Trees
is —t
u.
3)
W
1
§
m
^;
'a
£
2
cr
1
1 c
5
Boulders/rocks
Gravel/sand
C
o 3
r
0)
><
a>
CM O O
r- a a
fp
c
t
c
Q
5?
o
o
a
•
• vJ3 «
o> & £o o
£ 'to Is- S
cNDaa oioaoo cvaaaaa cMaoaaao
T-aaa i-oaao i-oooaa i-aooaoa
) ^
J D) «M CM
T-oaaa i-aa i-aaaaa . .,.
•£ CO •
CD £
2 fc
^ C '""*'
^ a> = 33 is - co
CO CO r- fn CO CD
^ D) S m ro "
u 03 5S ?• ^ S3 '
co "o £ ^ "o. £
'~ »- "55 D) .9- "g «
i § CD .£ o IS £
£2 O"§ " o "ceo SSw'sr'CDco
™>> Og ^T3*-J«: CD CD ScuS-^S
O "Jji ^j; Q- (Q QJ "ti c w .?? O) i~ •"— " -«-—
JP sP £ Q> "5."amCO OCO Ci-§Q.p>CO
•SlgSI ^§Sc .55 l^cfl
3Ji S -^ «p ¥2 11 .i |lf {? •: -
£S 1* = 1 1= S s ?^ § o|J|
5^ c^ « £ J .2. •§ fi o .2. S ~ -S2 o •£
S tl ^ - *? tnwff'3fv:.0--"Sf ^-^^Q-a3
i mil inn lit mill
S •§ w g.,2- wzcQOu: w2o o>2 5^ 2
0 *^ O
) Q
* U —J
CD
r-
^~;t±cNOnOO CNDD oiDDDOD
ij cij
03 ^J ^^^ ^^ ^^ ^^
Jd T~ D D D D T" D O T" O O D O n
O
-------
I §.
MACROINVERTEBRATES AND HABITAT
o 1
O c:
o
i ^
^^
•I
£l
ff
^
ii
n
m
T
9
i
0
D "aT
i3 §
. o
— I ca
el ™
? .£
^} <^
®
co S*
^ 3
a a
s. 1 .E
uj 8- | «P
> -^ 2 £2.
tt | | ! |
1 n\ la m~ '"5
ii
5 D D
£2 CO
^ T-
CO
h-
—
0-
"52
movemel
there any barriers to fish
£
4
0>
<2 CD
I 5
a o
f-~
3. i
i 1
Beaver dams Q Waterf
Dams Q Road I
D 0
CO
N.
0)
c?
a.
S^
1
^
"w
*
t
atic plants in the stream.
o-
o
CM
tb
~S i£
'•5 w
§ .§
Q. Z
O D
O)
« .£
.1 1
CO ^ ,-
«B en —
S 1 1
a a o
c
'S>
CO
•o e
J E
CD O CO
C CO CD
0 *3 i:
Z < CO
o o a
CO
CD
§
I
"CO
••-*
15
•JS
I
mt of algae in the stream,
0}
UJ
T-
CM
-
0)
_c
."2
'w _
0) 13
11 S
= {>• CD £
S> Eo, s: o
|| o a
•- sw
Si _ 1
"> fS CO CO
& t- C. O ^
„ O o o u>
re the submerged stones
ream coated with a layer
None Q Occas
Light coating Q Heavy
Brownish Q Greeni
< w 0 0 a
"5T
"~"
1
: 1
^' "£ ^
s ^ ~
§> Q- 0
^ a a
O
D)
•1 -a
£.: .1 .i
0)
O
O)
.s
_o
^~ 13
« '•£ CD
m CD £
CO Q. O
? DO
"(/)
1
>. "Jo
0 n ^z
5 O OD
,«• w c w co c
:£: CO
-------
4.3
Intensive Stream
Biosurvey
The Intensive Stream Biosurvey is
based on the habitat assessment and macro-
invertebrate sampling approach developed
by EPA in its Rapid Bioassesstnent Proto-
cols/or Streams and Rivers (Protocol II)
and adapted by volunteer monitoring
programs such as Maryland Save Our
Streams and River Watch Network.
Like the Stream Habitat Walk and
Streamside Biosurvey, this approach
includes a study of macroinvertebrates and
habitat. However, the Intensive Stream
Biosurvey approach is more rigorous; it
requires substantial volunteer training in
habitat and macroinvertebrate sampling
methods and in macroinvertebrate identifi-
cation. This approach also requires the
involvement of a stream biologist to advise
the program participants regarding every-
thing from the selection of reference
conditions to taxonomy and data analysis.
Because of the need for training and
professional assistance, the Intensive
Stream Biosurvey approach can be expen-
sive and labor-intensive for the volunteer
program. Its benefits, however, are equally
clear: with proper quality control and
volunteer training, the Intensive Stream
Biosurvey can yield credible information
on subtle stream impacts and water quality
trends. Key features of the Intensive Stream
Biosurvey are as follows:
• It relies on comparing the results for
the sampling site to regional or local
reference conditions. This type of
study is used to determine how
streams in a given area compare to
the best possible conditions. The
reference condition is a composite of
the best attainable (minimally
impaired) stream conditions within
the region and should be determined
by an experienced aquatic biologist
familiar with the characteristics of
the ecological region.
It includes a detailed habitat assess-
ment that requires the volunteer to
rate 10 parameters on a scale ofOto
20. The results of the habitat assess-
ment are compared to the score
received by the stream's reference
condition, and a percent similarity
score is calculated.
The methods for collecting macroin-
vertebrates are similar to those of the
Streamside Biosurvey. However,
rather than being processed stream-
side, the entire sample of
macroinvertebrates is preserved and
returned to a laboratory. A portion,
or subsample, of the total organisms
collected at each location is ran-
domly selected and identified to
taxonomic family level in the lab.
After identification, a series of
indices (or metrics) are calculated to
provide a broad range of information
about the stream site. The subsample
and the rest of the collected organ-
isms are maintained as a voucher
collection, which serves as a quality
assurance component.
The Intensive Stream Biosurvey
requires that volunteers be exten-
sively trained before habitat
assessment and macroinvertebrate
sampling and before attempting
macroinvertebrate identification in
the laboratory. An experienced
aquatic biologist is needed to deter-
mine and evaluate the regional
reference conditions; train volunteers
in habitat characteristics; and super-
vise and train volunteers in the
collection, processing, and identifica-
tion of sample macroinvertebrates. A
laboratory (with microscopes) and a
macroinvertebrate sample storage
facility are required.
-------
MACROINVERTEBRATES AND HABITAT
Step 1—Prepare for the Intensive
Stream Biosurvey field work
Preparing for the Intensive Stream
Biosurvey might take several months from
the initial planning stages to the time when
actual sampling occurs. An aquatic biolo-
gist should be centrally involved in all
aspects of technical program development.
Issues that should be considered in
planning the program include the follow-
ing:
• Availability of reference conditions
for your area
• Appropriate dates to sample in each
season
• Appropriate sampling gear
• Sampling station location
• Availability of laboratory facilities
and trainers
• Sample storage
• Data management
• Appropriate taxonomic keys,
metrics, or measurements for macro-
invertebrate analysis
• Habitat assessment consistency
Some of the preparation work for this
approach is similar to that of the Stream
Habitat Walk (section 4.1) and Streamside
Biosurvey (section 4.2). Refer back to .those
sections for relevant information on the
following tasks:
• Obtaining a USGS topographical
map
• Becoming familiar with safety
procedures
TASK1
Select monitoring locations
If possible, the program coordinator, in
conjunction with technical advisor(s),
should preselect sampling locations for
each stream. This adds an element of
quality control to the sampling process.
You might want to consider sampling at a
few locations that are also sampled by state
or local professionals, as a way to compare
your results to theirs. Be sure to secure
approval to do so, however, and coordinate
your sampling so as not to affect profes-
sional results.
Provide detailed hand-drawn maps of
the locations selected to the monitors.
Know the latitude and longitude of your
monitoring locations. This is critical for
mapping and for many data management
programs. Latitude and longitude can be
calculated manually (see Appendix C) or
by using a hand-held Global Positioning
System (GPS).
TASK 2 | Schedule the field portion of
"""""™~^™^ the biosurvey
Schedule your Intensive Stream
Biosurvey for a time of year for which
reference conditions have been established.
Reference conditions might vary by season.
It is also essential that seasonal data be
collected within the same index period, or
window of time, each year. In other words,
if you sample during the last two weeks of
March this year, do the same next year.
Another factor to keep in mind is
*• i
weather. It is best to wait at least a week
after a heavy rain or snow event before
sampling. Heavy rains can have a scouring
effect on macroinvertebrates, :washing them
downstream. If this happens, samples
collected will not accurately reflect biologi-
cal conditions. However, if you are study-
ing the possible impact of runoff from a
particular source (such as a construction
site), you might decide to sample within a
short time after heavy precipitation.
TASK 3 I Gather tools and equipment
for the Intensive Stream
Biosurvey
In addition to the basic sampling
equipment listed for the Stream Habitat
Walk, collect the following equipment
needed for the macroinvertebrate collection
and habitat assessment of the Intensive
Stream Biosurvey:
-------
MACROINVERTEBRATES AND HABITAT
Jars (2, at least quart size), plastic,
wide-mouth with tight cap; one
should be empty and the other filled
about two thirds full with 70 percent
ethyl alcohol. (Jars can be purchased
from a scientific supply company or
you might try using large pickle,
mayonnaise, or quart mason jars.)
Hand lens, magnifying glass, or field
microscope
Fine-point forceps
Heavy-duty rubber gloves (kitchen
gloves will work fine)
Plastic sugar scooper or ice-cream
scooper
Kick net (rocky bottom stream) or
dip net (muddy bottom stream) (see
Fig. 4.7, page 63)
Buckets (2)
String or twine (50 yards); tape
measure
Stakes (4)
Orange (a stick, an apple, or a fish
float may also be used in place of an
orange) to measure velocity
Reference maps indicating general
information pertinent to the sampling
area, including the surrounding
roadways, as well as hand-drawn
station map
Station ID tags
Spray water bottle
Pencils (at least 2)
Sieve Buckets
Most professional biological monitoring programs employ sieve
buckets as a holding container for composited samples. These buckets
have a mesh bottom that allows water to drain out while the organisms
and debris remain. This material can then be
easily transferred to the alcohol-filled jars.
However, sieve buckets can be expen-
sive. Many volunteer programs employ
alternative equipment, such as the two
regular buckets described in this section.
Regardless of the equipment, the
process for compositing and transferring
the sample is basically the same. The
decision is one of cost and convenience.
TASK 4
Become familiar with field
data sheets and instructions/
definitions for conducting
the macroinvertebrate
collection and Habitat
Assessment portions of the
Intensive Biosurvey
Step 2—Conduct the Intensive
Biosurvey field work
The method you use to collect macroin-
vertebrates using this approach depends on
the type of stream you are sampling.
Rocky-bottom streams are defined as
those with bottoms made up of gravel,
cobbles, and boulders in any combination.
They usually have definite riffle areas.
Riffle areas are fairly well oxygenated and,
therefore, are prime habitats for benthic
macroinvertebrates. In these streams, use
the Rocky-Bottom sampling method.
Muddy-bottom streams have muddy,
silty, or sandy bottoms that lack riffles.
Usually, these are slow-moving, low-
gradient streams (i.e., streams that flow
along flat terrain). In such streams, macro-
invertebrates generally attach to overhang-
ing plants, roots, logs, submerged vegeta-
tion, and stream substrate where organic
particles are trapped. In these streams, use
the Muddy Bottom sampling method.
Each method is detailed below. Regard-
less of which collection method is used, the
process for counting, identifying, and
analyzing the macroinvertebrate sample for
the Intensive Stream Biosurvey is the same.
Following the discussion of both ap-
proaches to macroinvertebrate collection
and habitat assessment procedures is a
section on analyzing the sample.
-------
MACROINVERTEBRATES AND HABITAT I 89
Rocky-Bottom Streams
Part 1: Macrpinvertebrate
Sampling Method
Use the following method of macroin-
vertebrate sampling in streams that have
riffles and gravel/cobble substrates. You
will collect three samples at each site and
composite them to obtain one large total
sample.
TASK1
Identify the sampling
location
You should already have located your
site on a map along with its latitude and
longitude (see Task 3, page 45)
1. You are going to sample in three
different spots within a 100-yard
stream site. These spots may be three
separate riffles; one large riffle with
different current velocities; or, if no
riffles are present, three run areas
with gravel or cobble substrate.
Combinations are also possible (if,
for example, your site has only one
small riffle and several run areas).
Mark off your 100-yard stream
site. If possible, it should begin at
least 50 yards upstream of any
human-made modification of the
channel, such as a bridge, dam, or
pipeline crossing, Avoid walking in
the stream, since this might dislodge
macroinvertebrates and alter your
sampling results.
2. Sketch the 100-yard sampling area.
Indicate the location of your three
sampling spots on the sketch. Mark
the most downstream site as Site 1,
the middle site as Site 2, and the
upstream site as Site 3. (See Fig.
4.8.)
TASK 2 | Get into place
and sample the site
farthest down-
stream first (Site 1).
This keeps you from
biasing your second
and third collections
with dislodged
sediment or macro-
invertebrates.
Always use a clean
kick-seine, relatively
free of mud and
debris from previous
uses. Fill a bucket
about one third full
with stream water
and fill your spray
bottle.
2. Select a 3-foot by 3-
foot riffle area for
sampling at Site 1.
One member of the
team, the net holder,
.should position the
net at the down-
stream end of this
sampling area. Hold
the net handles at a
45 degree angle to
the water's surface.
Be sure that the
bottom of the net fits
tightly against the
streambed so no
macroinvertebrates
escape under the net.
You may use rocks
from the sampling
area to anchor the
net against the
stream bottom.
Don't allow any
water to flow over
the net.
1. Always approach your sampling
locations from the downstream end
1. Approach the sample
site from the down-
stream end.
2. Position the net at a 45°
angle with the bottom
tight against the sub-
strate.
3. Dislodge macroinverte-
brates by rubbing rocks
thoroughly.
TASK 3 | Dislodge the
macroinver-
tebrates
-------
4. Disturb the substrate
thoroughly with your
feet.
5. Remove the net with a
forward scooping motion.
6. Flush out the net with
clean stream water.
1. Pick up any large
rocks in the 3-foot
by 3-foot sampling
area and rub them
thoroughly over the
partially-filled
bucket so that any
macroinvertebrates
clinging to the rocks
will be dislodged
into the bucket. Then
place each cleaned
rock outside of the
sampling area. After
sampling is com-
pleted, rocks can be
returned to the
stretch of stream
they came from.
2. The member of the
team designated as
the "kicker" should
thoroughly stir up
the sampling area
with their feet,
starting at the
upstream edge of the
3-foot by 3-foot
sampling area and
working down-
stream, moving
toward the net. All
dislodged organisms
will be carried by the
stream flow into the
net. Be sure to
disturb the first few
inches of stream
sediment to dislodge
burrowing organ-
isms. As a guide,
disturb the sampling
area for about 3
minutes, or until the
area is thoroughly
worked over.
3. Any large rocks used to anchor the
net should be thoroughly rubbed into
the bucket as above.
TASK 4
Remove the net
1
Next, remove the net without allow-
ing any of the organisms it contains
to wash away. While the net holder
grabs the top of the net handles, the
kicker grabs the bottom of the net
handles and the net's bottom edge.
Remove the net from the stream with
a forward scooping motion.
2. Roll the kick net into a cylinder
shape and place it vertically in the
partially filled bucket. Pour or spray
water down the net to flush its
contents into the bucket. If neces-
sary, pick debris and organisms from
the net by hand. Release back into
the stream any fish, amphibians, or
reptiles caught in the net.
TASK 5 I Collect the second and third
samples
Once you have removed all the organ-
isms from the net repeat these steps at Sites
2 and 3. Put the samples from all three sites
into the same bucket. Combining the debris
and organisms from all three sites into the
same bucket is called compositing.
Hint: If your bucket is nearly full of water
after you have washed the net clean, let
the debris and organisms settle to the
bottom of the bucket. Then cup the net
over the bucket and pour the water through
the net into a second bucket. Inspect the
water in the second bucket to be sure no
organisms came through.
TASK 6 I Preserve the sample
1. After collecting and compositing all
three samples, it is time to preserve
the sample. All team members
-------
MACROINVERTEBRATES AND HABITAT
should leave the stream and return to
a relatively flat section of stream
bank with all their equipment. The
next step will be to remove large
pieces of debris (leaves, twigs, and
rocks) from the sample. Carefully
remove the debris one piece at a
time. While holding the material over
the bucket, use the forceps, spray
bottle, and your hands to pick, rub,
and rinse the leaves, twigs, and rocks
to remove any attached organisms.
Use your magnifying lens and
forceps to find and remove small
organisms clinging to the debris.
When you are satisfied that the
material is clean, discard it back into
the stream.
2. You will need to drain off the water
before transferring material to the jar.
This process will require two team
members. Place the kick net over the
second bucket, which has not yet
been used and should be completely
empty. One team member should
push the center of the net into bucket
#2, creating a small indentation or
depression. Then, hold the sides of
the net closely over the mouth of the
bucket. The second person can now
carefully pour the remaining contents
of bucket #1 onto a small area of the
net to drain the water and concentrate
the organisms. Use care when
pouring so that organisms are not lost
over the side of the net (Fig. 4.16).
Use your spray bottle, forceps,
sugar scoop, and gloved hands to
remove all the material from bucket
#1 onto the net. When you are
satisfied that bucket #1 is empty, use
your hands and the sugar scoop to
transfer all the material from the net
into the empty jar.
Bucket #2 captured the water and
any organisms that might have fallen
through the netting during pouring.
As a final check, repeat the process
above, but this time, pour bucket #2
over the net, into bucket #1. Transfer
any organisms on the net into the jar.
3. Now, fill the jar (so that all material
is submerged) with the alcohol from
the second jar. Put the lid tightly
back onto the jar and gently turn the
jar upside down two or three times to
distribute the alcohol and remove air
bubbles. \
4. Complete the Sampling Station ID
tag. Be sure to use a pencil, not a
pen, because the ink will run in the
alcohol! The tag includes your
station number, the stream, location
(e.g., upstream from a road cross-
ing), date, time, and the'names of the
members of the collecting crew.
Place the ID tag into the sample
container—writing side facing out,
so that identification can be seen
clearly. :
Fig. 4.16
Pouring
sample water
through the net
-------
Rocky-Bottom Streams
Part 2: Habitat Assessment Method
You will conduct a habitat assessment
(which will include measuring general
characteristics and local land use) in a 100-
yard section of stream that includes the
riffles from which organisms were col-
lected.
TASK 1 I Delineate the habitat assess-
~"™^™~™^ ment boundaries
1. Begin by identifying the most
downstream riffle that was sampled
for macroinvertebrates. Using your
tape measure or twine, mark off a
100-yard section extending 25 yards
below the downstream riffle and
about 75 yards upstream.
2. Complete the identifying information
on your field data sheet for your
habitat assessment site. On your
stream sketch, be as detailed as
possible, and be sure to note which
riffles were sampled.
TASK 2
Complete the General
Characteristics and Local
Land Use sections of the
field sheet
For safety reasons as well as to protect
the stream habitat, it is best to estimate
these characteristics rather than actually
wading into the stream to measure them.
General Characteristics
1. Water appearance can be a physical
indicator of water pollution.
• Clear - colorless, transparent
• Milky - cloudy-white or grey, not
transparent; might be natural or
due to pollution
• Foamy - might be natural or due
to pollution, generally detergents
or nutrients (foam that is several
inches high and does not brush
apart easily is generally due to
pollution)
• Turbid - cloudy brown due to
suspended silt or organic material
• Dark brown - might indicate that
acids are being released into the
stream due to decaying plants
• Oily sheen -multicolored reflec-
tion might indicate oil floating in
the stream, although some sheens
are natural
• Orange - might indicate acid
drainage
• Green - might indicate excess
nutrients being released into the
stream
2. Water odor can be a physical indica-
tor of water pollution.
• None or natural smell
• Sewage - might indicate the
release of human waste material
• Chlorine - might indicate that a
sewage treatment plant is over-
chlorinating its effluent
• Fishy - might indicate the pres-
ence of excessive algal growth or
dead fish
• Rotten eggs - might indicate
sewage pollution (the presence of
a natural gas)
3. Water temperature can be particu-
larly important for determining
whether the stream is suitable as
habitat for some species of fish and
macroinvertebrates that have distinct
temperature requirements. Tempera-
ture also has a direct effect on the
amount of dissolved oxygen avail-
able to aquatic organisms. Measure
temperature by submerging a ther-
mometer for at least 2 minutes in a
typical stream run. Repeat once and
average the results.
-------
MACROINVERTEBRATES AND HABITAT
4. The width of the stream channel can
be determined by estimating the
width of the streambed that is
covered by water from bank to bank.
If it varies widely along the stream,
estimate an average width.
Local Land Use
5. Local land use refers to the part of
the watershed within 1/4 mile up-
stream of and adjacent to the site.
Note which land uses are present, as
well as which ones seem to be
having a negative impact on the
stream. Base your observations on
what you can see, what you passed
on the way to the stream, and, if
possible, what you notice as you
leave the stream.
TASKS
Conduct the habitat assess-
ment
The following information describes
the parameters you will evaluate for rocky-
bottom habitats. Use these definitions when
completing the habitat assessment field data
sheet.
The first two parameters should be
assessed directly at the riffle(s) or run(s)
that were used for the macroinvertebrate
sampling.
1. Attachment sites for macroinverte-
brates are essentially the amount of
living space or hard substrates
(rocks, snags) available for aquatic
insects and snails. Many insects
begin their life underwater in streams
and need to attach themselves to
rocks, logs, branches, or other
submerged substrates. The greater
the variety and number of available
living spaces or attachment sites, the
greater the variety of insects in the
stream. Optimally, cobble should
predominate and boulders and gravel
should be common. The availability
of suitable living spaces for macroin-
vertebrates decreases as cobble
becomes less abundant and boulders,
gravel, or bedrock become more
prevalent.
2. Embeddedness refers to the extent to
which rocks (gravel, cobble, and
boulders) are surrounded by, cov-
ered, or sunken into the silt, sand, or
mud of the stream bottom. Gener-
ally, as rocks become embedded,
fewer living spaces are available to
macroinvertebrates and fish for
shelter, spawning and egg incuba-
tion.
To estimate the percent of
embeddedness, observe the amount
of silt or finer sediments overlying
and surrounding the rocks. If kicking
does not dislodge the rocks or
cobbles, they might be greatly
embedded.
The following eight parameters should
be assessed in the entire 100-yard section
of the stream.
3. Shelter for fish includes the relative
quantity and variety of natural
structures in the stream, such as
fallen trees, logs, and branches;
cobble and large rocks; and undercut
banks that are available to fish for
hiding, sleeping, or feeding. A wide
variety of submerged structures in
the stream provide fish with many
living spaces; the more living spaces
in a stream, the more types of fish
the stream can support.
4. Channel alteration is basically a
measure of large-scale changes in
the shape of the stream channel.
Many streams in urban and agricul-
tural areas have been straightened,
deepened (e.g., dredged), or diverted
into concrete channels, often for
flood control purposes. Such streams
have far fewer natural habitats for
fish, macroinvertebrates, land plants
than do naturally meandering
-------
streams. Channel alteration is present
when the stream runs through a
concrete channel; when artificial
embankments, riprap, and other
forms of artificial bank stabilization
or structures are present; when the
stream is very straight for significant
distances; when dams, bridges, and
flow-altering structures such as
combined sewer overflow (CSO)
pipes are present; when the stream is
of uniform depth due to dredging;
and when other such changes have
occurred. Signs that indicate the
occurrence of dredging include
straightened, deepened, and other-
wise uniform stream channels, as
well as the removal of streamside
vegetation to provide dredging
equipment access to the stream.
5. Sediment deposition is a measure of
the amount of sediment that has been
deposited in the stream channel and
the changes to the stream bottom that
have occurred as a result of the
deposition. High levels of sediment
deposition create an unstable and
continually changing environment
that is unsuitable for many aquatic
organisms.
Sediments are naturally deposited
in areas where the stream flow is
reduced, such as pools and bends, or
where flow is obstructed. These
deposits can lead to the formation of
islands, shoals, or point bars (sedi-
ments that build up in the stream,
usually at the beginning of a mean-
der) or can result in the complete
filling of pools. To determine
whether these sediment deposits are
new, look for vegetation growing on
them: new sediments will not yet
have been colonized by vegetation.
6. Stream velocity and depth combina-
tions are important to the
maintenance of healthy aquatic
communities. Fast water increases
the amount of dissolved oxygen in
the water; keeps pools from being
filled with sediment; and helps food
items like leaves, twigs, and algae
move more quickly through the
aquatic system. Slow water provides
spawning areas for fish and shelters
macroinvertebrates that might be
washed downstream in higher stream
velocities. Similarly, shallow water
tends to be more easily aerated (i.e.,
it holds more oxygen), but deeper
water stays cooler longer. Thus the
best stream habitat includes all of the
following velocity/depth combina-
tions and can maintain a wide variety
of organisms.
slow (<1 ft/sec), shallow (<1.5 ft)
slow, deep
fast, deep
fast, shallow
Measure stream velocity by
marking off a 10-foot section of
stream run and measuring the time it
takes a stick, orange, or other float-
ing biodegradable object to float the
10 feet. Repeat 5 times, in the same
10-foot section, and determine the
average time. Divide the distance (10
feet) by the average time (seconds) to
determine the velocity in feet per
second.
Measure the stream depth by
using a stick of known length and
taking readings at various points
within your stream site, including
riffles, runs, and pools. Compare
velocity and depth at various points
within the 100-yard site to see how
many of the combinations are
present.
Channel flow status is the percent of
the existing channel that is filled with
water. The flow status changes as the
channel enlarges or as flow decreases
-------
MACROINVERTEBRATES AND HABITAT
as a result of dams and other obstruc-
tions, diversions for irrigation, or
drought. When water does not cover
much of the streambed, the living
area for aquatic organisms is limited.
For the last three parameters, evaluate
the condition of the right and left stream
banks separately. Define the " left" and
"right" banks by standing at the down-
stream end of your study stretch and
looking upstream. Each bank is evaluated
on a scale of 0-10.
8. Bank vegetative protection measures
the amount of the stream bank that is
covered by natural (i.e., growing
wild and not obviously planted)
vegetation. The root systems of
plants growing on stream banks help
hold soil in place, reducing erosion.
Vegetation on banks provides shade
for fish and macroinvertebrates and
serves as a food source by dropping
leaves and other organic matter into
the stream. Ideally, a variety of
vegetation should be present, includ-
ing trees, shrubs, and grasses.
Vegetative disruption can occur
when the grasses and plants on the
stream banks are mowed or grazed,
or when the trees and shrubs are cut
back or cleared.
9. Condition of banks measures erosion
potential and whether the stream
banks are eroded. Steep banks are
more likely to collapse and suffer
from erosion than are gently sloping
banks and are therefore considered to
have a high erosion potential. Signs
of erosion include crumbling,
unvegetated banks, exposed tree
roots, and exposed soil.
10. The riparian vegetative zone width is
defined here as the width of natural
vegetation from the edge of the
stream bank. The riparian vegetative
zone is a buffer zone to pollutants
entering a stream from runoff. It also
controls erosion and provides stream
habitat and nutrient input into the
stream.
A wide, relatively undisturbed
riparian vegetative zone .reflects a
healthy stream system; narrow, far
less useful riparian zones occur
when roads, parking lots; fields,
lawns, and other artificially culti-
vated areas, bare soil, rocks, or
buildings are near the stream bank.
The presence of "old fields" (i.e.,
previously developed agricultural
fields allowed to revert to natural
conditions) should rate higher than
fields in continuous or periodic use.
In arid areas, the riparian vegetative
zone can be measured by observing
the width of the area dominated by
riparian or water-loving plants, such
as willows, marsh grasses, and
cotton wood trees.
Note: Instructions on sample processing,
macroinvertebrate identification, and data
analysis follow the sections on muddy-
bottom macroinvertebrate sampling and
habitat assessment. (See Step 3, page
101)
Muddy-Bottom Sampling
Part 1: Macroinvertebrate Sampling
In muddy-bottom streams, as in rocky-
bottom streams, the goal is to sample the
most productive habitat available and look
for the widest variety of organisms. The
most productive habitat is the one that
harbors a diverse population of pollution-
sensitive macroinvertebrates. Volunteers
should sample by using a D-frame net to
jab at the habitat and scoop up the organ-
isms that are dislodged. The idea is to
collect a total sample that consists of 20
jabs taken from a variety of habitats.
-------
MACRO1NVERTEBRATES AND HABITAT
TASK 1 I Determine which habitats are
present
Figure 4.17
Four habitats
found in
muddy-bottom
streams
Volunteers will
likely find the
most macroin-
vertebrates in
vegetated
habitats and
snags and logs.
Muddy-bottom streams usually have
four habitats (Fig. 4.17). It is generally best
to concentrate sampling efforts on the most
productive habitat available, yet to sample
other principal habitats if they are present.
This ensures that you will secure as wide a
variety of organisms as possible. Not all
habitats are present in all streams or present
in significant amounts. If your sampling
areas have not been preselected, try to
determine which of the following habitats
are present. (Avoid standing in the stream
while making your habitat determinations.)
• Vegetated bank margins consist of
overhanging bank vegetation and
submerged root mats attached to
banks. The bank margins may also
contain submerged, decomposing
leaf packs trapped in root wads or
lining the streambanks. This is
generally a highly productive habitat
in a muddy-bottom stream, and it is
often the most abundant type of
habitat.
Snags and logs consist of submerged
wood, primarily dead trees, logs,
branches, roots, cypress knees and
leaf packs lodged between rocks or
logs. This is also a very productive
muddy-bottom stream habitat.
Aquatic vegetation beds and decay-
ing organic matter consist of beds of
submerged, green/leafy plants that
are attached to the stream bottom.
This habitat can be as productive as
vegetated bank margins, and snags
and logs.
Silt/sand/gravel substrate includes
sandy, silty, or muddy stream
bottoms; rocks along the stream
bottom; and/or wetted gravel bars.
This habitat may also contains algae-
covered rocks (sometimes called
Aufwuchs). This is the least produc-
tive of the four muddy-bottom
stream habitats, and it is always
present in one form or another (e.g.,
silt, sand, mud, or gravel might
predominate).
TASK 2
Determine how many times
to jab in each habitat type
Your goal is to jab a total of 20 times.
The D-frame net is 1 foot wide, and a jab
should be approximately 1 foot in length.
Thus, 20 jabs equals 20 square feet of
combined habitat.
• If all four habitats are present in
plentiful amounts, jab the vegetated
banks 10 times and divide the
remaining 10 jabs among the remain-
ing 3 habitats.
-------
MACROINVERTEBRATES AND HABITAT
• If three habitats are present in
plentiful amounts and one is absent,
jab the silt/sand/gravel substrate—
the least productive habitat—5 times
and divide the remaining 15 jabs
among the other two more produc-
tive habitats.
• If only two habitats are present in
plentiful amounts, the silt/sand/
gravel substrate will most likely be
one of those habitats. Jab the silt/
sand/gravel substrate 5 times and the
more productive habitat 15 times.
• If some habitats are plentiful and
others are sparse, sample the sparse
habitats to the extent possible, even
if you can take only one or two jabs.
Take the remaining jabs from the
plentiful habitat(s). This rule also
applies if you cannot reach a habitat
because of unsafe stream conditions.
Jab a total of 20 times.
Because you might need to make an
educated guess to decide how many jabs to
take in each habitat type, it is critical that
you note, on the field data sheet, how many
jabs you took in each habitat. This informa-
tion can be used to help characterize your
findings.
After every few jabs, the sampler should
hand the net to the second person, who then
can rinse the contents of the net into the
bucket.
TASK3 I Get into place
Outside and downstream of your first
sampling location (1st habitat), rinse the dip
net and check to make sure it does not
contain any macroinvertebrates or debris
from the last time it was used. Fill a bucket
approximately one-third full with clean
stream water. Also, fill the spray bottle with
clean stream water. This bottle will be used
to wash down the net between jabs and
after sampling is completed.
This method of sampling requires only
one person to disturb the stream habitats.
While one person is sampling, a second
person should stand outside the sampling
area, holding the bucket and spray bottle.
TASK 4
Dislodge the macroinverte-
brates
Approach the first sample site from
downstream, and sample as you walk
upstream. Here is how to sample in the four
habitat types:
• Sample vegetated bank margins by
jabbing vigorously, with an upward
motion, brushing the net against
vegetation and roots along the bank.
The entire jab motion should occur
underwater.
• To sample snags and logs, hold the
net with one hand under the section
of submerged wood you are sam-
pling (Fig. 4.18). With the other
hand (which should be gloved), rub
about 1 square foot of area on the
snag or log. Scoop organisms, bark,
twigs, or other organic matter you
dislodge into your net. Each combi-
nation of log rubbing and net
scooping is one jab.
• To sample aquatic vegetation beds,
jab vigorously, with an upward
motion, against or through the plant
bed. The entire jab motion should
occur underwater.
• To sample a silt/sand/gravel sub-
strate, place the net with one edge
against the stream bottom and push
it forward about a foot (in an up-
stream direction) to dislodge the first
few inches of silt, sand,.gravel, or
rocks. To avoid gathering a netful of
mud, periodically sweep the mesh
bottom of the net back and forth in
the water, making sure that water
does not run over the top of the net.
This will allow fine silt to rinse out
of the net. When you have com-
-------
Figure 4.18
Collecting a
sample from a
log
Volunteer rubs
the log with one
hand and
catches dis-
lodged organ-
isms and other
material in the
net.
pleted all 20 jabs, rinse the net
thoroughly into the bucket. If neces-
sary, pick any clinging organisms
from the net by hand and put them in
the bucket.
TASK 5 | Preserve the sample
1. Look through the material in the
bucket and immediately return any
fish, amphibians, or reptiles to the
stream. Carefully remove large
pieces of debris (leaves, twigs, and
rocks) from the sample. While
holding the material over the bucket,
use the forceps, spray bottle, and
your hands to pick, rub, and rinse the
leaves, twigs, and rocks to remove
any attached organisms. Use your
magnifying lens and forceps to find
and remove small organisms clinging
to the debris. When you are satisfied
that the material is clean, discard it
back into the stream.
2. You will need to drain off the water
before transferring material to the jar.
This process will require two team
members. One person should place
the net into the second bucket, like a
sieve (this bucket, which has not yet
been used, should be completely
empty) and hold it securely. The
second person can now carefully
pour the remaining contents of
bucket #1 onto the center of the net
to drain the water and concentrate the
organisms.
Use care when pouring so that
organisms are not lost over the side
of the net. Use your spray bottle,
forceps, sugar scoop, and gloved
hands to remove all the material from
bucket #1 onto the net. When you are
satisfied that bucket #1 is empty, use
your hands and the sugar scoop to
transfer all the material from the net
into the empty jar. You can also try
to carefully empty the contents of the
net directly into the jar by turning the
net inside out into the jar.
Bucket #2 captured the water and
any organisms that might have fallen
through the netting. As a final check,
repeat the process above, but this
time, pour bucket #2 over the net,
into bucket #1. Transfer any organ-
isms on the net into the jar.
3. Fill the jar (so that all material is
submerged) with alcohol. Put the lid
tightly back onto the jar and gently
turn the jar upside down two or three
times to distribute the alcohol and
remove air bubbles.
4. Complete the sampling station ID
tag. Be sure to use a pencil, not a
pen, because the ink will run in the
alcohol. The tag should include your
station number, the stream, location
(e.g., upstream from a road crossing),
date, time, and the names of the
members of the collecting crew.
Place the ID tag into the sample
container, writing side facing out, so
that identification can be seen clearly
(Fig. 4.19).
-------
MACROINVERTEBRATES AND HABITAT
Muddy-Bottom Streams
Part 2: Habitat Assessment
You will conduct a habitat assessment
(which will include measuring general
characteristics and local land use) in a 100-
yard section of the stream that includes the
habitat areas from which organisms were
collected.
TASK 1 I Delineate the habitat assess-
ment boundaries
1. Begin by identifying the most
downstream point that was sampled
for macroinvertebrates. Using your
tape measure or twine, mark off a
100-yard section extending 25 yards
below the downstream sampling
point and about 75 yards upstream.
2. Complete the identifying information
on your field data sheet for your
habitat assessment site. On your
stream sketch, be as detailed as
possible, and be sure to note which
habitats were sampled.
TASK 2
Complete the General
Characteristics and Local
Land Use sections of the
field sheet
For safety reasons as well as to protect
the stream habitat, it is best to estimate
these characteristics rather than actually
wading into the stream to measure them.
For instructions on completing these
sections of the field data sheet, see the
rocky-bottom habitat assessment instruc-
tions.
TASKS
Conduct the habitat assess-
ment
The following information describes
the parameters you will evaluate for
muddy-bottom habitats. Use these defini-
tions when completing the habitat assess-
ment field data sheet.
STATION ID TAG
Station #:
Stream:
Location:
Date/Time: L_
Team members:
Shelter for fish and attachment sites
for macroinvertebrates are essen-
tially the amount of living space and
shelter (rocks, snags, and undercut
banks) available for fish, insects, and
snails. Many insects attach them-
selves to rocks, logs, branches, or
other submerged substrates. Fish can
hide or feed in these areas. The
greater the variety and number of
available shelter sites or attachment
sites, the greater the variety offish
and insects in the stream.
Many of the attachnient sites
result from debris falling into the
stream from the surrounding vegeta-
tion. When debris first falls into the
water, it is termed new fall and it has
not yet been "broken down" by
microbes (conditioned) for macroin-
vertebrate colonization. Leaf
material or debris that is conditioned
is called old fall. Leaves that have
been in the stream for some time
lose their color, turn brown or dull
yellow, become soft and supple with
Figure 4.19
Example of a
Station ID tag
To prevent
samples from
being mixed up,
volunteers
should place the
IDtag/ns/otethe
sample jar.
-------
100 I MACROINVERTEBRATES AND HABITAT
age, and might be slimy to the touch.
Woody debris becomes blackened or
dark in color; smooth bark becomes
coarse and partially disintegrated,
creating holes and crevices. It might
also be slimy to the touch.
2. Pool substrate characterization
evaluates the type and condition of
bottom substrates found in pools.
Pools with firmer sediment types
(e.g., gravel, sand) and rooted
aquatic plants support a wider variety
of organisms than do pools with
substrates dominated by mud or
bedrock and no plants. In addition, a
pool with one uniform substrate type
will support far fewer types of
organisms than will a pool with a
wide variety of substrate types.
3. Pool variability rates the overall
mixture of pool types found in the
stream according to size and depth.
The four basic types of pools are
large-shallow, large-deep, small-
shallow, and small-deep. A stream
with many pool types will support a
wide variety of aquatic species.
Rivers with low sinuosity (few
bends) and monotonous pool charac-
teristics do not have sufficient
quantities and types of habitats to
support a diverse aquatic community.
4. Channel alteration (See description
in habitat assessment for rocky-
bottom streams.)
5. Sediment deposition (See description
for rocky-bottom streams.)
6. Channel sinuosity evaluates the
sinuosity or meandering of the
stream. Streams that meander
provide a variety of habitats (such as
pools and runs) and stream velocities
and reduce the energy from current
surges during storm events. Straight
stream segments are characterized by
even stream depth and unvarying
velocity, and they are prone to
flooding. To evaluate this parameter,
imagine how much longer the stream
would be if it were straightened out.
7. Channel flow status (See description
in habitat assessment for rocky-
bottom streams.)
8. Bank vegetative protection (See
description for rocky-bottom
streams.)
9. Condition of banks (See description
for rocky-bottom streams.)
10. The riparian vegetative zone width
(See description for rocky-bottom
streams.)
Reference Collection
A reference collection is a sample of locally-found macroinvertebrates that have been
identified, labelled, and preserved in alcohol. The program advisor, along with a professional
biologist/entomologist, should assemble the reference collection, properly identify all samples,
preserve them in vials, and label them. This collection may then be used as a training tool and, in
the field, as an aid in macroinvertebrate identification.
-------
MACROINVERTEBRATES AND HABITAT I 101
Step 3—Leave the field, complete
data forms, clean the site, and
return material
After completing the stream character-
ization and habitat assessment, make sure
that all of the field data sheets have been
completed properly and that the informa-
tion is legible. Be sure to include the site's
identifying name and the sampling date on
each sheet. These will function as a quality
control element. If you can't determine how
to answer a question on the field data sheet,
just leave the space blank.
Before you leave the stream location,
make sure that all your equipment has been
collected and rinsed properly. Double-
check to see that sample jars are tightly
closed and properly identified. All samples,
field sheets, and equipment should be
returned to. the coordinator at this point.
You might want to keep a copy of the field
data sheet for comparison with future
monitoring trips and for personal records.
Step 4—Prepare for macro-
invertebrate laboratory work
This step includes all the work needed
to set up a laboratory for processing
samples into subsamples and identifying
macroinvertebrates to the family level. A
professional biologist/entomologist or the
program advisor should supervise the
'identification procedure. All interested
volunteers should be encouraged to partici-
pate. In general it is a good idea to train
volunteers in identification procedures
before each lab session and to start new
volunteers with less diverse samples.
Refresher workshops for experienced
volunteers are strongly encouraged.
TASK1
Gather tools arid equipment
for the laboratory
The following lab equipment is recom-
mended for the macroinvertebjrate identifi-
cation process. Enough of each will need to
be provided for each volunteer work
station:
• Reference collection and taxonomic
keys
• Fine-point forceps
• Petri dishes or small, shallow, clear
container
• Alcohol preservative (used in field
and lab): 70 percent ethyl alcohol,
denatured; no other preservatives
used
• Microscope, dissecting microscope,
and magnifying glass, or hands lens
• Sample containers, preferably
shatterproof with poly-seal caps that
prevent evaporation of the preserva-
tive (jars or vials are used in field
and lab). Shatterproof vials with
poly-seal caps are available from
scientific supply houses.
• Wash bottles or spray bpttles
• Shallow, rectangular white pans
(large enough to hold entire macro-
invertebrate sample)
• Additional shallow white containers
(heavy duty plastic plates with a rim,
white pans, or cafeteria trays are all
possible choices).
• Plastic spoons or unslotted spatulas
• Sieve, purchased from scientific
supply company (#30) 6r homemade
(with same mesh size afe sampling
net) ;
• Permanent ink markers:
• Ruler
• Macroinvertebrate assessment
worksheet
• Pencils
• Note paper for counting
-------
102 I MACROINVERTEBRATES AND HABITAT
TASK 2 I Create gridded subsampling
pans
Figure 4.20
A gridded
subsampling
pan
Volunteers
collect a
subsample of
organisms by
picking them
from randomly
selected grid
squares.
Using the ruler, measure the inside
width and length of the large rectangular
white pan. Draw a grid of evenly sized
squares on the inside of the pan, using
permanent ink. The grid should fill the
entire inside of the pan. Number each
square. One pan will be needed for each
work station. Volunteers will use these pans
for randomizing the sample and selecting a
subsample of organisms.
TASKS
Prepare the lab and the
individual work stations
Before volunteers enter the lab, the
program manager will need to prepare work
stations. Make sure that all microscopes are
functioning properly and that each station
has access to all other equipment. The
reference collection should be centrally
located as should any other visual training
displays. The lab itself should be well lit
and well ventilated. A copy of lab safety
instructions should be visible to all volun-
teers.
Step 5—Conduct macro-
invertebrate processing and
identification
If possible, before beginning the
subsampling and identification processes,
all volunteers should become familiar with
the lab equipment, microscope(s), the
reference collection, and the taxonomic key
chosen by the advisor.
Processing a subsample and identifying
the organisms are two separate activities.
Some programs might prefer to split these
tasks into separate lab sessions.
Session 1:
Picking a subsample of
aquatic organisms
TASK 1 | Prepare the sample
1. Carefully remove the station ID tag
from the sample container and put it
aside. You will need it later.
2. Cover the bottom of the gridded pan
with about 1/4 inch of clean water.
3. Pour the preserved sample (alcohol
and debris) into the sieve and wash
off preservative over a sink, using a
spray or wash bottle filled with
water.
4. Transfer the sample to the white
gridded pan by turning the sieve
upside down over the pan. Tap it
several times to empty the contents
onto the pan. Squirt a small amount
of water over the bottom of the sieve
to flush the organisms into the pan.
5. With your hands and by gently
shaking the pan, evenly disperse the
sample over the entire bottom of the
pan, making sure that even the
corners are covered. The water will
help in distributing the sample
throughout the pan. This is called
randomizing the sample.
-------
MACROINVERTEBRATES AND HABITAT I 103
TASK 2 I Randomly select a square for
""""""""""""^ the subsample
1. Randomly choose a square to start
sorting organisms. You may use a
random numbers table, draw num-
bers from a hat, or roll a pair of dice.
The most important thing to remem-
ber is that the grid selection should
be random. Indicate the square
number selected on the lab sheet.
2. Using a plastic spoon or unslotted
spatula, remove all the material from
the square and transfer it to another
container (another pan, tray, or plate)
for sorting. The organisms in this
container will become your
subsample.
TASK 3 I Pick the subsample
1. Prepare a container to house the
subsample by filling a vial or jar one-
half full of alcohol. Place the new
label into the vial, writing side out.
Keep the vial on a flat, stable area.
2. Using forceps, carefully and system-
atically remove all organisms from
the pan or tray and place them one by
one into the prepared subsample vial.
Examine all debris such as leaves or
sticks for clinging organisms. Count
each organism as it is transferred.
Keep a written count of the number
of organisms you have transferred.
The objective is have at least 100
individual organisms in your
subsample. If you reach 100 and
there are still organisms remaining in
your subsample plate or tray, con-
tinue picking until all the organisms
are removed even though you might
end up with more than 100.
When you think all the organisms
have been transferred from the plate
or tray to the subsample vial, have a
second volunteer check to confirm
that all organisms have been re-
moved. On your lab sheet, record
how many organisms are in the
subsample.
3. If you finish picking the contents of
the first square selected :and have
fewer than 100 organisms, randomly
select another square and repeat the
process of removing the contents of
the square to the subsample plate or
tray; picking organisms with the
forceps and transferring them to the
vial (all organisms that will be part
of the subsample should be trans-
ferred to the same vial). Record the
number of organisms you obtain
from the second square. Repeat this
process until at least 100 organisms
have been placed into the vial or
until the entire sample in the gridded
pan has been picked clean. Remem-
ber, any square started must be
picked clean.
If, after picking the entire gridded
pan clean, you have fewer than 100
organisms, and your reference site
SUBSAMPLE ID TAG
Station #: ___
Stream: '
Location: •
Date/Time: '
Subsample team members:
Figure 4.21
Example of a
Subsample ID
tag
To prevent
subsamples
from being
mixed up,
volunteers
should place the
ID tag inside the
subsample jar.
-------
104 I MACROINVERTEBRATES AND HABITAT
produced 100 or more organisms,
either your site is impaired or your
sampling technique is flawed. It is
also possible that recent heavy rains
might have washed many organisms
downstream. If you do not find 100
organisms in the entire sample, be
sure to note the potential cause for
such a problem on the Habitat
Assessment Data Sheet.
TASK 4
Label and store the
subsample
Fill out a new Subsample ID Tag (Fig.
4.21) for the subsample. Remember to use
pencil because ink will run in the alcohol.
The vial housing the subsample must be
labeled with the same station number,
stream name, location, and date found on
the original sample ID tag. The vial tag
should also include information on when
the subsample was picked (i.e., 100 or more
organisms counted) and by whom. Place
the tag in the vial with the writing side out.
Make sure the vial is tightly closed before
giving the subsample in the vial to the
program coordinator.
TASK 5 I Replace remainder of
original sample back into the
sample jar
Place the remaining sample back into
the original container. Be sure that the
original station ID tag is included, writing
side out. Fill the jar with 70 percent alco-
hol. This sample will be retained as part of
a voucher collection. Make sure the jar is
tightly closed before returning it to the
program coordinator.
Session 2: Identifying the subsample to
family level , ,
TASK 1 | Prepare for the ID
1. Make sure that you have several petri
dishes, fresh alcohol, and fresh water
close at hand. Also have your
taxonomic keys handy for all stages
of the ID process. Check to make
sure that your microscope is working
properly.
2. Carefully remove the station ID tag
from the subsample vial and put it
aside. You will need it later. Be sure
no organisms are clinging to it. If
they are, remove them with forceps.
3. Using the information on the station
ID tag, complete the first section of
the Macroinvertebrate Assessment
Sheet with your name, date, the
stream name, station number, and
any other information requested.
TASK 2
Identify the sample to order
level
1. Place a few of the macroinverte-
brates in a petri dish (or other small,
shallow container) and examine them
under the microscope. Include some
ethyl alcohol in the dish to ensure
that the organisms do not dry out.
Compare the organisms in the dish to
those in the taxonomic key and/or
reference collection.
2. Roughly sort organisms by taxo-
nomic order into petri dishes. Many
volunteers find it helpful to use one
dish for every major taxonomic order
found in the subsample. Place any
organism that you cannot identify
into another dish for the biological
advisor to examine.
-------
MAfcROINVERTEBRATES AND HABITAT I 105
TASK 3
1
Identify the organisms within
each order to family level
Starting with one order, and using
the taxonomic keys, reference
collection, and assistance of the
biological advisor, identify each
individual to family level.
2. Keep a running count of how many
individuals there are in each family
on a piece of scratch paper.
3. Place any organisms that you cannot
identify into a separate container.
Make sure that the biological advisor
sees them and assists you with the
ID.
4. After all organisms have been
identified, note the total number of
organisms in each family on the
Macroinvertebrate Assessment
Sheet. Write in pencil and make sure
your writing is legible. These lab
sheets will be the basis for the data
analysis. It is important that they are
accurate and easy to read.
TASK 4 I Return the organisms to the
——• vial
1. After you have identified and
counted all organisms in the
subsample, return them to the
subsample vial and replace the
subsample ID Tag, writing side out.
2. Refill the subsample vial with 70
percent ethyl alcohol (new or re-
cycled). Be sure to secure the caps on
the vial tightly to prevent the organ-
isms from drying out.
3. Return the subsample vial and the
assessment worksheet to the program
manager.
Voucher Collection
Maintaining a voucher collection adds another layer of
credibility to the program by documenting the accuracy of the
volunteer identifications. It substantiates and provides evidence to
support the analysis of the data—a powerful quality control
element. However, an important issue to consider is how long to
keep the samples. Program managers, in collaboration with
technical advisors, will have to consider the following in keeping a
voucher collection.
• Sample maintenance. Even jars and vials with tight fitting
lids require maintenance on a regular basis (every 2-3
months) to ensure that alcohol levels are adequate.
• Fire safety. When you are dealing with alcohol, you will
need to consider fire safety and ventilation issues to make
sure that you are in line with local codes.
• Availability of storage space. In addition to needing well-
ventilated and fire-proof storage cabinet, you will need a
well-ventilated room to store samples. Samples should not
be stored in someone's office for any length of time.
• Length of storage. How long samples should be maintained
is an issue determined by program goals. Data collected for
regulatory purposes will probably require longer storage
than other samples. Generally, 1-5 years is recommended
for storage.
Step 6—Performing habitat
assessment data analysis
To evaluate the condition of your
stream site properly, you should compare it
to an optimal or best condition found in the
region. This is called a reference condition.
In an ideal world, the reference condition
would reflect the water quality', habitat, and
aquatic life characteristics of pristine sites
in the same ecological region as your
stream. In real life, however, few pristine
sites remain. The reference condition is,
therefore, a composite of sites that reflect
the best physical, chemical, and biological
conditions existing in your ecological
region. State water quality or natural
resource agencies might have already
established reference conditions for the
ecological regions in your state.
-------
106 I MACROINVERTEBRATES AND HABITAT
Table 4.5
Reference
scores for
sampling site
comparison
If a score falls at
or near the
break between
categories, use
your best
judgement to
determine the
appropriate
score.
% Similarity to
Reference Score
> 90 %
75-88%
60-73%
< 58%
Habitat Quality
Category
Excellent
Good
Fair
Poor
Attributes
Comparable to the best situation to be expected
within an ecoregion. Excellent overall habitat
structure conducive to supporting healthy
biological community.
Habitat structure slightly impaired. Generally,
diverse instream habitat well-developed; some
degradation of riparian zone and banks; a small
amount of channel alteration may be present.
Loss of habitat compared to reference. Habitat is a
major limiting factor to supporting a healthy
biological community.
Severe habitat alteration at all levels.
Your program's consulting biologist
should work in cooperation with the state
agency to identify the reference
condition(s) you will need to conduct an
Intensive Stream Biosurvey. The biologist
will use the reference condition to establish
a water quality rating system against which
to rank your monitored stream sites.
To perform the habitat assessment data
analysis for the Intensive Stream
Biosurvey, perform the following tasks.
TASK1
Determine the habitat index
score
Add together the scores of all 10 habitat
parameters. This sum is the habitat index
score for the study stretch.
TASK 2
Determine the percent
similarity to the reference
score
Divide the habitat index score by the
reference index score and then multiply the
result by 100. This number is the percent
similarity to the reference score.
TASK 3 I Determine the stream habitat
quality rating
Compare the percent similarity of your
results with the range of percent similarity
numbers in the stream habitat rating table to
obtain the habitat quality category for your
site(s) (Table 4.5). Enter the appropriate
descriptive rating (excellent, good, fair, or
poor) on the field data sheet. If your score
falls at or near the break between habitat
quality categories, use your best judgment
to determine an appropriate rating.
Step 7—Conduct
macroinvertebrate data analysis
In general, the program's biological
advisor, rather than the volunteers, should
analyze the results of the Intensive Stream
Biosurvey's macroinvertebrate identifica-
tion. The advisor's knowledge of local
ecological conditions will help in the
interpretation of the data findings and will
lend additional credibility to the sampling
effort. Volunteers can contribute signifi-
cantly to the advisor's data analysis by
interpreting field notes, assisting with
-------
MACROINVERTEBRATES AND HABJTAT I 107
macroinvertebrate identification, and
counting organisms on the aquatic macroin-
vertebrate assessment worksheet. Relay the
results of the data analysis to the volunteers
as soon after the sampling date as possible.
TASK 1 j Determine which metrics or
"^^"^^""^ measurements are appropri-
ate
A number of metrics (or measures) can
be used to calculate stream health using
benthic macroinvertebrates. These metrics
should be calculated for both the sample
site and the reference condition. By com-
paring the two, the program advisor can
reach a clear understanding of the biologi-
cal health of the sampling site.
The Intensive Stream Biosurvey
recommends the use of four basic metrics
(taxa richness, number of EPT taxa, percent
abundance of EPT, and sensitive taxa
index) plus two optional metrics (percent
abundance of scrapers and percent abun-
dance of shredders). These metrics are
discussed briefly below. Refer to the
reference list for more information.
The term taxa (plural for taxon), used
below, refers to the specific taxonomic
groupings to which organisms have been
identified. For the Intensive Stream
Biosurvey, organisms are identified to the
taxon of family. Your volunteer monitoring
program should identify organisms to a
specific taxonomic grouping if it is to
compare results over time and between
sites. The following metrics are generally
applicable throughout the country (but
confirm this with a local biologist).
1. Number of taxa (taxa richness)—this
measure is a count of the number of
taxa (e.g., families) found in the
sample. A high diversity or variety is
good.
2. Number of EPT taxa (EPT rich-
ness)—this measure is a count of the
number of taxa in each of three
generally pollution-sensitive orders:
Ephemeroptera (mayflies),
Plecoptera (stoneflies), and
Trichoptera (caddisflies). A high
diversity or variety is good.
3. Percent dominance—this measure is
the percent composition of the most
abundant family from your station. It
indicates how dominant a single
taxon is at a particular site. A high
percent dominance is not good.
4. Sensitive taxa index (modified
Hilsenhojf index)—this measure is
calculated by multiplying the
number of organisms in each taxon
by the pollution tolerance value
assigned to each taxon, adding these
for all taxa represented in the
sample, and dividing by the total
number of taxa in the sample. A high
index number is not good.
Sensitive taxa index =
where:
n =
the summation of X,t
the number of individuals
in each taxon
tolerance value for each
taxon in the sample
number of individuals in
the sample
The following optional metrics can be
used in rocky-bottom streams frf at least 10
scraper and shredder organisms are col-
lected.
5. Percent abundance of scrapers—in
the majority of rocky-bottom
streams, the basic food source for
many aquatic organisms is algae
covering the rocks in the stream.
Macroinvertebrates that "scrape"
or graze on these algae are known as
scrapers. To compute the percent
-------
108 I MACROINVERTEBRATES AND HABITAT
Selecting Metrics to Determine Stream Health
Metrics are used to analyze and interpret biological data by condensing lists of organisms into relevant biological information. In order to be !
useful, metrics must be proven to respond in predictable ways to various types and intensities of stream impacts. This manual recommends
using a multimetric approach that combines several metrics into a total Biosurvey Score. The four primary and two optional metrics discussed in
this chapter have been tested extensively in the mid-Atlantic region and have been shown to respond in predictable ways to stream impacts. In
olher parts of the country, other metrics and scoring systems may be more appropriate. For example, the Benthic Index of Biotic Integrity (B-IBI),
developed by Dr. James Karr, is another multimetric approach, using different metrics, that has been tested in the Tennessee Valley, the
Midwest, and the northwest. The River Watch Network suggests that, while you should always use multiple metrics to summarize your data, you
shouldn't rely solely on an overall score to interpret your data; individual metrics can also provide a wealth of information. In any case you will
need to select metrics that have been proven to respond predictably to various impacts. As always, consult with your program's biological
advisor for help in selecting appropriate metrics for your region and for determining whether an overall biosurvey score is recommended.
Below are metrics that are commonly used in rocky bottom streams. This is only a partial list of the dozens of metrics used by monitoring
programs throughout the country. These metrics fall under four general categories: 1) taxa richness and composition, 2) pollution tolerance and
Intolerance, 3) feeding ecology, and 4) population attributes. Metrics marked with a (*) are included in the recommended suite of metrics in this
manual. The River Watch Network's Benthic Macroinvertebrate Monitoring Manual contains detailed guidance on selecting, calculating,
aggregating, and interpreting the metrics discussed below. (See Dates, G. and J. Byrne in References and Further Reading)
Taxa Richness and Composition Metrics
• Total Number of Taxa *: the total number of taxa found in the sample.
• Number of EPT Taxa *: the combined number of mayf ly (E), stonefly (P) and caddisfly (T) taxa found in the sample. The number of taxa
in each of these macroinvertebrate orders can also be reported separately since each order may respond differently to various impacts.
• Number of Long-Lived Taxa: the number of organism families found in the sample (such as giant stoneflies and dobson flies) that live
more than one season.
• Percent Abundance of the Major Groups *: the percent of the sample that is comprised of individuals in each of the selected major groups
(mostly orders).
• Percent Model Affinity (Bode, 1991): used in conjunction with Percent Composition of the Major Groups, this metric measures the
similarity of the sample to a model "nonimpacted" community of organisms (adjusted for ecoregional conditions) based on the percent
composition of the major groups.
• Quantitative Similarity Index (from Shackleford, 1988): used in conjunction with Percent Composition of the Major Groups, this metric
shows the percent similarity between two sites based on the percent of the sample in each of the major groups.
• Dominants in Common (from Shackleford, 1988): the number of dominant (5 most abundant families) families common to two sites.
Tolerance and intolerance Metrics
• Number of Intolerant Taxa: the number of taxa in the sample that are in the 10-15% of the least tolerant taxa in a region or that have a
pollution tolerance value of 1 (based on the Hilsenhoff scale of 0-10).
• Percent of Individuals in Tolerant 7axa;the number of taxa in the sample that are in the 10-15% of the most tolerant taxa in a region or
that have a pollution tolerance value of 10 (based on the Hilsenhoff scale of 0-10).
• Number of Clinger Taxa: the number of families in the sample that live by clinging to the bottom of the stream.
• Sensitive Taxa Index*: the pollution tolerance values (based on the Hilsenhoff scale of 0-10) assigned to each family aggregated into an
overall pollution tolerance value for the sample.
Feeding Ecology Metrics
• Percent Composition of Functional Feeding Groups: the percentage of the total number of individuals in the sample that belong to each of
the five functional feeding groups (scrapers, shredders, filtering collectors, gathering collectors, and predators).
• Percent Abundance of Scrapers *: the percent of the total number of individuals in the sample that use bottom-growing algae as their
primary food source.
• Percent Abundance of Shredders *: the percent of the total number of individuals in the sample that use leaves and other plant debris as
their primary food source.
• Percent Abundance of Predators: the percent of the total number of individuals in the sample that eat other animals as their primary food
source.
Population Attributes Metrics
• Percent Dominance (of the most abundant family) *: the percentage of the total number of individuals in the sample that are in the
sample's most abundant family.
• Percent Dominance (of the three most abundant families): the percentage of the total number of individuals in the sample that are in the
sample's three most abundant families.
• Organism Density Per Sample (total abundance): the total number of individuals in the sample (calculated if a subsample is used).
-------
MACROINVERTEBRATES AND HABITAT I 109
abundance of the scrapers in the
macro-invertebrate community,
divide the number of organisms
classified as grazers or scrapers by
the total number of organisms in the
sample. A high percent abundance of
scrapers is good.
6. Percent abundance of shredders—
leaf litter and other plant debris are
broken down and processed by
organisms called shredders. To
compute the percent abundance of
shredders in the macroinvertebrate
community, divide the number of
organisms classified as shredders by
the total number of organisms in the
sample. A high percent abundance of
shredders is good.
The following optional metrics can be
used in muddy-bottom streams as addi-
tional metrics to provide more information
about the condition of the macroinverte-
brate assemblage.
7. Percent abundance ofEPT—this
measure compares the number of
organisms in the EPT orders to the
total number of organisms in the
sample. (The number of organisms in
the EPT orders is divided by the total
number of organisms in the sample
to calculate a percent abundance.) A
high percent abundance of EPT
orders is good.
8. Percent abundance of midge lar-
vae—this measure compares the
number of midges to the total
number of organisms in the sample.
(The number of organisms in the
chironomidae family is divided by
the total number of organisms in the
sample to calculate a percent compo-
sition.) A low percent abundance of
midge larvae is good.
TASK 2
Calculate a score for the site
The metric worksheets Tables 4.6 and
4.7 are designed to help calculate a total
score for the monitored site. Table 4.8
provides an example of a sample metric
worksheet for the fictional Volunteer Creek
(rocky-bottom stream). This score should
be compared to reference conditions to
determine the biological condition of the
stream at that site. You should also note
that these worksheets were developed for
use in mid-Atlantic states; they might need
to be modified to reflect local conditions.
To calculate a score for your stream
site using one of these worksheets, enter
the metric values at the monitored site in
the (M) column. Compare each metric
value from your monitored siteto the value
ranges presented in the biosurvey score
columns. Choose the matching range and
circle it; this gives you the corresponding
score (6, 3, or 0) for your metric value.
Add the metric scores to obtain the total
biosurvey score (see instructions in Tables
4.6 and 4.7).
TASKS
Determine the biological
condition
To determine the biological condition
of the site, refer to Table 4.9, Biosurvey
Scoring Guide.
TASK 4
Return the lab sheets and
metric worksheets to the
program coordinator
All remaining worksheets should be
returned to the program coordinator once
the site's final score has been determined.
The program coordinator will determine
how to proceed with the findings of the
biological assessment (e.g., the data may be
entered into a database or shared with a
state or local agency). It is important that
the biological advisor include documenta-
tion of any problems encountered in the
process of monitoring, identifying macroin-
vertebrates, or analyzing the data.
-------
110 I MACROINVERTEBRATES AND HABITAT
Table 4.6
Metric
worksheet for
rocky-bottom
streams
Primary Metrics
No. of Taxa
No.ofEPTTaxa
% Dominance
Sensitive Taxa Index
Optional Metrics
% Abundance of Scrapers
% Abundance of Shredders
(M)
Monitored
Site Values
Biosurvey Score
(Circle the appropriate range for each metric)
COLUMN SCORE (multiply no. of circled values
by the biosurvey score)
TOTAL SCORE (Sum all the column scores)
>8
< 34%
<4.8
15-8
8-4
34 - 67%
4.8 - 6.4
18-10%
9 - 5%
<8
<4
> 67%
>6.4
Notes: If fewer than 60 individuals in the monitored site, don't calculate metrics for any of the sites.
Biosurvey scoring ranges determined for the summer index period.
Table 4.7
Metric
worksheet for
muddy-bottom
streams
Primary Metrics
No. of Taxa
No.ofEPTTaxa
% Dominance
Sensitive Taxa Index
Optional Metrics
% Abundance of EPT
% Abundance of Midge Larvae
(M)
Monitored
Site Values
Biosurvey Score
(Circle the appropriate range for each metric)
COLUMN SCORE (multiply no. of circled values
by the biosurvey score)
TOTAL SCORE (Sum all the column scores)
>7
< 30%
<5.0
> 39%
< 24%
7-4
30 - 50%
5.0 - 6.8
39 - 20%
24 - 60%
<4
> 50%
>6.8
< 20%
> 60%
Notes: If fewer than 60 individuals in the monitored site, don't calculate metrics for any of the sites.
Biosurvey scoring ranges determined for the summer index period.
-------
MACROINVERTEBRATES AND HABITAT I 111
Primary Metrics
No. of Taxa
No.ofEPTTaxa
% Dominance (81 individuals)
Sensitive Taxa Index
(M)
Monitored
Site Values
Biosurvey Score
(Circle the appropriate range foteach metric)
12
67%
3.83
COLUMN SCORE (multiply no. of circled values
by the biosurvey score)
TOTAL SCORE (Sum all the column scores)
>8
<34%
9
<4
> 67%
>6.4
Biosurvey Score for this site is 15
This site scores in the Fair range, 9-15
Table 4.8
Sample metric
worksheet for
Volunteer
Creek
(hypothetical
rocky-bottom
stream).
There were 119
macroinverte-
brates in this
sample.
Total Score
From Metrics
>18-24
9-15
0-6
Condition
Category
Good
Table 4.9
Fair
Poor
Attributes
Comparable to the best situation to be expected
within an ecoregion. Balanced trophic structure.
Optimum community structure (composition and
dominance) for stream size and habitat quality.
Community structure less than expected. Compo-
sition (species richness) and diversity lower than
expected due to loss of some pollution- intolerant
forms. Percent contribution of tolerant forms
increased. Reduction in EPT index.
Few species present. If high densities of organ-
isms, then dominated by one or two pollution-
tolerant taxa.
Biosurvey
Scoring Guide
This guide is
based on the
four primary
metrics. If your
score falls on
the boundary of
two categories,
consider the
site's habitat
assessment
results and
chemical data, if
available, in
confirming your
assignment to a
particular
category.
-------
112 I MACROINVERTEBRATES AND HABITAT
INTENSIVE BIOSURVEY:
MACROINVERTEBRATE ASSESSMENT
Stream Name:
County:
Investigators:
Site (description):
Latitude:.
Site or Map Number:
Date:
State:
Longitude:
Time:
Weather in past 24 hours:
Q Storm (heavy rain)
D Rain (steady rain)
Q Showers (intermittent)
Q Overcast
Q Clear/Sunny
Weather now:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent)
Q Overcast
Q Clear/Sunny
Type of Sampling (check one)
Rocky bottom Muddy bottom
Muddy Bottom Sampling Only: Record the number of
jabs taken in each habitat type.
Vegetated bank margin
Snags and logs
Aquatic vegetation beds
Silt/sand/gravel substrate
-------
MACROINVERTEBRATES AND HABITAT I 113
Q. 0.
CD O I Q_
O CO CO U-
Q. Q.
LO
CM CO LO IO IO
D. O. Q. Q.
T- CO IO
I CD CD
CO O O
O
LJ-
to
CO
<
o
DC
O
Megaloptera
Corydalidae (Dobsonflies/Fishflies)
Slalidae (Alderflies)
Water Beetles (Coleoptera)
Elmidae
Psephenidae
Ptilodactylidae
Gyrinidae
Haliplidae
Hydrophilidae
Dytiscidae
Dragonflies & Damselflies (Odonata)
Gomphidae (Clubtails)
Aeshnidae (Darners)
Calopterygidae (BW-damselflies)
Coenagrionidae (NW-damselflies)
Waterbugs (Hemiptera)
Crustacea:
Sowbugs (Isopoda) Aseliidae
Scuds (Amphipoda) Gammaridae
Crayfish (Decapoda) Astacidae
P. ANNELIDA
Earthworms, C. Oligochaeta
Leeches, C. Hirudinea
P. MOLLUSCA, C. Gastropoda (Snails]
P. MOLLUSCA, C. Bivalvia
(Clams & Mussels)
Other Animals (Invertebrates Only)
CD
.C
O
o a?
I i
si
§1
CO CO
CO CO
if
c2
1°
£ c
O)
_ CD
^§
?l
UJ E
b!
Z 0_
QC
UJ
t =
£
CD
J=
to
CD
^i
o
o *-
0) CD
8 I
11 'IF
CD _
O a.
o
t3
jCD
"o
O
.
Bi i
EC W U-
C5 II II
O O O
CO U-
< |*
•^ .t;
Ui co
O g
5 w
oc
UI
to
a
ui
ui >-
U. 0
-J -a
< £
icS M
p n a-
O I II
Z CO OL
LL
^
CD D)
d> D)
O c
i «
s •-
» o"
.0 V
O
CO
D- a.
CO O O
I 10 CD
CO CO U- O
T- CD CM
OOCD
cocoo
CD, CD
oo
CD CD
Oo
cococococococo oii-co
t-CMOT-;OCNCMT-O
8
co
.CD
ir
CD
CD J£ S D).2 2 .2
I
"^
CD
•D
ichop
. C CD
(n s?iroir
l|
fa|gs«stl|l
0 . CO « o-g S g §^V
-<-& = 2 ;§ o 8
0 •" i- CO T3
0 S '>»
2 II
ill
CO CO •»--
I- I- CO
® a*
-------
114 I MACROINVERTEBRATES AND HABITAT
INTENSIVE BIOSURVEY:
HABITAT ASSESSMENT
Stream Name:
County:.
Investigators:
Site (description):
Latitude:.
Site or Map Number:.
Date:
State:
Longitude:
Time:
Weather in past 24 hours:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
Weather now:
Q Storm (heavy rain)
Q Rain (steady rain)
Q Showers (intermittent rain)
Q Overcast
Q Clear/Sunny
-------
MACROINVERTEBRATES AND HABITAT I 115
Sketch of site
On your sketch, note features that affect stream habitat, such as: riffles, runs, pools, ditches, wetlands, dams, riprap,
outfalls, tributaries, landscape features, logging paths, vegetation, and roads.
-------
116 I MACROINVERTEBRATES AND HABITAT
i:
I
a.
c
Q>
I
03
Q>
'as
ter
m.
1
t^
g
•D
"5
0)
*V"
D)
c
1
^
'E
.2
d>
o>
D)
'I
Q
.C
.§•
ar
»i
=!
^1
CO
1
U.
1
1
1
f 5
uatuiuoc
XI
5
-3
-8
a «>
w"
(0
o
QC
m
1
0.
CO
CD
•o
3
C
_O
o
Ta
o
O
CD
Q.
.O
CD
1
DJ
C
'«
=3
o
I
k__
CD
Q.
O
2
CD
73
.I
a
E
O
O
o
onstructi
o
I
J5
•D
CD
O
CC
holding areas
'§
_ o
(0 "P J2
3 J2 .2
*±
o .— ^ —
~ y U Q.
±=. CO fl> O
O) i- CD £
< G LL O
ields
nactive agricul
ion
boating
O)
C '
'CD
o
!
ti
n>.s g.
Hill
I 11 i 11
_ O O CO
CC 0. 0 O
S
Hi
01
•5.
§
ca
-
0
Il
0 2
ill
o £ J3
TO
1
<5 E
CM
a a a CM a a
s= v-aaaa r-aa
^11^
cMaaa cMaaaa cMaaaaa cMaaaaaa
r-aaa ^-aaaa i-aaaao r-aaaaaa
in
S
8,
^CjjjJ^B ,c*
m -s-
•*•• ooo
CM CM CO
O) O) O)
CD CD 0)
O) D) O)
TO CO CO
Q- D. Q.
1
CO
2 CD m
o ^ ^ a
zo
M oca aa -g
H i
H ic i I i
2
CO
8.
a.
ra
o
I
a aa
s^-i
oil
aaa
w o
iT cc
aa
o
o
£
03
a a
S
1
0)
a.
o
ra
£
w
•»•
o
S
CO
2
o.
a.
a
CO
-------
MACROINVERTEBRATES AND HABITAT I 117
HABITAT ASSESSMENT FIELD DATA SHEET
ROCKY BOTTOM SAMPLING
Habitat
Parameter
1 . Attachment
Sites for Macro-
invertebrates
Page 93 I
srnRF
2. Embeddedness
Page 93 |
SCORE
3. Shelter for Fish
Page 93 |
SCORE
4. Channel
Alteration
Page 93 I
srnpp
5. Sediment
Deposition
Page 94 |
SCORE
Category
Optimal
Well-developed riffle
and run; riffle is as
wide as stream and
length extends 2 times
the width of stream;
cobble predominate;
boulders and gravel
common.
^20^9-1:8 ;:v:f 7; :^6:.
Fine sediment ,
surrounds and fills in
0-25% of the living
spaces around and in
between the gravel,
cobble, and boulders.
20 19 18 17 16
Snags, submerged
logs, undercut banks,
cobble and large
rocks, or other stable
habitat are found in
over 50% of the site.
.20 19 18 17 16
Stream straightening,
dredging, artificial
embankments, dams
or bridge abutments
absent or minimal;
stream with
.meandering pattern.
2O i%9 .T8>\17 Vlff
Little or no
enlargement of
islands or point bars
and less than 5% of
the bottom affected
by sediment
deposition.
20 19 ;|8 17 ;i€
Suboptimal
Riffle is as wide as
stream but length is less
than 2 times width;
cobble less abundant;
boulders and gravel
common.
; • r5;v;i.m:j3 si:2'; n 1: '.
Fine sediment
surrounds and fills in
25-50% of the living
spaces around and in
between the gravel,
cobble, and boulders.
15 14 13 12 11
Snags, submerged
logs, undercut banks,
cobble and large rocks,
or other stable habitat
are found in over 30-
50% of the site.
15 14 13 12 11
Some stream
straightening,
dredging, artificial
embankments or dams
present, usually in
areas of bridge
abutments; no
evidence of recent
channel alteration
activity.
:0$m4-^&::^2?;3^z
Some new increase in
bar formation, mostly
from coarse gravel;
5-30% of the bottom
affected; slight
deposition in pools.
15. :-;i:4 •:i:3;::;i-2,,;'i;tij?.
Marginal
Run area may be
lacking; riffle not as
wide as stream and its
length is less than 2
times the stream width;
gravel or large boulders
and bedrock prevalent;
some cobble present.
v:*/iv;.- -Q Ova'..'v:-r „.;»*..•...
••- |:y :•::'. .H 13 1 .. sD •••
Fine sediment
surrounds and fills in
50-75% of the living
spaces around and in
between the gravel,
cobble, and boulders.
i
10 9 8 7 6 :
Snags, submerged
logs, undercut banks,
cobble and large rocks,
or other stable habitat
are found in over 1 0-
30% of the site.
10 9 8 7 6
Artificial embankments
present to some extent
on both banks; and 40
to 80% of stream site
straightened, dredged,
or otherwise altered.
;aO •9:^.8;:-.:.' ;7 y--6", ,
Moderate deposition of
new gravel, coarse
sand on old and new
bars; 30-50% of the
bottom affected;
sediment deposits at
stream obstructions
and bends; moderate
deposition in pools.
;;;TO/9..;: /,8; . T -.••••&$•••
Poor
Riffles or run virtually
nonexistent; large
boulders and bedrock
prevalent; cobble
lacking.
5 43 21 0
Fine sediment
surrounds and fills in
more than 75% of the
living spaces around
and in between the
gravel, cobble, and
boulders.
5 : 4 ,3 :2,: 1 0
Snags, submerged
logs, undercut banks,
cobble and large
rocks, or other stable
habitat are found in
less than 10% of the
site.
5 4, .3 ,2:;;KJ1 0 :
Banks shored with
gabion or cement;
over 80% of the
stream site
straightened .and
disrupted.
5 4 3. -Z;. .1 O
Heavy deposits of fine
material, increased bar
development; more
than 50% of the
bottom affected;
pools almost absent
due to substantial
sediment deposition.
5 4: 3 .;2 1 0
-------
118 I MACROINVERTEBRATES AND HABITAT
ROCKY BOTTOM SAMPLING
Habitat
Parameter
6, Stream Velocity
and Depth
Combinations
Page 94
SCORE
7. Channel Flow
Status
Paga 94
SCORE
8. Bank
Vegetative
Protection (score
each bank)
Pago 95
Note: determine
left or right side
by facing
downstream
SOORP (1 RJ
SCORE (RB)
9. Condition of
Banks (score each
bank)
I I i
Paga 95
SC0RF , (IP)
SCORE (RB)
10. Riparian
Vegetative Zone
Width (score each
bank riparian
zone)
Paga 95
SCORE (LB)
SCORE (RB)
Category
Optimal
Slow «1 ft/s)/deep
(>1.5ft);
slow/shallow;
fast/deep;
fast/shallow
combinations all
present.
20 '19,> 18;::-i1;?-fi-:$
Water reaches base
of both lower banks
and minimal amount
of channel substrate
is exposed.
20 .19 18. 17:;:16.
More than 90% of
the streambank
surfaces covered by
natural vegetation,
including trees,
shrubs, or other
plants; vegetative
disruption, through
grazing or mowing,
minimal or not
evident; almost all
plants allowed to
grow naturally.
Left Bank .,10 9
Right Bank 10 ' 9
Banks stable; no
evidence of erosion
or bank failure; little
potential for future
problems.
Left Bank 10. ,9
Right Bank 1 6 . -9 .
Width of riparian zone
>50 feet; no
evidence of human
activities (i.e., parking
lots, roadbeds, clear-
cuts, mowed areas,
or crops) within the
riparian zone.
Left Bank 10 9
Right Bank '10 9
Suboptimal
3 of the 4
velocity/depth
combinations are
present; fast current
areas generally
dominate.
^m^i^^s-'-^,-^
-'<. ; :< .•;...:• •<". '• .•.;.:..•.•.•.„-.._• •.'- .•-..... -.-•- ; «.••.
Water fills >75% of
the available channel;
<25% of channel
substrate is exposed.
;'1S: X^X^j.*™*^
70-90% of the
streambank surfaces
covered by natural
vegetation, but one
class of plants is not
well-represented; some
vegetative disruption
evident; more than
one-half of the
potential plant stubble
height remaining.
•:•::&•"• :•;:••/•••:•• > 6 .•
.^8-V;.:yC,:U:VVe:,..
Moderately stable;
infrequent, small areas
of erosion mostly
healed over.
8 7, 6
8.7 6
Width of riparian zone
35-40 feet.
8 7 6
v8,;,. 7 " •' "'".'^ :•
Marginal
Only 2 of the 4
velocity/depth
combinations present.
Score lower if fast
current areas missing.
&10 '.,&..: .8-^.7 "i/6--'.
Water fills 25-75% of
the available channel
and/or riffle substrates
are mostly exposed.
ti$&$9 .:..:&^ ??-^:6- :."-.
50-70% of the
streambank surfaces
covered by vegetation;
patches of bare soil or
closely cropped
vegetation common;
less than one-half of
the potential plant
stubble height
remaining.
^•:.6:., .,4;-'.:;::;,:; .-3
'.. •.••6.':" ':'C:':' '. '3,.
Moderately unstable;
up to 60% of banks in
site have areas of
erosion; high erosion
potential during floods.
5 4 3
5 : 4. ';>;;., .$.'•*;
Width of riparian zone
20-35 feet.
.•,'.'•5 4, : •.;•••.•... ;3:":":
:.::,. •'& 4 ;'•"•;•/ ;.'^>:
Poor
Dominated by 1
velocity/depth
category (usually
slow/shallow areas).
:X';5'. ..:4; :.:,3 2 1 0 ;
Very little water in
channel and mostly
present as standing
pools.
:5'£.-4:. 3 2 1,"0
Less than 50% of the
streambank surfaces
covered by
vegetation; disruption
of streambank
vegetation is very
high; vegetation has
been removed to 2
inches or less in
average stubble
height.
••••&•.• ••':.. ':1 .- ..-0 • :i
2 i '. o ;
Unstable; many
eroded areas; "raw"
areas frequent along
straight sections and
bends; obvious bank
collapse or failure; 60-
100% of bank has
erosional scars.
2 1 0
'2/r.... :ri. ;,;•;?©• -fi -
Width of riparian zone
< 20 feet.
.2,, , , 1 .. o;^, :s.
.,2"- 1 0
Total Score
-------
MACROINVERTEBRATES AND HABITAT I 119
HABITAT ASSESSMENT FIELD DATA SHEET
MUDDY BOTTOM SAMPLING
Habitat
Parameter
1 . Shelter for
Fish and Macro-
invertebrates
Page 99
SCORE
2. Pool Substrate
Characterization
| Page 100
SCORE
3. Pool
Variability
| Page 100
SCORE
4. Channel
Alteration
Page 1 00
SOORF
5. Sediment
Deposition
| Page 1 00
SCORE
6. Channel
Sinuosity
Page 100
SCORE
Category
Optimal
Snags, submerged logs,
undercut banks, rubble
or other stable habitat
found over 50% of the
site; logs/snags are old
fall.
20 19 18 17 16
Pools have mixture of
substrate materials,
with gravel and firm
sand prevalent; root
mats and submerged
vegetation common.
20 19 18 17 16
Even mix of large-
shallow, large-deep,
small-shallow, small-
deep pools.
,;20: 19 18 17 16
Stream straightening,
dredging, artificial
embankments, dams or
bridge abutments
absent or minimal;
stream with
meandering pattern.
'&:'2Q^19*J?18.- .1.7."?T6.:;;-;
Less than 20% of
stream bottom
affected by extensive
sediment deposition;
minor accumulation of
fine and coarse
material at snags and
submerged vegetation;
little or no enlargement
of islands or point
bars.
;"2Q-n9<^i.8"A1:7:--;T6"::
The bends in the
stream would increase
the stream length 3 to
4 times longer than if
it was in a straight
line.
20 19 18 17 16
Suboptimal
Snags, submerged logs,
undercut banks, rubble
or other stable habitiat
found over 30-50% of
the site; some old fall,
but preponderance of
new fall.
15 14 13 12 11
Pools have mixture of
soft sand, mud, or clay
substrate; mud may be
dominant; some root
mats and submerged
vegetation present.
15 14 13 12 11 :
Majority of pools large-
deep; very few
shallow.
15 14 13 12 11
Some stream
straightening, artificial
embankments or dams
present, usually in
areas of bridge
abutments; no evidence
of recent channel
alteration activity.
:,;p'5:i;;i1,4x: :-T$--^Z:,J-T ':'l
20-50% of stream
bottom affected by
extensive sediment
deposition; moderate
accumulation;
substantial sediment
movement only during
major storm event;
some new increase in
bar formation.
-i;SCt4^3^^2 .11
The bends in the
stream would increase
the stream length 2 to
3 times longer than if it
was in a straight line.
15 14 13 12 11
' Marginal
Snags, submerged
logs, undercut banks,
rubble or other stable
habitiat found over 1 0-
30% of the site;
appears unstable;
some new fall.
10 9 8 7 6
Pools have all mud or
clay or sand
substrate; little or no
root mat; no
submerged
vegetation.
TO 9 8 7 6
Shallow pools much
more prevalent than
deep pools.
1O 9 8 7 6
Artificial
embankments present
to some extent on
both banks; and 40
to 80% of stream
site straightened,
dredged, or otherwise
altered.
J!l:0:?--9; n8<.'.; "C- 3'"V2. ::•! -0: •;?
Pools have hard-pan
clay or bedrock
substrate; no root mat
or vegetation.
5 43210 ;;
Majority of pools
small-shallow or pools
absent.
5p^.-4>-3.'.,2 ' .1;. .0.,-;;|
Banks shored with
gabion or cement; over
80% of the stream
site straightened and
disrupted.
::S ,;:4,;:3:p2' •?i::*.-:;cr :'
Greater than 80% of
stream bottom
affected by extensive
sediment deposition;
Heavy deposits; mud,
silt, and/or sand in
braided or nonbraided
channels; pools almost
absent due to
deposition.
5 43210
Channel straight;
waterway has been
channelized.
54321 Ov:
-------
120 I MACROINVERTEBRATES AND HABITAT
MUDDY BOTTOM SAMPLING
Habitat
Parameter
7. Channel Flow
Status
Pagaioo)
SRORP
8. Bank
Vegetative
Protection
| Page 100|
Note: determine
left or right side
by facing
downstream
SCORE (LB)
SCORE (RB)
9. Condition of
Banks
| Page 100J
SCORE (LB)
SCORE (RB)
10. Riparian
Vegetative1 Zone
Width {score
each bank
riparian zone]
(Page 100|
SCORE (I.BJ
SCORE {RB}
Category
Optimal
Water reaches base of
both lower banks and
minimal amount of
channel substrate is
exposed.
20 19 18 17 16
More than 90% of the
streambank surfaces
covered by native
vegetation, including
trees, understory
shrubs, or non-woody
macrophytes;
vegetative disruption
through grazing or
mowing, minimal or
not evident; almost all
plants allowed to grow
naturally.
Left Bank 10 9
Right Bank 1 0 9
Banks stable; no
evidence of erosion or
bank failure; little
potential for future
problems.
Left Bank 10 9
Right Bank . 109
Width of riparian zone
> 50 feet; human
activities (i.e. parking
lots, roadbeds, clear-
cuts, lawns, or crops)
have not affected
riparian zone.
Left Bank 10 9
Right Bank 10 9
Suboptimal
Water fills >75% of
the available channel;
<25% of channel
substrate is exposed.
15 14 13 12 11
70-90% of the
streambank surfaces
covered by native
vegetation, but one
class of plants is not
well-represented; some
vegetative disruption
evident; more than one-
half of the potential
plant stubble height
remaining.
87-6
8 7 6 '
Moderately stable;
infrequent, small areas
of erosion mostly
healed over.
876
876
Width of riparian zone
35-40 feet.
8.7 6
;8 7 6
Marginal
Water fills 25-75% of
the available channel
and/or riffle
substrates are mostly
exposed.
10 9 8 7 6
50-70% of the
streambank surfaces
covered by
vegetation; patches
of bare soil or closely
cropped vegetation
common; less than
one-half of the
potential plant
stubble height
remaining.
543
543
Moderately unstable;
up to 60% of banks
in site have areas of
erosion; high erosion
potential during
floods.
543
543
Width of riparian zone
20-35 feet.
543
543
Poor
Very little water in
channel and mostly
present as standing
pools.
543210
Less than 50% of the
streambank surfaces
covered by vegetation;
disruption of stream-
bank vegetation is
very high; vegetation
has been removed to 2
inches or less in
average stubble
height.
210
2 1 0
Unstable; many eroded
areas; "raw" areas
frequent along straight
sections and bends;
obvious bank collapse
or failure; 60-100% of
bank has erosional
scars.
210
2 1 0
Width of riparian zone
< 20 feet.
2 1 0
2 1 0
Total Score
-------
MACROINVERTEBRATES AND HABITAT I 121
z
UJ
o
o
^^^H s
^^^^H -a '5)
Comparable to the best situation to be expecte
within an ecoregion. Excellent overall habitat
structure conducive to supporting healthy biolo
community.
m Hii
•=1 K>>2fxl uj
K?3 m^BSM
BUB ^
v4 l^^^l
^^^^^^^^^^H P 'F Cli —
|| 3{ •s • •«
Q> 3: v» O O *- O
•° o (ft X O '(g O
^ -D ^ UJ O U- CL
CO C ^^^
**~
-------
122 I MACROINVERTEBRATES AND HABITAT
References and Further Reading
Note: References marked with (k)
contain macroinvertebrate taxonomic keys.
Brigham, A. R., W. U. Brigham, and A.
Gnilka. 1982. Aquatic Insects and
Oligochaetes of North and South Caro-
lina. Midwest Enterprises, Mahomet, DL.
(k)
Cummins, Kenneth W. and Margaret A.
Wilzbach. 1985. Field Procedures for
Analysis of Functional Feeding Groups
of Stream Macroinvertebrates. Univer-
sity of Maryland, Frostburg. (k)
Dates, G. and J. Byrne. 1995. River Watch
Network Benthic Macroinvertebrate
Monitoring Manual. River Watch
Network. 153 State St., Montpelier, VT
05602 ($25). (k)
Delaware Nature Education Center. 1996.
Delaware Stream Watch Guide. Dela-
ware Nature Society, P.O. Box 700,
Hockessin, DE 19707.
Fore, L., J. Karr, and R. Wiseman. 1996.
Assessing Invertebrate Responses to
Human Activities: Evaluating Alternative
Approaches. Journal of the North
American Benthological Society.
15(2):212-231.
Hilsenhoff, William L. 1982. Using a
Biotic Index to Evaluate Water Quality in
Streams. Wisconsin Department of
Natural Resources, Madison, WI. Tech-
nical Bulletin No. 132.
Hilsenhoff, William L. 1988. Rapid Field
Assessment of Organic Pollution With a
Family-level Biotic Index. Journal of the
North American Benthological Society,
7:65-68.
Izaak Walton League of America (IWLA).
1992. A Monitor's Guide to Aquatic
Macroinvertebrates. Izaak Walton
League of America Save Our Streams.
707 Conservation Lane, Gaithersburg,
MD 20878. (k)
Izaak Walton League of America (IWLA).
Stream Insects and Crustaceans Card.
Izaak Walton League of America Save
Our Streams. 707 Conservation Lane,
Gaithersburg, MD 20878. (k)
Karr, J. R. In press. Rivers As Sentinels:
Using the Biology of Rivers to Guide
Landscape Management. In The Ecology
and Management of Streams and Rivers
in the Pacific Northwest Coastal Ecore-
gion. Springer-Verlag, NY
Klemm, D.J., et al. 1990. Macroinverte-
brate Field and Laboratory Methods for
Evaluating the Biological Integrity of
Surface Waters. EPA/600/4-90/030. U.S.
Environmental Protection Agency,
Office of Research and Development,
Cincinnati, OH.
Lathrop, J. 1989. A Naturalist's Key to
Stream Macroinvertebrates for Citizen
Monitoring Programs in the Midwest. In
Proceedings of the 1989 Midwest Pollu-
tion Control Biologists Meeting, Chicago
IL, EPA 9059-89/007, ed. W.S. Davis
and T.P. Simon, USEPA Region 5
Instream Biocriteria and Ecological
Assessment Committee. Chicago,
Illinois, (k)
Maryland Save Our Streams. 1994. Project
Heartbeat Volunteer Monitoring Hand-
book. Maryland Save Our Streams, 258
Scotts Manor Dr., Glen Burnie, MD
21061.
McCafferty, W. P. 1981. Aquatic Entomol-
ogy: The Fishermen's and Ecologists'
Illustrated Guide to Insects and Their
Relatives. Science Books International,
Boston, (k)
McDonald, B., W. Borden, and J. Lathrop.
Citizen Stream Monitoring: A Manual
for Illinois. ILENR/RE-WR90/18.
Illinois Department of Energy and
Natural Resources.
Merritt, R. W. and K. W. Cummins, eds.
1984. An Introduction to the Aquatic
Insects of North America. 2d. ed.
Kendall/Hunt Publishing Company,
Dubuque. (k)
-------
MACROINVERTEBRATES AND HABITAT I 123
Moen, C. and J. Schoen. 1994. Habitat
Monitoring. The Volunteer Monitor
Needham, James C. and Paul R. Needham.
1988. A Guide to the Study of Fresh-
Water Biology. Reiter's Scientific and
Professional Books, Washington, D.C.
(k)
Peckarsky, Barbara L. et al., 1990. Fresh-
water Macroinvertebrates of Northeast-
ern North America. Cornell University
Press, Ithaca, New York, (k)
Pennak, Robert W. 1989. Fresh-Water
Invertebrates of the United States:
Protoza to Mollusca. 3rd. ed. John
Wiley and Sons, New York, (k)
Plafkin, J.L., M.T. Barbour, K.D. Porter.
S.K. Gross, and R.M. Hughes. 1989.
Rapid Bioassessment Protocols for Use
in Streams and Rivers: Benthic Macroin-
vertebrates and Fish. EPA 440/4-89-001.
U.S. Environmental Protection Agency,
Office of Wetlands, Oceans, and Water-
sheds, 4503F, Washington, DC 20460.
River Watch Network. 1992. A Simple
Picture Key: Major Groups of Benthic
Macroinvertebrates Commonly Found in
Freshwater New England Streams. River
Watch Network, 153 State St., Montpe-
lier, VT 05602 (k)
Tennessee Valley Authority (TVA). 1994.
Common Aquatic Flora and Fauna of
the Tennessee Valley. Water Quality
Series Booklet 4. TVA, Chattanooga,
TN. (k)
Tennessee Valley Authority (TVA). 1988.
Homemade Sampling Equipment. Water
Quality Series Booklet 2. TVA, Chatta-
nooga, TN.
Thorp, J.H. and A.P. Covich, eds. 1991.
Ecology and Classification of North
American Freshwater Invertebrates.
Academic Press, NY. (Especially
Chapter 17 by W.L. Hilsenhoff) (k)
USEPA. 1992. Streamwalk Manual.
March. U.S. Environmental Protection
Agency Region 10, Water Management
Division, Seattle, WA.
USEPA. 1994. Biological Criteria:. Techni-
cal Guidance for Small Streams and
Rivers. EPA 822-B-94-001/U.S. Envi-
ronmental Protection Agency, Office of
Wetlands, Oceans, and Watersheds,
4503F, Washington, DC 20460.
USEPA. 1996. The Volunteer Monitor's
Guide to Quality Assurance Project
Plans. EPA 841-B-96-003. U.S. Envi-
ronmental Protection Agency, Office of
Wetlands, Oceans, and Watersheds,
4503F, Washington, DC 20460.
-------
124 I MACROINVERTEBRATES AND HABITAT
-------
WATER QUALITY CONDITIONS I 125
-------
126 I WATER QUALITY CONDITIONS
Water quality monitoring is
defined here as the sampling
and analysis of water constitu-
ents and conditions. These may include:
• Introduced pollutants, such as
pesticides, metals, and oil
• Constituents found naturally in water
that can nevertheless be affected by
human sources, such as dissolved
oxygen, bacteria, and nutrients
The magnitude of their effects can be
influenced by properties such as pH and
temperature. For example, temperature
influences the quantity of dissolved oxygen
that water is able to contain, and pH affects
the toxicity of ammonia.
Volunteers, as well as state and local
water quality professionals, have been
monitoring water quality conditions for
many years. In fact, until the past decade or
so (when biological monitoring protocols
were developed and began to take hold),
water quality monitoring was generally
considered the primary way of identifying
water pollution problems. Today, profes-
sional water quality specialists and volun-
teer program coordinators alike are moving
toward approaches that combine chemical,
physical, and biological monitoring meth-
ods to achieve the best picture of water
quality conditions.
Water quality monitoring can be used
for many purposes:
• To identify whether waters are
meeting designated uses. All states
have established specific criteria
(limits on pollutants) identifying
what concentrations of chemical
pollutants are allowable in their
waters. When chemical pollutants
exceed maximum or minimum
allowable concentrations, waters
might no longer be able to support
the beneficial uses—such as fishing,
swimming, and drinking—for which
they have been designated. Desig-
nated uses and the specific criteria
that protect them (along with
antidegradation statements that say
waters should not be allowed to
deteriorate below existing or antici-
pated uses) together form water
quality standards. State water quality
professionals assess water quality by
comparing the concentrations of
chemical pollutants found in streams
to the criteria in the state's standards,
and so judge whether streams are
meeting their designated uses.
Water quality monitoring, how-
ever, might be inadequate for
determining whether aquatic life uses
are being met in a stream. While
some constituents (such as dissolved
oxygen and temperature) are impor-
tant to maintaining healthy fish and
aquatic insect populations, other
factors, such as the physical structure
of the stream and the condition of the
habitat, play an equal or greater role.
Biological monitoring methods (see
Chapter 4) are generally better suited
to determining whether aquatic life is
supported.
To identify specific pollutants and
sources of pollution. Water quality
monitoring helps link sources of
pollution to a stream quality problem
because it identifies specific problem
pollutants. Since certain activities
tend to generate certain pollutants
(e.g., bacteria and nutrients are more
likely to come from an animal
feedlot than an automotive repair
shop), a tentative link might be made
that would warrant further investiga-
tion or monitoring.
To determine trends. Chemical
constituents that are properly moni-
tored (i.e., consistent time of day and
on a regular basis, using consistent
methods) can be analyzed for trends
over time.
-------
WATER QUALITY CONDITIONS I 127
• To screen for impairment. Finding
excessive levels of one or more
chemical constituents can serve as an
early warning "screen" of potential
pollution problems.
Designing a water quality
monitoring program
The first step in designing a water
quality monitoring program is to determine
the purpose of the monitoring. This will
help you select which parameters to moni-
tor. The program steering committee should
make this decision based on factors such as:
• Types of water quality problems and
pollution sources that will likely be
encountered (Table 5.1)
• Cost of available monitoring equip-
ment
• Precision and accuracy of available
monitoring equipment
• Capabilities of the volunteers
Because of the expense and difficulty
involved, volunteers generally do not
monitor for toxic substances such as heavy
metals and organic chemicals (e.g., pesti-
cides, herbicides, solvents, and PCBs).
They might, however, collect water
samples for analysis at accredited labs.
The parameters most commonly
monitored by volunteers in streams are
discussed in detail in this chapter. They
include stream flow, dissolved oxygen and
biochemical oxygen demand, temperature,
pH, turbidity, phosphorus, nitrates, total
solids, conductivity, total alkalinity, and
fecal bacteria. Of these, the first five are
the most basic and should form the founda-
tion of almost any volunteer water quality
monitoring program.
Relatively inexpensive and simple-to-
use kits are available from scientific supply
houses to monitor these pollutants. Many
volunteer programs use these kits effec-
tively. Meters and sophisticated lab equip-
ment may be more accurate, but they are
also more expensive, less flexible (e.g.,
meters generally have to be read in the
field), and require periodic calibration. This
chapter discusses specific equipment and
sampling considerations for each param-
eter, and usually describes several ap-
Source
Cropland
Forestry harvest
Grazing land
Industrial discharge
Mining
Septic systems
Common Associated Chemical Pollutants
Sewage treatment plants
Construction
Urban runoff
Turbidity, phosphorus, nitrates, temperature, total solids
Turbidity, temperature, total solids
Fecal bacteria, turbidity, phosphorus, nitrates, temperature
Temperature, conductivity, total solids, toxics, pH
pH, alkalinity, total dissolved solids
Fecal bacteria (i.e., Escherichia coli, enterococcis), nitrates, phosphorus,
dissolved oxygen/biochemical oxygen demand, conductivity, temperature
Dissolved oxygen and biochemical oxygen demand, turbidity, conductivity,
phosphorus, nitrates, fecal bacteria, temperature, total solids, pH
Turbidity, temperature, dissolved oxygen and biochemical oxygen demand,
total solids, and toxics !
Turbidity, phosphorus, nitrates, temperature, conductivity, dissolved oxygen
and biochemical oxygen demand
Table 5.1
•^••I^BBi
Sources and
associated
pollutants
A volunteer
water quality
monitoring
program should
be geared to the
types of water-
shed land uses
most often
encountered.
-------
128 I WATER QUALITY CONDITIONS
Figure 5.1
Sketch of a
Whirl-pak* bag
Volunteers can
be easily trained
to use these
factory-sealed,
disposable
water sample
collection bags.
proaches to monitor them. Table 5-2 lists
methods available for monitoring key
parameters, including the preferred testing
site (lab or field).
General preparation and
sampling considerations
The sections that follow will detail
specific sampling and equipment consider-
ations and analytical procedures for each of
the most common water quality parameters.
There are, however, two general tasks that
are accomplished anytime water samples
are taken. These are discussed below.
Task 1 | Preparation of Sampling
•—«•«••• Containers
Reused sample containers and glass-
ware must be cleaned and rinsed before the
first sampling run and after each run by
following either Method A or Method B
described below. The most suitable method
depends on the parameter being measured.
Method A: General Preparation of Sampling
Containers
The following method should be used
when preparing all sample containers and
glassware for monitoring conductivity, total
solids, turbidity, pH, and total alkalinity.
Wear latex gloves!
1. Wash each sample bottle or piece
UU-
.- Perforation
Wire Tab
--- Pull Tab
of glassware with a brush and
phosphate-free detergent.
2. Rinse three times with cold tap
water.
3. Rinse three times with distilled or
deionized water.
Method B: Acid Wash Procedure for
Preparing Sampling Containers
This method should be used when
preparing all sample containers and glass-
ware for monitoring nitrates and phospho-
rus. Wear latex gloves!
1. Wash each sample bottle or piece
of glassware with a brush and
phosphate-free detergent.
2. Rinse three times with cold tap
water.
3. Rinse with 10 percent hydrochlo-
ric acid.
4. Rinse three times with deionized
water.
Task 2 I Collecting Samples
In general, sample away from the
streambank in the main current. Never
sample stagnant water. The outside curve of
the stream is often a good place to sample
since the main current tends to hug this
bank. In shallow stretches, carefully wade
into the center current to collect the sample.
A boat will be required for deep sites.
Try to maneuver the boat into the center of
the main current to collect the water
sample.
When collecting a water sample for
analysis in the field or at the lab, follow the
steps below.
For Whirl-pak® Bags
1. Label the bag with the site num-
ber, date, and time.
2. Tear off the top of the bag along
the perforation above the wire tab
just prior to sampling (Fig. 5.1).
-------
WATER QUALITY CONDITIONS I 129
Location 1
Method . (Lab or Field) Comments 1
Dissolved Oxygen (DO)
Winkler with eye dropper
Winkler with digital titrator or buret
Meter
Either
Either
Field
If lab, the sample is fixed in field and titrated in lab;
must be measured within 8 hours of collection.
The meter is fragile and must be handled carefully.
Biochemical Oxygen Demand (BOD)
Winkler with eye dropper
Winkler with digital titrator or buret
Meter
1st part -Either
2nd part - Lab
1st part -Either
2nd part - Lab
1st part -Either
2nd part - Lab
If lab, the sample is fixed in field and titrated in lab;
must be measured within 6 hours of Collection.
If lab, the sample is fixed in field and titrated in lab;
must be measured within 6 hours of collection.
The meter is fragile and must be handled carefully;
must be measured within 6 hours of collection.
Temperature
Thermometer
Field
Cannot be done in the lab.
pH
Color comparator
pH "Pocket Pal"
Meter
Either
Either
Either
If lab, measured ASAP within 2 hours of collection.
If lab, measured ASAP within 2 hours of collection.
If lab, measured ASAP within 2 hours of collection.
Turbidity
Meter
Either
If lab, measured within 24 hours of collection.
Total Orthophosphate
Ascorbic acid w/ color comparator
Ascorbic acid w/ spectrophotometer
Either
Either
If lab, measured within 48 hours of collection.
If lab, measured within 48 hours of collection.
Nitrate ;
Cadmium reduction w/ color comparator
Cadmium reduction w/ spectrophotometer
Either
Either
If lab, measured within 48 hours of collection.
If lab, measured within 48 hours of collection.
Total Solids
Oven drying/weighing
Lab
Must be measured within 7 days of collection.
Conductivity
Meter
Either
If lab, measured within 28 days of collection.
Total Alkalinity
Titration
• Either
If lab, measured within 24 hours of collection.
Fecal Bacteria
Membrane filtration
Lab
Must be measured within 6 hours of collection.
Table 5.2
•••^^•••i
Summary of
chemical
monitoring
methods
Volunteers can
measure some
parameters in
the field or in
the laboratory.
-------
130 I WATER QUALITY CONDITIONS
Avoid touching the inside of the
bag. If you accidentally touch the
inside of the bag, use another one.
3. Wading. Try to disturb as little
bottom sediment as possible. In
any case, be careful not to collect
water that contains bottom sedi-
ment. Stand facing upstream.
Collect the water sample in front
of you.
Boat. Carefully reach over the side
and collect the water sample on
the upstream side of the boat.
4. Hold the two white pull tabs in
each hand and lower the bag into
the water on your upstream side
with the opening facing upstream.
Open the bag midway between the
surface and the bottom by pulling
the white pull tabs. The bag should
begin to fill with water. You may
need to "scoop" water into the bag
by drawing it through the water
upstream and away from you. Fill
the bag no more than 3/4 full!
5. Lift the bag out of the water. Pour
out excess water. Pull on the wire
tabs to close the bag. Continue
holding the wire tabs and flip the
bag over at least 4-5 times quickly
to seal the bag. Don't try to
squeeze the air out of the top of
the bag. Fold the ends of the wire
tabs together at the top of the bag,
being careful not to puncture the
bag. Twist them together, forming
a loop.
6. Fill in the bag number and/or site
number on the appropriate field
data sheet. This is important! It is
the only way the lab coordinator
know which bag goes with which
site.
7. If samples are to be analyzed in a
lab, place the sample in the cooler
with ice or cold packs. Take all
samples to the lab.
For Screw-cap Bottles
To collect water samples using screw-
cap sample bottles, use the following
procedures (Fig. 5.2 and 5.3):
Figure 5.2
Getting into
position to
take a water
sample
Volunteers
should sample
in the main
current, facing
upstream.
-------
WATER QUALITY CONDITIONS I 131
2.
4.
Figure 5.3
Taking a water
sample
Turn the bottle
into the current
and scoop in an
upstream
direction.
1. Label the bottle with the site num-
ber, date, and time.
2. Remove the cap from the bottle just
before sampling. Avoid touching the
inside of the bottle or the cap. If you
accidentally touch the inside of the
bottle, use another one.
3. Wading. Try to disturb as little
bottom sediment as possible. In any
case, be careful not to collect water
that has sediment from bottom
disturbance. Stand facing upstream.
Collect the water sample on your
upstream side, in front of you. You
may also tape your bottle to an
extension pole to sample from
deeper water.
Boat. Carefully reach over the side
and collect the water sample on the
upstream side of the boat.
4. Hold the bottle near its base and
plunge it (opening downward)
below the water surface. If you are
using an extension pole, remove the
cap, turn the bottle upside down,
and plunge it into the water, facing
upstream. Collect a water sample 8
to 12 inches beneath the surface or
mid-way between the surface and
the bottom if the stream reach is
shallow.
5. Turn the bottle underwater into the
current and away from you. In
slow-moving stream reaches, push
the bottle underneath the surface
and away from you in an upstream
direction.
6. Leave a 1-inch air space (Except for
DO and BOD samples). Do not fill
the bottle completely (so that the
sample can be shaken just before
analysis). Recap the bottle care-
fully, remembering not to touch the
inside.
7. Fill in the bottle number and/or site
number on the appropriate field data
sheet. This is important because it
tells the lab coordinator which
bottle goes with which site.
8. If the samples are to be analyzed in
the lab, place them in the cooler for
transport to the lab.
-------
132 I WATER QUALITY CONDITIONS
QUALITY ASSURANCE, QUALITY CONTROL, and QUALITY ASSESSMENT MEASURES
Quality assurance/quality control measures are those activities you undertake to demonstrate the accuracy (how close to the real result you
are) and precision (how reproducible your results are) of your monitoring. Quality Assurance (QA) generally refers to a broad plan for maintaining
quality in all aspects of a program. This plan should describe how you will undertake your monitoring effort: proper documentation of all your
procedures, training of volunteers, study design, data management and analysis, and specific quality control measures. Quality Control (QC)
consists of the steps you will take to determine the validity of specific sampling and analytical procedures. Quality assessment is your assess-
ment of the overall precision and accuracy of your data, after you've run the analyses.
Quality Control and Assessment Measures: Internal Checks
Internal checks are performed by the project field volunteers, staff, and lab.
• Reid Blanks. A trip blank (also known as a field blank) is de-ionized water which is treated as a sample. It is used to identify errors or
contamination in sample collection and analysis.
• Negative and Positive Plates (for bacteria). A negative plate results when the buffered rinse water (the water used to rinse down the
sides of the filter funnel during filtration) has been filtered the same way as a sample. This is different from a field blank in that it
contains reagents used in the rinse water. There should be no bacteria growth on the filter after incubation. It is used to detect labora-
tory bacteria contamination of the sample. Positive plates result when water known to contain bacteria (such as wastewater treatment
plant influent) is filtered the same way as a sample. There should be plenty of bacteria growth on the filter after incubation. It is used to
detect procedural errors or the presence of contaminants in the laboratory analysis that might inhibit bacteria growth.
• Reid Duplicates. A field duplicate is a duplicate river sample collected by the same team or by another sampler or team at the same
place, at the same time. It is used to estimate sampling and laboratory analysis precision.
• Lab Replicates. A lab replicate is a sample that is split into subsamples at the lab. Each subsample is then analyzed and the results
compared. They are used to test the precision of the laboratory measurements. For bacteria, they are used to obtain an optimal number
of bacteria colonies on filters for counting purposes.
• Spike Samples. A known concentration of the indicator being measured is added to the sample. This should increase the concentration
in the sample by a predictable amount. It is used to test the accuracy of the method.
• Calibration Blank. A calibration blank is de-ionized water processed like any of the samples and used to "zero" the instrument. It is the
first "sample" analyzed and used to set the meter to zero. This is different from the field blank in that it is "sampled" in the lab. It is used
to check the measuring instrument periodically for "drift" (the instrument should always read "0" when this blank is measured). It can
also be compared to the field blank to pinpoint where contamination might have occurred.
• Calibration Standards. Calibration standards are used to calibrate a meter. They consist of one or more "standard concentrations"
(made up in the lab to specified concentrations) of the indicator being measured, one of which is the calibration blank. Calibration
standards can be used to calibrate the meter before running the test, or they can be used to convert the units read on the meter to the
reporting units (for example, absorbance to milligrams per liter).
Quality Control And Assessment Measures: External Checks
Bctemal checks are performed by non-volunteer field staff and a lab (also known as a "quality control lab"). The results are compared with
those obtained by the project lab.
• External Field Duplicates. An external field duplicate is a duplicate river sample collected and processed by an independent (e.g.,
professional) sampler or team at the same place at the same time as regular river samples. It is used to estimate sampling and
laboratory analysis precision.
• Split Samples. A split sample is a sample that is divided into two subsamples at the lab. One subsample is analyzed at the project lab
and the other is analyzed at an independent lab. The results are compared.
• Outside Lab Analysis of Duplicate Samples. Either internal or external field duplicates can be analyzed at an independent lab. The
results should be comparable with those obtained by the project lab. '
-------
WATER QUALITY CONDITIONS 1133
• Knowns. The quality control lab sends samples for selected indicators, labeled with the concentrations, to the project lab for analysis
• prior to the first sample run. These samples are analyzed and the results compared with the known concentrations. Problems are
reported to the quality control lab.
• iMnowna'The quaiity.control lab sends samples to the project lab for analysis for selected indicators, prior to the first sample run. The
concentrations of these samples are unknown to the project lab. These samples are analyzed and the results reported to the quality
control lab. Discrepancies are reported to the project lab and a problem-identification and solving process follows.
The table below shows the applicability of common quality control measures to the water quality indicators covered in this manual
Steps To Quality Control
1. Consult with your technical committee and/or program advisor to help you determine quality assurance/quality control measures you
will use to answer your questions and meet your data quality requirements
2. Locate a quality control lab—an independent lab that can run external checks for you.
3. Determine which quality checks you have the resources and capabilities to carry out. Your human and financial resources and expertise
might limit the water quality indicators your can monitor. :
References
APHA. 1992. Standard Methods for the Examination of Water and Wastewater. 18th ed. American Public Health Association, Washington, DC.
Intergovernmental Task Force on Monitoring Water Quality. 1994. Water quality monitoring in the United States. 1993 report and technical
appendixes. Washington, DC.
Mattspn, M. 1992. The basics of quality control. The Volunteer Monitor. 4(2) Fall 1992.
USEPA. 1983. Methods for chemical analysis of water, and wastes. EPA-600/4-79-020. U.S. Environmental Protection Agency, Environmental
Monitoring and Support Laboratory, Cincinnati, OH. March.
USEPA. 1984. Guidance for preparation of combined work/quality assurance project plans for environmental monitoring. ORWS QA-1, U.S.
Environmental Protection Agency, Office of Water Regulations and Standards. Washington DC, May.
USEPA. 1996. The Volunteer Monitor's Guide to Quality Assurance Project Plans. EPA-841-B-96-003. Environmental Protection Agency, Office
of Water, Washington, DC.
Common Quality Control Measures
Dissolved terno- Tur- Phos-
Oxygen erature pH bidity phorus
Total Con- Total Fecal
Nitrates Solids ductivity Alkalinity Bacteria
Internal Checks
Field blanks
Field duplicates
Lab replicates
Positive plates
Negative plates
Spike samples
Calibration blank
Calibration standard
External Checks
External field duplicates
Split samples
Outside lab analysis
Verification
Knowns
Unknowns
/
/
/
/
/
/
a - using an oxygen-saturated sample
b - using subsamples of different sizes
-------
134 I WATER QUALITY CONDITIONS
5.1
Stream Flow
What is stream flow and why is it
important?
Stream flow, or discharge, is the
volume of water that moves over a desig-
nated point over a fixed period of time. It is
often expressed as cubic feet per second
(ftVsec).
The flow of a stream is directly related
to the amount of water moving off the
watershed into the stream channel. It is
affected by weather, increasing during
rainstorms and decreasing during dry
periods. It also changes during different
seasons of the year, decreasing during the
summer months when evaporation rates are
high and shoreline vegetation is actively
growing and removing water from the
ground. August and September are usually
the months of lowest flow for most streams
and rivers in most of the country.
Water withdrawals for irrigation
purposes can seriously deplete water flow,
as can industrial water withdrawals. Dams
used for electric power generation, particu-
larly facilities designed to produce power
during periods of peak need, often block the
flow of a stream and later release it in a
surge.
Flow is a function of water volume and
velocity. It is important because of its
impact on water quality and on the living
organisms and habitats in the stream. Large,
swiftly flowing rivers can receive pollution
discharges and be little affected, whereas
small streams have less capacity to dilute
and degrade wastes.
Stream velocity, which increases as the
volume of the water in the stream increases,
determines the kinds of organisms that can
live in the stream (some need fast-flowing
areas; others need quiet pools). It also
affects the amount of silt and sediment
carried by the stream. Sediment introduced
to quiet, slow-flowing streams will settle
quickly to the stream bottom. Fast moving
streams will keep sediment suspended
longer in the water column. Lastly, fast-
moving streams generally have higher ,
levels of dissolved oxygen than slow
streams because they are better aerated.
This section describes one method for
estimating flow in a specific area or reach
of a stream. It is adapted from techniques
used by several volunteer monitoring
programs and uses a float (an object such as
an orange, ping-pong ball, pine cone, etc.)
to measure stream velocity. Calculating
flow involves solving an equation that
examines the relationship among several
variables including stream cross-sectional
area, stream length, and water velocity.
One way to measure flow is to solve the
following equation:
Flow =
ALC
Where:
A = Average cross-sectional area of the
stream (stream width multiplied by
average water depth).
L = Length of the stream reach mea-
sured (usually 20 ft.)
C = A coefficient or correction factor (0.8
for rocky-bottom streams or 0.9 for
muddy-bottom streams). This allows
you to correct for the fact that water
at the surface travels faster than
near the stream bottom due to
resistance from gravel, cobble, etc.
Multiplying the surface velocity by a
correction coefficient decreases the
value and gives a better measure of
the stream's overall velocity.
T = Time, in seconds, for the float to
travel the length of L
-------
WATER QUALITY CONDITIONS I 135
How to Measure and Calculate
Stream Flow
TASK1
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, when measuring and calculat-
ing flow, include the following equipment:
• Ball of heavy-duty string, four
stakes, and a hammer to drive the
stakes into the ground. The string
will be stretched across the width of ,
the stream perpendicular to shore at
two locations. The stakes are to
anchor the string on each bank to
form a transect line.
• Tape measure (at least 20 feet)
• Waterproof yardstick or other
implement to measure water depth
• Twist ties (to mark off intervals on
the string of the transect line)
• An orange and a fishing net (to scoop
the orange out of the stream)
• Stopwatch (or watch with a second
hand)
• Calculator (optional)
TASK 2
Select a stretch of stream
The stream stretch chosen for the
measurement of discharge should be
straight (no bends), at least 6 inches deep,
and should not contain an area of slow
water such as a pool. Unobstructed riffles
or runs are ideal. The length that you select
will be equal to L in solving the flow
equation. Twenty feet is a standard length
used by many programs. Measure your
length and mark the upper and lower end by
running a transect line across the stream
perpendicular to the shore using the string
and stakes (Fig. 5.4). The string should be
taut and near the water surface. The
upstream transect is Transect #1 and the
downstream one is Transect #2.
TASK 3 I Calculate the average cross-
™~~"™~™"^ sectional area
Cross-sectional area (A in the formula)
is the product of stream width multiplied
by average water depth. To calculate the
average cross-sectional area for the study
stream reach, volunteers should determine
the cross-sectional area for each transect,
add the results together, and then divide by
2 to determine the average cross-sectional
area for the stream reach.
To measure cross-sectional area:
1. Determine the average depth along
the transect by marking off equal
intervals along the string with the
twist ties. The intervals can be one-
fourth, one-half, and three-fourths of
the distance across the stream.
Measure the water's depth at each
interval point (Fig. 5.5). To calculate
average depth for each transect,
divide the total of the three depth
measurements by 4. (You divide by
4 instead of 3 because you need to
account for the 0 depths that occur at
the shores.) In the example shown in
Figure 5.4
A diagram of a
20- foot
transect
-------
136 I WATER QUALITY CONDITIONS
•TOTAL MOTH VFEET-
INTtlWtl- WIDTH
A cross sec-
tion view to
measure
stream width
and depth
Figure 5.6
A sample
calculation of
average cross-
sectional area.
Figure 5.6, the average depth of
Transect #1 is 0.575 feet and the
average depth of Transect #2 is 0.625
feet.
2. Determine the width of each transect
by measuring the distance from
shoreline to shoreline. Simply add
together all the interval widths for
each transect to determine its width.
In the Figure 5.6 example, the width
of Transect #1 is 8 feet and the width
of Transect #2 is 10 feet.
3. Calculate the cross-sectional area of
each transect by multiplying width
times average depth. The example
given in Figure 5.6 shows that the
average cross-sectional area of
Transect #1 is 4.60 square feet and
the average cross-sectional area of
Transect #2 is 6.25 square feet.
4. To determine the average cross-
sectional area of the entire stream
reach (A in the formula), add to-
gether the average cross-sectional
area of each transect arid then divide
by 2. The average cross-sectional
area for the stream reach in Figure
5.6 is 5.42 square feet.
Determining Average C
Transect #1 (upstream)
Interval width Depth
(feet) (feet)
AtoB = 2.0 1.0 (atB)
BtoC = 2.0 0.8 (atC)
CtoD = 2.0 0.5 (atD)
D to E = 2.0 0.0 (shoreline)
Totals 8.0 2.3
Average depth = 2.3 / 4 = 0.575 feet
Cross-sectional area of Transect #1
= Total width X Average depth
= 8 ft X 0.575
= 4.60 ft2
Average area = (Cross-sectional area of Transe
= (4.60 ft
= 5.42 ft2
ross-Sectional Area (A)
Transect #2 (downstream)
Interval width Depth
(feet) (feet)
AtoB = 2.5 1.1 (atB)
BtoC = 2.5 1.0 (atC)
CtoD = 2.5 0.4 (atD)
D to E = 2.5 0.0 (shoreline)
10.0 2.5
Average depth = 2.5 / 4 = 0.625 feet
Cross-sectional area of Transect #2
= Total width X Average depth
= 10.0 ft X 0.625
= 6.25 ft2
ct #1 + Cross-sectional area of Transect #2) / 2
! + 6.25 ft2)/ 2
-------
WATER QUALITY CONDITIONS I 137
Task 4
Measure travel time
Volunteers should time with a stop-
watch how long it takes for an orange (or
some other object) to float from the up-
stream to the downstream transect. An
orange is a good object to use because it
has enough buoyancy to float just below the
water surface. It is at this position that
maximum velocity typically occurs.
The volunteer who lets the orange go at
the upstream transect should position it so it
flows into the fastest current. The clock
stops when the orange passes fully under
the downstream transect line. Once under
the transect line, the orange can be scooped
out of the water with the fishing net. This
"time of travel" measurement should be
conducted at least three times and the
results averaged—the more trials you do,
the more accurate your results will be. The
averaged results are equal to T in the
formula. It is a good idea to float the orange
at different distances from the bank to get
various velocity estimates. You should
discard any float trials if the object gets
hung up in the stream (by cobbles, roots,
debris, etc.)
Task 5 I Calculate flow
Recall that flow can be calculated
using the equation:
ALC
Flow =
Flow =
(5.42 ft ) (20 ft) (0.8)
15 sec.
86.72 ft
15 sec.
Flow = 5,78 ft3/sec.
Task6
Record flow on the data form
On the following page is a form
volunteers can use to calculate flow of a
stream.
References
Adopt-A-Stream Foundation. Field Guide:
Watershed Inventory and Stream Moni-
toring Methods, by Tom Murdoch and
Martha Cheo. 1996. Everett, .WA.
Mitchell, M.K., and W. Stapp. Field
Manual for Water Quality Monitoring.
5th Edition. Thompson Shore; Printers.
Missouri Stream Teams. Volunteer Water
Quality Monitoring. Missouri Depart-
ment of Natural Resources, P.O. Box
176, Jefferson City, MO 65102.
Flow =
Continuing the example in Fig. 5.6. say
the average time of travel for the orange
between Transect #1 and #2 is 15 seconds
and the stream had a rocky bottom. The
calculation of flow would be:
A = 5.42 ft2
L = 20 ft
C = 0.8 (coefficient for a rocky-
bottom stream)
T = 15 seconds
-------
138 I WATER QUALITY CONDITIONS
DATA FORM FOR CALCULATING FLOW
Solving the equation: Flow =
ALC
Where:
A = Average cross-sectional area of the stream. L = Length of the stream reach measured (usually 20 ft.).
C = A coefficient or correction factor (0.8 for rocky-bottom streams or 0.9 for muddy-bottom streams). T = Time, in
seconds, for the float to travel the length of L.
A: Average Cross-Sectional Area
Transect #1 (upstream)
Depth
(feet)
(atB)
(at C)
(atD)
Interval width
(feet)
AtoB =
B to C =
C to D =
D to E =
(shoreline)
Totals I |
= Avg. depth I I ft
Cross-sectional area of Transect #1
= Total width (ft) X Avg. depth (ft)
EH X I""1 • I I ff
Transect #2 (downstream)
Interval width
(feet)
A to B =
BtoC =
C to D = .
D to E =
Totals
Depth
(feet)
(atB)
(atC)
(atD)
(shoreline)
n
= Avg. depth
Cross-sectional area of Transect #2
= Total width (ft) X Avg. depth (ft)
n x n = d
ft2
(Cross-sectional area of Transect #1 + Cross-sectional area of Transect #2) •*• 2 = Average Cross-sectional area
I I X [ I =
ft2
L: Length of Stream Reach
|
C: Coefficient
||
ft
T: Travel Time Travel Time
of Float (sec.)
Trial #1
Trial #2
Trial #3
Total | ~\ •* 3
= Avg. time I | sec.
Flow =
ALC
ftVsec.
-------
WATER QUALITY CONDITIONS I 139
5.2
Dissolved Oxygen and
Biochemical Oxygen
Demand
What is dissolved oxygen and
why is it important?
The stream system both produces and
consumes oxygen. It gains oxygen from the
atmosphere and from plants as a result of
photosynthesis. Running water, because of
its churning, dissolves more oxygen than
still water, such as that in a reservoir behind
a dam. Respiration by aquatic animals,
decomposition, and various chemical
reactions consume oxygen.
Wastewater from sewage treatment
plants often contains organic materials that
are decomposed by microorganisms, which
use oxygen in the process. (The amount of
oxygen consumed by these organisms in
breaking down the waste is known as the
biochemical oxygen demand or BOD. A
discussion of BOD and how to monitor it is
included at the end of this section.) Other
sources of oxygen-consuming waste
include stormwater runoff from farmland or
urban streets, feedlots, and failing septic
systems.
Oxygen is measured in its dissolved
form as dissolved oxygen (DO). If more
oxygen is consumed than is produced,
dissolved oxygen levels decline and some
sensitive animals may move away, weaken,
or die.
DO levels fluctuate seasonally and over
a 24-hour period. They vary with water
temperature and altitude. Cold water holds
more oxygen than warm water (Table 5.3)
and water holds less oxygen at higher
altitudes. Thermal discharges, such as water
used to cool machinery in a manufacturing
plant or a power plant, raise the tempera-
ture of water and lower its oxygen content.
Aquatic animals are most vulnerable to
lowered DO levels in the early morning on
hot summer days when stream flows are
low, water temperatures are high, and
aquatic plants have not been producing
oxygen since sunset. ;
Sampling and Equipment
Considerations
In contrast to lakes, where DO levels
are most likely to vary vertically in the
water column, the DO in rivers and streams
changes more horizontally along the course
of the waterway. This is especially true in
smaller, shallower streams. In larger,
deeper rivers, some vertical stratification of
dissolved oxygen might occur. The DO
levels in and below riffle areas,, waterfalls,
or dam spillways are typically higher than
those in pools and slower-moving
stretches. If you wanted to measure the
effect of a dam, it would be important to
sample for DO behind the dam,' immedi-
ately below the spillway, and upstream of
the dam. Since DO levels are critical to
fish, a good place to sample is in the pools
that fish tend to favor or in the spawning
areas they use.
An hourly time profile of DO levels at
a sampling site is a valuable set of data
because it shows the change in JDO levels
from the low point just before sunrise to
the high point sometime in the midday.
However, this might not be practical for a
volunteer monitoring program. It is impor-
tant to note the time of your DO sampling
to help judge when in the daily cycle the
data were collected.
DO is measured either in milligrams
per liter (mg/L) or "percent saturation."
Milligrams per liter is the amount of
oxygen in a liter of water. Percent satura-
tion is the amount of oxygen in a liter of
water relative to the total amount of oxygen
that the water can hold at that temperature.
-------
140 I WATER QUALITY CONDITIONS
Temperature DO Temperature DO
fC) (mg/L) (°C) (mg/L)
0
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
14.60
14.19
13.81
13.44
13.09
12.75
12.43
12.12
11.83
11.55
11.27
11.01
10.76
10.52
10.29
10.07
9.85
9.65
9.45
9.26
9.07
8.90
8.72
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
8.56
8.40
8.24
8.09
7.95
7.81
7.67
7.54
7.41
7.28
7.16
7.05
6.93
6.82
6.71
6.61 '
6.51
6.41
6.31
6.22
6.13
6.04
5.95
Table 5.3
Maximum
dissolved
oxygen con-
centrations
vary with
temperature
DO samples are collected using a
special BOD bottle: a glass bottle with a
"turtleneck" and a ground glass stopper.
You can fill the bottle directly in the stream
if the stream is wadable or boatable, or you
can use a sampler that is dropped from a
bridge or boat into water deep enough to
submerse the sampler. Samplers can be
made or purchased.
Dissolved oxygen is measured prima-
rily either by using some variation of the
Winkler method or by using a meter and
probe.
Winkler Method
The Winkler method involves filling a
sample bottle completely with water (no air
is left to bias the test). The dissolved
oxygen is then "fixed" using a series of
reagents that form an acid compound that is
titrated. Titration involves the drop-by-drop
addition of a reagent that neutralizes the
acid compound and causes a change in the
color of the solution. The point at which the
color changes is the "endpoint" and is
equivalent to the amount of oxygen dis-
solved in the sample. The sample is usually
fixed and titrated in the field at the sample
site. It is possible, however, to prepare the
sample in the field and deliver it to a lab for
titration.
Dissolved oxygen field kits using the
Winkler method are relatively inexpensive,
especially compared to a meter and probe.
Field kits run between $35 and $200, and
each kit comes with enough reagents to run
50 to 100 DO tests. Replacement reagents
are inexpensive, and you can buy them
already measured out for each test in plastic
pillows.
You can also buy the reagents in larger
quantities, in bottles, and measure them out
with a volumetric scoop. The advantage of
the pillows is that they have a longer shelf
life and are much less prone to contamina-
tion or spillage. The advantage of buying
larger quantities in bottles is that the cost
per test is considerably less.
The major factor in the expense of the
kits is the method of titration they use—
eyedropper, syringe-type titrator, or digital
titrator. Eyedropper and syringe-type
titration is less precise than digital titration
because a larger drop of titrant is allowed to
pass through the dropper opening and, on a
micro-scale, the drop size (and thus the
volume of titrant) can vary from drop to
-------
WATER QUALITY CONDITIONS I 141
drop. A digital titrator or a buret (which is a
long glass tube with a tapered tip like a
pipet) permits much more precision and
uniformity in the amount of titrant that is
allowed to pass.
If your program requires a high degree
of accuracy and precision in DO results, use
a digital titrator. A kit that uses an eye
dropper-type or syringe- type titrator is
suitable for most other purposes. The lower
cost of this type of DO field kit might be
attractive if you are relying on several
teams of volunteers to sample multiple sites
at the same time.
Meter and Probe
A dissolved oxygen meter is an elec-
tronic device that converts signals from a
probe that is placed in the water into units
of DO in milligrams per liter. Most meters
and probes also measure temperature. The
probe is filled with a salt solution and has a
selectively permeable membrane that
allows DO to pass from the stream water
into the salt solution. The DO that has
diffused into the salt solution changes the
electric potential of the salt solution and
this change is sent by electric cable to the
meter, which converts the signal to milli-
grams per liter on a scale that the volunteer
can read.
DO meters are expensive compared to
field kits that use the titration method.
Meter/probe combinations run between
$500 and $1,200, including a long cable to
connect the probe to the meter. The advan-
tage of a meter/probe is that you can
measure DO and temperature quickly at any
point in the stream that you can reach with
the probe. You can also measure the DO
levels at a certain point on a continuous
basis. The results are read directly as
milligrams per liter, unlike the titration
methods, in which the final titration result
might have to be converted by an equation
to milligrams per liter.
However, DO meters are more fragile
than field kits, and repairs to a damaged
meter can be costly. The meter/probe must
be carefully maintained, and it must be
calibrated before each sample 'run and, if
you are doing many tests, in between
samplings. Because of the expense, a
volunteer program might have only one
meter/probe. This means that only one
team of samplers can sample DO and they
will have to do all the sites. With field kits,
on the other hand, several teams can
sample simultaneously. '.
Laboratory Testing of Dissolved Oxygen
If you use a meter and probe, you must
do the testing in the field; dissolved oxygen
levels in a sample bottle change quickly
due to the decomposition of organic
material by microorganisms or the produc-
tion of oxygen by algae and other plants in
the sample. This will lower your DO
reading. If you are using a variation of the
Winkler method, it is possible ;to "fix" the
sample in the field and then deliver it to a
lab for titration. This might be! preferable if
you are sampling under adverse conditions
or if you want to reduce the time spent
collecting samples. It is also a little easier
to titrate samples in the lab, and more
quality control is possible because the same
person can do all the titrations.
How to collect and analyze
samples
The procedures for collecting and
analyzing samples for dissolved oxygen
consist of the following tasks:'.
TASK1
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, when sampling for dissolved
oxygen, include the following equipment:
-------
142 I WATER QUALITY CONDITIONS
If Using the Winkler Method
• Labels for sample bottles
• Field kit and instructions for DO
testing
• Enough reagents for the number of
sites to be tested
• Kemmerer, Van Dorn, or home-made
sampler to collect deep-water
samples
• A numbered glass BOD bottle with a
glass stopper (1 for each site)
• Data sheet for dissolved oxygen to
record results
If Using a Meter and Probe
• DO meter and probe (electrode)
(NOTE: Confirm that the meter has
been calibrated according to the
manufacturer's instructions.)
• Operating manual for the meter and
probe
• Extra membranes and electrolyte
solution for the probe
• Extra batteries for the meter
• Extension pole
• Data sheet for dissolved oxygen to
record results
| TASK 2 I Confirm that you are at the
^^"^""""^ proper location
The directions for sampling should
provide specific information about the
exact point in the stream from which you
are to sample; e.g., "approximately 6 feet
out from the large boulder downstream
from the west side of the bridge." If you are
not sure you are in the exact spot, record a
detailed description of where you took the
sample so that it can be compared to the
actual site later.
TASK 3
Collect samples and fill out
the field data sheet
Winkler Method
Use a BOD bottle to collect the water
sample. The most common sizes are 300
milliliters (mL) and 60 mL. Be sure that
you are using the correct volume for the
titration method that will be used to deter-
mine the amount of DO. There is usually a
white label area on the bottle, and this may
already be numbered. If so, be sure to
record that number on the field data sheet.
If your bottle is not already numbered,
place a label on the bottle (not on the cap
because a cap can be inadvertently placed
on a different bottle) and use a waterproof
marker to write in the site number.
If you are collecting duplicate samples,
label the duplicate bottle with the correct
code, which should be determined prior to
sampling by the lab supplying the bottles.
Use the following procedure for collecting
a sample for titration by the Winkler
method:
1. Remember that the water sample
must be collected in such a way that
you can cap the bottle while it is still
submerged. That means that you
must be able to reach into the water
with both arms and the water must be
deeper than the sample bottle.
2. Carefully wade into the stream.
Stand so that you are facing one of
the banks.
3. Collect the sample so that you are
not standing upstream of the bottle.
Remove the cap of the BOD bottle.
Slowly lower the bottle into the
water, pointing it downstream, until
the lower lip of the opening is just
submerged. Allow the water to fill
the bottle very gradually, avoiding
any turbulence (which would add
oxygen to the sample). When the
-------
WATER QUALITY CONDITIONS I 143
Figure 5.7
Taking a water
sample for DO
analysis
Point the bottle
downstream
and fill gradu-
ally. Cap
underwater
when full.
water level in the bottle has stabi-
lized (it won't be full because the
bottle is tilted), slowly turn the bottle
upright and fill it completely. Keep
the bottle under water and allow it to
overflow for 2 or 3 minutes to ensure
that no air bubbles are trapped.
4. Cap the bottle while it is still sub-
merged. Lift it out of the water and
look around the "collar" of the bottle
just below the bottom of the stopper.
If you see an air bubble, pour out the
sample and try again.
5. "Fix" the sample immediately
following the directions in your kit:
• Remove the stopper and add the
fixing reagents to the sample.
• Immediately insert the stopper so
air is not trapped in the bottle and
invert several times to mix. This
solution is caustic. Rinse your
hands if you get any solution on
them. An orange-brown flocculent
precipitate will form if oxygen is
present.
• Wait a few minutes until the floe
in the solution has settled. Again
invert the bottle several times and
wait until the floe has settled. This
ensures complete reaction of the
sample and reagents. The sample
is now fixed, and atmospheric
oxygen can no longer affect it.
If you are taking the sample to the
lab for titration, no further action is
necessary. You can store the sample
in a cooler for up to 8 hours before
titrating it in a lab. If you are titrat-
ing the sample in the field, see Task
4: Analyze the Samples.
Using a DO Meter
If you are using a dissolved oxygen
meter, be sure that it is calibrated immedi-
ately prior to use. Check the cable connec-
tion between the probe and the meter.
Make sure that the probe is filled with
electrolyte solution, that the membrane has
no wrinkles, and that there are no bubbles
trapped on the face of the membrane. You
can do a field check of the meter's accu-
racy by calibrating it in saturated air
according to the manufacturer's instruc-
tions. Or, you can measure a water sample
that is saturated with oxygen, as follows.
(NOTE: You can also use this procedure
for testing the accuracy of the Winkler
method.)
1. Fill a 1-liter beaker or bucket half full
of tap water. (You may want to bring
a gallon jug with water in it for this
purpose.) Mark the. bottle number as
"tap" on the lab sheet.
2. Pour this water back and forth into
another beaker 10 times to saturate
the water with oxygen.
-------
144 I WATER QUALITY CONDITIONS
3. Use the meter to measure the water
temperature and record it in the water
temperature column on the field data
sheet.
4. Find the water temperature of your
"tap" sample in Table 5.3. Use the
meter to compare the dissolved
oxygen concentration of your sample
with the maximum concentration at
that temperature in the table. Your
sample should be within 0.5 mg/L. If
it is not, repeat the check and if there
is still an error, check the meter's
batteries and follow the troubleshoot-
ing procedures in the manufacturer's
manual.
Once the meter is turned on, allow 15
minute equilibration before calibrating.
After calibration, do not turn the meter off
until the sample is analyzed. Once you have
verified that the meter is working properly,
you are ready to measure the DO levels at
the sampling site.
Figure 5.8
Titrating a DO
sample using a
buret
You might need an extension pole (this
can be as simple as a piece of wood) to get
the probe to the proper sampling point.
Simply secure the probe to the end of the
extension pole. A golfer's ball retriever
works well because it is collapsible and
easy to transport. To use the probe, proceed
as follows:
1. Place the probe in the stream below
the surface.
2. Set the meter to measure tempera-
ture, and allow the temperature
reading to stabilize. Record the
temperature on the field data sheet.
3. Switch the meter to read dissolved
oxygen.
4. Record the dissolved oxygen level on
the field data sheet.
TASK 4 | Analyze the samples
Three types of titration apparatus can
be used with the Winkler method: droppers,
digital titrators, and burets. The dropper and
digital titrator are suited for field use. The
buret is more conveniently used in the lab
(Fig. 5.8) Volunteer programs are most
likely to use the dropper or digital titrator.
For titration with a dropper or syringe,
which is relatively simple, follow the
manufacturer's instructions. The following
procedure is for using a digital titrator to
determine the quantity of dissolved oxygen
in a fixed sample:
1. Select a sample volume and sodium
thiosulfate titration cartridge for the
digital titrator corresponding to the
expected dissolved oxygen concen-
tration according to Table 5.4. In
most cases, you will use the 0.2 N
cartridge and the 100-mL sample
volume.
2. Insert a clean delivery tube into the
titration cartridge.
3. Attach the cartridge to the titrator
body.
-------
WATER QUALITY CONDITIONS I 145
4. Hold the titrator with the cartridge tip
up. Turn the delivery knob to eject
air and a few drops of titrant. Reset
the counter to 0 and wipe the tip.
5. Use a graduated cylinder to measure
the sample volume (from the "fixed"
sample in the 300-mL BOD bottle)
according to Table 5.4.
6. Transfer the sample into a 250-mL
Erlenmeyer flask, and place the flask
on a magnetic stirrer with a stir bar.
If you are in the field, you can
manually swirl the flask to mix.
7. Place the delivery tube tip into the
solution and turn the stirrer on to stir
the sample while you're turning the
delivery knob.
8. Titrate to a pale yellow color.
9. Add two dropperfuls of starch
indicator solution and swirl to mix. A
strong blue color will develop.
10. Continue to titrate until the sample is
clear. Record the number of digits
required. (The color might reappear
after standing a few minutes, but this
is not a cause for concern. The "first"
disappearance of the blue color is
considered the endpoint.)
11. Calculate mg/L of DO = digits
required X digit multiplier (from
Table 5.4).
12. Record the results in the appropriate
column of the data sheet.
Some water quality standards are
expressed in terms of percent saturation. To
calculate percent saturation of the sample:
1. Find the temperature of your water
sample as measured in the field.
2. Find the maximum concentration of
your sample at that temperature as
given in Table 5.3.
3. Calculate the percent saturation, by
dividing your actual dissolved
oxygen by the maximum concentra-
tion at the sample temperature.
Expected . Sample ' Titration Digit
Range Volume . Cartridge Multiplier
•••••••••••••••••••••••••i
1-5 mg/L
2-10 mg/L
10+ mg/L
200 mL ;
100mL ;
200 mL
0.2 N
0.2 N
2.0 N
0.01
0.02
0.10
Example: You measured a dissolved
oxygen concentration of 5 mg/L at 20 °C.
Divide 5 mg/L by 9.07, the maximum
concentration at 20 °C. The percent
saturation would be 55 percent.
4. Record the percent saturation in the
appropriate column on the data
sheet.
TASKS
Return the samples and the
field data sheets to the lab/
drop-off point
If you are using the Winkler method
and delivering the samples to \SL lab for
titration, double-check to make sure that
you have recorded the necessary informa-
tion for each site on the field data sheet,
especially the bottle number and corre-
sponding site number and the times the
samples were collected. Deliver your
samples and field data sheets to the lab. If
you have already obtained the dissolved
oxygen results in the field, send the data
sheets to your sampling coordinator.
What /s biochemical oxygen
demand and why /s it important?
Biochemical oxygen demand, or BOD,
measures the amount of oxygen consumed
by microorganisms in decomposing
organic matter in stream water. BOD also
measures the chemical oxidatjon of inor-
ganic matter (i.e., the extraction of oxygen
from water via chemical reaction). A test
is used to measure the amount of oxygen
consumed by these organisms during a
Table 5.4
Sample volume
selection and
corresponding
values for
Winkler titra-
tion
-------
146 I WATER QUALITY CONDITIONS
specified period of time (usually 5 days at
20 °C). The rate of oxygen consumption in
a stream is affected by a number of vari-
ables: temperature, pH, the presence of
certain kinds of microorganisms, and the
type of organic and inorganic material in
the water.
BOD directly affects the amount of
dissolved oxygen in rivers and streams. The
greater the BOD, the more rapidly oxygen
is depleted in the stream. This means less
oxygen is available to higher forms of
aquatic life. The consequences of high
BOD are the same as those for low dis-
•solved oxygen: aquatic organisms become
stressed, suffocate, and die.
Sources of BOD include leaves and
woody debris; dead plants and animals;
animal manure; effluents from pulp and
paper mills, wastewater treatment plants,
feedlots, and food-processing plants; failing
septic systems; and urban stormwater
runoff.
Sampling Considerations
BOD is affected by the same factors
that affect dissolved oxygen (see above).
Aeration of stream water—by rapids and
waterfalls, for example—will accelerate the
decomposition of organic and inorganic
material. Therefore, BOD levels at a
sampling site with slower, deeper waters
might be higher for a given volume of
organic and inorganic material than the
levels for a similar site in highly aerated
waters.
Chlorine can also affect BOD measure-
ment by inhibiting or killing the microor-
ganisms that decompose the organic and
inorganic matter in a sample. If you are
sampling in chlorinated waters, such as
those below the effluent from a sewage
treatment plant, it is necessary to neutralize
the chlorine with sodium thiosulfate. (See
APHA, 1992.)
BOD measurement requires taking two
samples at each site. One is tested immedi-
ately for dissolved oxygen, and the second
is incubated in the dark at 20 °C for 5 days
and then tested for the amount of dissolved
oxygen remaining. The difference in
oxygen levels between the first test and the
second test, in milligrams per liter (mg/L),
is the amount of BOD. This represents the
amount of oxygen consumed by microor-
ganisms to break down the organic matter
present in the sample bottle during the
incubation period. Because of the 5-day
incubation, the tests should be conducted in
a laboratory.
Sometimes by the end of the 5-day
incubation period the dissolved oxygen
level is zero. This is especially true for
rivers and streams with a lot of organic
pollution. Since it is not known when the
zero point was reached, it is not possible to
tell what the BOD level is. In this case it is
necessary to dilute the original sample by a
factor that results in a final dissolved
oxygen level of at least 2 mg/L. Special
dilution water should be used for the
dilutions. (See APHA, 1992.)
It takes some experimentation to
determine the appropriate dilution factor for
a particular sampling site. The final result is
the difference in dissolved oxygen between
the first measurement and the second after
multiplying the second result by the dilu-
tion factor. More details are provided in the
following section.
How to Collect and Analyze
Samples
The procedures for collecting samples
for BOD testing consist of the same steps
described for sampling for dissolved
oxygen (see above), with one important
difference. At each site a second sample is
collected in a BOD bottle and delivered to
the lab for DO testing after the 5-day
incubation period. Follow the same steps
used for measuring dissolved oxygen with
these additional considerations:
-------
WATER QUALITY CONDITIONS I 147
• Make sure you have two BOD
bottles for each site you will sample.
The bottles should be black to
prevent photosynthesis. You can
wrap a clear bottle with black
electrician's tape if you do not have a
bottle with black or brown glass.
• Label the second bottle (the one to be
incubated) clearly so that it will not
be mistaken for the first bottle.
• Be sure to record the information for
the second bottle on the field data
sheet.
The first bottle should be analyzed just
prior to storing the second sample bottle in
the dark for 5 days at 20 °C. After this time,
the second bottle is tested for dissolved
oxygen using the same method that was
used for the first bottle. The BOD is
expressed in milligrams per liter of DO
using the following equation:
DO (mg/L) of first bottle
- DO (mg/L) of second bottle
= BOD (mg/L)
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
-------
148 I WATER QUALITY CONDITIONS
5.3
Temperature
Why is temperature important?
The rates of biological and chemical
processes depend on temperature. Aquatic
organisms from microbes to fish are
dependent on certain temperature ranges for
their optimal health. Optimal temperatures
for fish depend on the species: some
survive best in colder water, whereas others
prefer warmer water. Benthic macroinverte-
brates are also sensitive to temperature and
will move in the stream to find their
optimal temperature. If temperatures are
outside this optimal range for a prolonged
period of time, organisms are stressed and
can die. Temperature is measured in de-
grees Fahrenheit (F) or degrees Celsius (C).
For fish, there are two kinds of limiting
temperatures—the maximum temperature
for short exposures and a weekly average
temperature that varies according to the
time of year and the life cycle stage of the
fish species. Reproductive stages (spawning
and embryo development) are the most
sensitive stages. Table 5.5 provides tem-
perature criteria for some species.
Temperature affects the oxygen content
of the water (oxygen levels become lower
as temperature increases); the rate of
photosynthesis by aquatic plants; the
metabolic rates of aquatic organisms; and
the sensitivity of organisms to toxic wastes,
parasites, and diseases.
Causes of temperature change include
weather, removal of shading streambank
vegetation, impoundments (a body of water
confined by a barrier, such as a dam), dis-
charge of cooling water, urban storm water,
and groundwater inflows to the stream.
Sampling and Equipment
Considerations
Temperature in a stream will vary with
width and depth. It can be significantly
different in the shaded portion of the water
on a sunny day. In a small stream, the
temperature will be relatively constant as
long as the stream is uniformly in sun or
shade. In a large stream, temperature can
vary considerably with width and depth
regardless of shade. If it is safe to do so,
temperature measurements should be
collected at varying depths and across the
surface of the stream to obtain vertical and
horizontal temperature profiles. This can be
done at each site at least once to determine
the necessity of collecting a profile during
each sampling visit. Temperature should be
measured at the same place every time.
Temperature is measured in the stream
with a thermometer or a meter. Alcohol-
filled thermometers are preferred over
mercury-filled because they are less hazard-
ous if broken. Armored thermometers for
field use can withstand more abuse than
unprotected glass thermometers and are
worth the additional expense. Meters for
other tests, such as pH (acidity) or dis-
solved oxygen, also measure temperature
and can be used instead of a thermometer.
How to sample
The procedures for measuring tempera-
ture consist of the following tasks.
TASK1
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, when measuring temperature
you will need:
• A thermometer or meter
• A data sheet for temperature to
record results
-------
WATER QUALITY CONDITIONS I 149
Species
Max. weekly
average temp.
for growth
(juveniles)
Max. temp, for,
survival of
short exposure
(juveniles)
Table 5.5
Max. weekly
average temp.
for spawninga
Max. temp.
for embryo
spawning b
Atlantic salmon
Bluegill
Brook trout
Common carp
Channel catfish
Largemouth bass
Rainbow trout
Smallmouth bass
Sockeye salmon
20°C (68°F)
32°C (90°F)
19°C (66°F)
...
32°C (90°F)
32°C (90°F)
19°C (66°F)
29°C (84°F)
18°C (64°F)
23°C (73°FJ
35°C (95°F)
24°C (75°F)
...
35°C (95°F)
34°C (93°F)
24°C (75°F)
—
22°C (72°F)
5°C (41 °F)
25°C (77°F)
9°C (48bF)
21 °C (70°F)
27°C (81°F)
21 °C (70°F)
9°C (48°F)
17°C (63°F)
10°C (50°F)
1t°C (52°F)
34°C (93°F)
13°C (55°F)
33°C (91 °F)
29°C (84°F) c
27°C (81°F) c
13°C (55°F)
23°C (73°F) c
13°C (55°F)
a Optimum or mean of the range of spawning temperatures reported for the species
b Upper temperature for successful incubation and hatching reported for the species
0 Upper temperature for spawning ' (Brungsand Jones 1977)
Maximum
weekly average
temperatures
for growth and
short-term
maximum
temperatures
for selected
fish (°C and °F)
Be sure to let someone know where you
are going and when you expect to return
TASK 2 I Measure the temperature
In general, sample away from the
streambank in the main current. The outside
curve of the stream is often a good place to
sample since the main current tends to hug
this bank. In shallow stretches, wade into
the center current carefully to measure
temperature. If wading is not possible, tape
your thermometer to an extension pole or
use a boat. Reach out from the shore or boat
as far as safely possible. If you use an
extension pole, read the temperature
quickly before it changes to the air tem-
perature.
If you are doing a horizontal or vertical
temperature profile, make sure you can
safely reach all the points where a measure-
ment is required before trying.
Measure temperature as follows:
1. Place the thermometer or meter
probe in the water as least 4 inches
below the surface or halfway to the
bottom if in a shallow stream.
2. If using a thermometer, allow
enough time for it to reach a stable
temperature (at least 1 minute). If
using a meter, allow the temperature
reading to stabilize at a constant
temperature reading.
3. If possible, try to read the tempera-
ture with the thermometer bulb
beneath the water surface. If it is not
possible, quickly remove the ther-
mometer and read the temperature.
4. Record the temperature on the field
data sheet.
TASK 3
Return the field data sheets
to the lab/dropoff point.
References
Brangs, W.S. and B.R. Jones. 1977.
Temperature Criteria for Freshwater
Fish: Protocols and Procedures. EPA-
600/3-77-061. Environ. Research Lab,
Ecological Resources Service, U.S.
Environmental Protection Agency,
Office of Research and Development,
Duluth, MN.
-------
150 I WATER QUALITY CONDITIONS
5.4
PH
What Is pH and why is it
important?
pH is a term used to indicate the
alkalinity or acidity of a substance as
ranked on a scale from 1.0 to 14.0. Acidity
increases as the pH gets lower. Fig. 5.9
present the pH of some common liquids.
pH affects many chemical and biologi-
cal processes in the water. For example,
different organisms flourish within different
ranges of pH. The largest variety of aquatic
animals prefer a range of 6.5-8.0. pH
outside this range reduces the diversity in
the stream because it stresses the physi-
ological systems of most organisms and can
reduce reproduction. Low pH can also
allow toxic elements and compounds to
become mobile and "available" for uptake
by aquatic plants and animals. This can
produce conditions that are toxic to aquatic
life, particularly to sensitive species like
rainbow trout. Changes in acidity can be
caused by atmospheric deposition (acid
rain), surrounding rock, and certain waste-
water discharges.
The pH scale measures the logarithmic
concentration of hydrogen (H+) and hy-
droxide (OH") ions, which make up water
(H+ + OH' = HjO). When both types of
ions are in equal concentration, the pH is
7.0 or neutral. Below 7.0, the water is
acidic (there are more hydrogen ions than
hydroxide ions). When the pH is above 7.0,
the water is alkaline, or basic (there are
more hydroxide ions than hydrogen ions).
Since the scale is logarithmic, a drop in the
pH by 1.0 unit is equivalent to a 10-fold
increase in acidity. So, a water sample with
a pH of 5.0 is 10 times as acidic as one with
a pH of 6.0, and pH 4.0 is 100 times as
acidic as pH 6.0.
Analytical and equipment
considerations
pH can be analyzed in the field or in the
lab. If it is analyzed in the lab, you must
measure the pH within 2 hours of the
sample collection. This is because the pH
will change due to the carbon dioxide from
the air dissolving in the water, which will
bring the pH toward 7.
If your program requires a high degree
of accuracy and precision in pH results, the
pH should be measured with a laboratory
quality pH meter and electrode. Meters of
this quality range in cost from around $250
to $1,000. Color comparators and pH
Figure 5.9
pH of selected
liquids
1M
Hcl
NEUTRAL
I I I I I I I
I I
9 10
I I
11
12
I I I
gastric
juices
oranges I
tomatoes
urine pure
water
blood
I I
seawater
household
ammonia
I I
13 14
I I I I i I I I I I I I I \ I
1M
NaOH
-------
WATER QUALITY CONDITIONS I 151
"pocket pals" are suitable for most other
purposes. The cost of either of these is in
the $50 range. The lower cost of the
alternatives might be attractive if you are
relying on several teams of volunteers
sampling multiple sites at the same time.
pH Meters
A pH meter measures the electric
potential (millivolts) across an electrode
when immersed in water. This electric
potential is a function of the hydrogen ion
activity in the sample. Therefore, pH meters
can display results in either millivolts (mV)
or pH units.
A pH meter consists of a. potentiometer,
which measures electric current; a glass
electrode, which senses the electric poten-
tial where it meets the water sample; a
reference electrode, which provides a
constant electric potential; and a tempera-
ture compensating device, which adjusts the
readings according to the temperature of the
sample (since pH varies with temperature).
The reference and glass electrodes are
frequently combined into a single probe
called a combination electrode.
There is a wide variety of meters, but
the most important part of the pH meter is
the electrode. Buy a good, reliable electrode
and follow the manufacturer's instructions
for proper maintenance. Infrequently used
or improperly maintained electrodes are
subject to corrosion, which makes them
highly inaccurate.
pH "Pocket Pals" and Color Comparators
pH "pocket pals" are electronic hand-
held "pens" that are dipped in the water and
provide a digital readout of the pH. They
can be calibrated to one pH buffer (lab
meters, on the other hand, can be calibrated
to two or more buffer solutions and thus are
more accurate over a wide range of pH
measurements).
Color comparators involve adding a
reagent to the sample that colors the sample
water. The intensity of the color is propor-
tional to the pH of the sample. This color is
then matched against a standard color
chart. The color chart equates particular
colors to associated pH values. The pH can
be determined by matching the colors from
the chart to the color of the sample.
How to collect and analyze
samples
The field procedures for collecting and
analyzing samples for pH consist of the
following tasks.
TASK 1 I Prepare the sample contain-
ers
Sample containers (and all glassware
used in this procedure) must be cleaned and
rinsed before the first run and after each
sampling run by following the procedure
described under Method A on page 128.
Remember to wear latex gloves.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, picking
up and checking supplies, and checking
weather and directions. In addition to the
standard sampling equipment and apparel,
when sampling for pH, include the follow-
ing equipment:
• pH meter with combination tempera-
ture and reference electrode, or pH
"pocket pal" or color comparator
• Wash bottle with deionized water to
rinse pH meter electrode (if appro-
priate)
• Data sheet for pH to record results
Before you leave for the sampling site,
be sure to calibrate the pH meter or "pocket
pal." The pH meter and "pocket pal"
should be calibrated prior to sample
analysis and after every 25 samples accord-
ing to the instructions that come with them.
-------
152 I WATER QUALITY CONDITIONS
If you are using a "pocket pal," use the
buffer recommended by the manufacturer.
If you are using a laboratory grade meter,
use two pH standard buffer solutions: 4.01
and 7.0. (Buffers can be purchased from
test kit supply companies, such as Hach or
LaMotte.) Following are notes regarding
buffers.
• The buffer solutions should be at
room temperature when you calibrate
the meter.
• Do not use a buffer after its expira-
tion date.
• Always cap the buffers during
storage to prevent contamination.
• Because buffer pH values change
with temperature, the meter must
have a built-in temperature sensor
that automatically standardizes the
pH when the meter is calibrated.
• Do not reuse buffer solutions!
TASK 3 I Collect the sample
Refer to page 128 for details on how to
collect water samples using screw-cap
bottles or Whirl-pak® bags.
TASK 4 I Measure pH
The procedure for measuring pH is the
same whether it is conducted in the field or
lab.
If you are using a "pocket pal" or color
comparator, follow the manufacturer's
instructions. Use the following steps to
determine the pH of your sample if you are
using a meter.
1. Rinse the electrode well with deion-
ized water.
2. Place the pH meter or electrode into
the sample. Depress the dispenser
button once to dispense electrolyte.
Read and record the temperature and
pH in the appropriate column on the
data sheet. Rinse the electrode well
with deionized water.
3. Measure the pH of the 4.01 and 7.0
buffers periodically to ensure that the
meter is not drifting off calibration.
If it has drifted, recalibrate it.
TASK 4
Return the field data sheets
and samples to the lab or
drop-off point.
Samples for pH must be analyzed
within 2 hours of collection. If the samples
cannot be analyzed in the field, keep the
samples on ice and take them to the lab or
drop-off point as soon as possible within
the 2-hour limit.
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
River Watch Network. 1992. Total alkalin-
ity and pH field and laboratory proce-
dures (based on University of Massachu-
setts Acid Rain Monitoring Project). July
1.
-------
WATER QUALITY CONDITIONS I 153
5.5
Turbidity
What is turbidity and why is it
important?
Turbidity is a measure of water clar-
ity—how much the material suspended in
water decreases the passage of light through
the water. Suspended materials include soil
particles (clay, silt, and sand), algae,
plankton, microbes, and other substances.
These materials are typically in the size
range of 0.004 mm (clay) to 1.0 mm (sand).
Turbidity can affect the color of the water.
Higher turbidity increases water
temperatures because suspended particles
absorb more heat. This, in turn, reduces the
concentration of dissolved oxygen (DO)
because warm water holds less DO than
cold. Higher turbidity also reduces the
amount of light penetrating the water,
which reduces photosynthesis and the
production of DO. Suspended materials can
clog fish gills, reducing resistance to
disease in fish, lowering growth rates, and
affecting egg and larval development. As
the particles settle, they can blanket the
stream bottom, especially in slower waters,
and smother fish eggs and benthic macroin-
vertebrates. Sources of turbidity include:
• Soil erosion
• Waste discharge
• Urban runoff
• Eroding stream banks
• Large numbers of bottom feeders
(such as carp), which stir up bottom
sediments
• Excessive algal growth.
Sampling and equipment
considerations
Turbidity can be useful as an indicator
of the effects of runoff from construction,
agricultural practices, logging activity,
discharges, and other sources. Turbidity
often increases sharply during a rainfall,
especially in developed watersheds, which
typically have relatively high proportions
of impervious surfaces. The flow of
stormwater runoff from impervious sur-
faces rapidly increases stream velocity,
which increases the erosion rates of
streambanks and channels. Turbidity can
also rise sharply during dry weather if
earth-disturbing activities are occurring in
or near a stream without erosion control
practices in place.
Regular monitoring of turbidity can
help detect trends that might indicate
increasing erosion in developing water-
sheds. However, turbidity is closely related
to stream flow and velocity and should be
correlated with these factors. Comparisons
of the change in turbidity over tjme,
therefore, should be made at the same point
at the same flow.
Turbidity is not a measurement of the
amount of suspended solids present or the
rate of sedimentation of a steam since it
measures only the amount of light that is
scattered by suspended particles. Measure-
ment of total solids is a more direct mea-
sure of the amount of material suspended
and dissolved in water (see section 5.9).
Turbidity is generally measured by
using a turbidity meter. Volunteer pro-
grams may also take samples to a lab for
analysis. Another approach is to measure
transparency (an integrated measure of
light scattering and absorption) instead of
turbidity. Water clarity/transparency can be
measured using a Secchi disk or transpar-
ency tube. The Secchi disk can only be
used in deep, slow moving rivers; the
transparency tube, a comparatively new
development, is gaining acceptance in
-------
154 I WATER QUALITY CONDITIONS
Figure 5.10
Using a
Secchi disk to
measure
transparency.
The disk is
lowered until it
is no longer
visible. That
point is the
Secchi disk
depth.
Meters
Figure 5.11
Using a
transparency
tube.
(A) Prepare the
transparency
tube to take a
reading. Place
the tube on a
white surface
and look
vertically down
the tube to see
the wave
pattern at the
bottom.
(B) Slowly pour
water sample
into the tube
stopping
intermittently to
see if the wave
pattern has
disappeared.
Marked in
tenth of
a meter
Increments
Meters
Meter
programs around the country but is not yet
in wide use (see Using a Secchi Disk or
Tranparency Tube).
A turbidity meter consists of a light
source that illuminates a water sample and
a photoelectric cell that measures the
intensity of light scattered at a 90° angle by
the particles in the sample. It measures
turbidity in nephelometric turbidity units or
NTUs. Meters can measure turbidity over a
wide range—from 0 to 1000 NTUs. A clear
mountain stream might have a turbidity of
around 1 NTU, whereas a large river like
the Mississippi might have a dry-weather
turbidity of around 10 NTUs. These values
can jump into hundreds of NTU during
runoff events. Therefore, the turbidity
meter to be used should be reliable over the
range in which you will be working. Meters
of this quality cost about $800. Many
meters in this price range are designed for
field or lab use.
Although turbidity meters can be used
in the field, volunteers might want to
collect samples and take them to a central
B
-------
WATER QUALITY CONDITIONS I 155
Using a Secchi Disk or Transparency Tube
SecchiDisk
A Secchi disk is a black and white disk that is lowered by hand into the water to the depth at which it vanishes from
sight (Figure 5.10). The distance to vanishing is then recorded. The clearer the water, the greater the distance. Secchi
disks are simple to use and inexpensive. For river monitoring they have limited use, however, because in most cases
the river bottom will be visible and the disk will not reach a vanishing, point. Deeper, slower moving rivers are the most
appropriate places for Secchi disk measurement although the current might require that the disk be extra-weighted so it
does not sway and make measurement difficult. ;Secchi disks cost about $50 and can be homemade.
The line attached to the Secchi disk must be marked according to units designated by the volunteer program, in
waterproof ink. Many programs require volunteers to measure to the nearest 1/10 meter. Meter intervals can be tagged
(e;g., with duct tape) for ease of use.
To measure water clarity with a Secchi disk: ,
• Check to make sure that the Secchi disk is securely attached to the measured line.
• Lean over the side of the boat and lower the Secchi disk into the water, keeping your back toward the sun to
block glare.
• Lower the disk until it disappears from view. Lower it one third of a meter and then slowly raise the disk until it
just reappears. Move the disk up and down until the exact vanishing point is found.
• Attach a clothespin to the line at the point where the line enters the water. Record the measurement on your data
sheet. Repeating the measurement will provide you with a quality control check.
The key to consistent results is to train volunteers to follow standard sampling procedures and, if possible, have the
same individual take the reading at the same site throughout the season.
Transparency Tube
Pioneered by Australia's Department of Conservation, the transparency tube is a clear, narrow plastic tube marked
in units with a dark pattern painted on the bottom. Water is poured into the tube until the pattern disappears (Figure
5.11). Some U.S. volunteer monitoring programs (e.g., the Tennessee Valley Authority (TVA) Clean Water Initiative and
the Minnesota Pollution Control Agency (MPCA)) are testing the transparency tube in streams and rivers. MPCA uses
tubes marked in centimeters, and has found tube readings to relate fairly well to lab measurements of turbidity and total
suspended solids (although they do not recommend the transparency tube for applications where precise and accurate
measurement is required or in highly colored waters).
The TVA and MPCA recommend the following sampling considerations:
• Collect the sample in a bottle or bucket in mid-stream and mid-depth if possible. Avoid stagnant water and
sample as far from the shoreline as is safe. Avoid collecting sediment from the bottom of the stream.
• Face upstream as you fill the bottle or bucket.
• Take readings in open but shaded conditions. Avoid direct sunlight by turning your back to the sun.
• Carefully stir or swish the water in the bucket or bottle until it is homogeneous, taking care not to produce air
bubbles (these will scatter light and affect the measurement). Then pour the water slowly in the tube while looking
down the tube. Measure the depth of the water column in the tube when the symbol just disappears.
For more information on using a transparency tube, see the references at the end of this section. Many programs
have begun making their own tubes. They now may also be purchased in the U.S. (see Appendix B—Scientific Supply
Houses).
-------
156 I WATER QUALITY CONDITIONS
point for turbidity measurements. This is
because pf the expense of the meter .(most
programs can afford only one and would
have to pass it along from site to site,
complicating logistics and increasing the
risk of damage to the meter) and because
the meter includes glass cells that must
remain optically clear and free of scratches.
Volunteers can also take turbidity
samples to a lab for meter analysis at a
reasonable cost.
How to sample
The procedures for collecting samples
and analyzing turbidity consist of the
following tasks:
TASK 3 I Collect the sample
TASK1
Prepare the sample contain-
ers
If factory-sealed, disposable Whirl-
pak® bags are used to sample, no prepara-
tion is needed. Reused sample containers
(and all glassware used in this procedure)
must be cleaned before the first run and
after each sampling run by following
Method A described on page 128.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
consideration, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, when sampling for turbidity,
include the following equipment:
• Turbidity meter
• Turbidity standards
• Lint-free cloth to wipe the cells of
the meter
• Data sheet for turbidity to record
results
Be sure to let someone know where you
are going and when you expect to return.
Refer to page 128 for details on how to
collect water samples using screw-cap
bottles or Whirkpak® bags.
TASK 4 I Analyze the sample
The following procedure applies to
field or lab use of the turbidity meter.
1. Prepare the turbidity meter for use
according to the manufacturer's
directions.
2. Use the turbidity standards provided
with the meter to calibrate it. Make
sure it is, reading accurately in the
range in which you will be working.
3. Shake the sample vigorously and
wait until the bubbles have disap-
peared. You might want to tap the
sides of the bottle gently to acceler-
ate the process.
4. Use a lint-free cloth to wipe the
outside of the tube into which the
sample will be poured. Be sure not to
handle the tube below the line where
the light will pass when the tube is
placed in the meter.
5. Pour the sample water into the tube.
Wipe off any drops on the outside of
the tube.
6. Set the meter for the appropriate
turbidity range. Place the tube in the
meter and read the turbidity measure-
ment directly from the meter display.
7. Record the result on the field or lab
sheet.
8. Repeat steps 3-7 for each sample.
TASKS
Return the samples and the
field data sheets to the lab/
drop-off point.
If you are sending your samples to a lab
for analysis, they must be tested within 24
hours of collection. Keep samples in the
dark and on ice or refrigerated.
-------
WATER QUALITY CONDITIONS I 157
References and Further Reading
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
Minnesota Pollution Control Agency. 1997.
An Attempt to Classify Transparency
Tube Readings for Southern Minnesota,
by Lee Ganske. Contact Louise Hotka,
MPCA, Tel: (612) 296-7223, E-mail:
louise.hotka@pca.state.mn.us.
Mississippi Headwaters River Watch. 1991.
Water quality procedures. Mississippi
Headwaters Board. March.
Mitchell, M.K., and W. Stapp. Field
manual for water quality monitoring. 5th
ed. Thompson Shore Printers.
Tennessee Valley Authority (TVA). 1995
(draft). Clean Water Initiative Volunteer
Stream Monitoring Methods Manual.
TVA, 1101 Market Street, CST 17D,
Chattanooga, TN 37402-2801
USEPA. 1991. Volunteer lake monitoring:
A methods manual. EPA 440/4-91-002.
Office of Water, U. S. Environmental
Protection Agency, Washington, DC.
White, T. 1994. Monitoring a watershed:
Nationwide turbidity testing in Australia.
Volunteer Monitor. 6(2):22-23.
-------
158 I WATER QUALITY CONDITIONS
5.6
Phosphorus
Why is phosphorus important?
Both phosphorus and nitrogen are
essential nutrients for the plants and
animals that make up the aquatic food web.
Since phosphorus is the nutrient in short
supply in most fresh waters, even a modest
increase in phosphorus can, under the right
conditions, set off a whole chain of undesir-
able events in a stream including acceler-
ated plant growth, algae blooms, low
dissolved oxygen, and the death of certain
fish, invertebrates, and other aquatic
animals.
There are many sources of phosphorus,
both natural and human. These include soil
and rocks, wastewater treatment plants,
runoff from fertilized lawns and cropland,
failing septic systems, runoff from animal
manure storage areas, disturbed land areas,
drained wetlands, water treatment, and
commercial cleaning preparations.
Forms of phosphorus
Phosphorus has a complicated story.
Pure, "elemental" phosphorus (P) is rare. In
nature, phosphorus usually exists as part of
a phosphate molecule (PO4). Phosphorus in
aquatic systems occurs as organic phos-
phate and inorganic phosphate. Organic
phosphate consists of a phosphate molecule
associated with a carbon-based molecule, as
in plant or animal tissue. Phosphate that is
not associated with organic material is
inorganic. Inorganic phosphorus is the form
required by plants. Animals can use either
organic or inorganic phosphate.
Both organic and inorganic phosphorus
can either be dissolved in the water or
suspended (attached to particles in the
water column).
The phosphorus cycle
Phosphorus cycles through the environ-
ment, changing form as it does so (Fig.
5.12). Aquatic plants take in dissolved
inorganic phosphorus and convert it to
organic phosphorus as it becomes part of
their tissues. Animals get the organic
phosphorus they need by eating either
aquatic plants, other animals, or decompos-
ing plant and animal material.
Figure 5.12
The phospho-
rus cycle
Phosphorus
changes form
as it cycles
through the
aquatic environ-
ment.
THE PHOSPHORUS CYCLE
Inorganic phosphorus C==c> Intake by plants £={? Grazing and predation by animals
(from various natural and human sources) (converted to organic P) (organic P)
Inorganic P
returned to
water column
Death
I
Death
Excretion
Decomposition
(organic P converted to inorganic P by bacterial action)
-------
WATER QUALITY CONDITIONS I 159
As plants and animals excrete wastes or
die, the organic phosphorus they contain
sinks to the bottom, where bacterial decom-
position converts it back to inorganic
phosphorus, both dissolved and attached to
particles. This inorganic phosphorus gets
back into the water column when the
bottom is stirred up by animals, human
activity, chemical interactions, or water
currents. Then it is taken up by plants and
the cycle begins again.
In a stream system, the phosphorus
cycle tends to move phosphorus down-
stream as the current carries decomposing
plant and animal tissue and dissolved
phosphorus. It becomes stationary only
when it is taken up by plants or is bound to
particles that settle to the bottom of pools.
In the field of water quality chemistry,
phosphorus is described using several
terms. Some of these terms are chemistry
based (referring to chemically based
compounds), and others are methods-based
(they describe what is measured by a
particular method).
The term "orthophosphate" is a chemis-
try-based term that refers to the phosphate
molecule all by itself. "Reactive phospho-
rus" is a corresponding method-based term
that describes what you are actually mea-
suring when you perform the test for
orthophosphate. Because the lab procedure
isn't quite perfect, you get mostly ortho-
phosphate but you also get a small fraction
of some other forms.
More complex inorganic phosphate
compounds are referred to as "condensed
phosphates" or "polyphosphates." The
method-based term for these forms is "acid
hydrolyzable."
Monitoring phosphorus
Monitoring phosphorus is challenging
because it involves measuring very low
concentrations-^-down to 0.01 milligram
per liter (mg/L) or even lower. Even such
very low concentrations of phosphorus can
have a dramatic impact on streams. Less
sensitive methods should be used only to
identify serious problem areas.
While there are many tests for phos-
phorus, only four are likely to be per-
formed by volunteer monitors.
1. The total orthophosphdte test is
largely a measure of orthophosphate.
Because the sample is riot filtered,
the procedure measures both dis-
solved and suspended
orthophosphate. The EPA-approved
method for measuring total ortho-
phosphate is known as the ascorbic
acid method. Briefly, a reagent
(either liquid or powder) containing
ascorbic acid and ammonium
molybdate reacts with orthophos-
phate in the sample to form a blue
compound. The intensity of the blue
color is directly proportional to the
amount of orthophosphate in the
water.
2. The total phosphorus test measures
all the forms of phosphorus in the
sample (orthophosphate, condensed
phosphate, and organic phosphate).
This is accomplished by first "di-
gesting" (heating and acidifying) the
sample to convert all the other forms
to orthophosphate. Then the ortho-
phosphate is measured by the
ascorbic acid method. Because the
sample is not filtered, the procedure
measures both dissolved and sus-
pended orthophosphate.
3. The dissolved phosphorus test
measures that fraction of the total
phosphorus which is in isolution in
the water (as opposed to being
attached to suspended particles). It is
determined by first filtering the
sample, then analyzing the filtered
1 sample for total phosphbrus.
4. Insoluble phosphorus is calculated
by subtracting the dissolved phos-
phorus result from the total
phosphorus result.
-------
160 I WATER QUALITY CONDITIONS
All these tests have one thing in
common—they all depend on measuring
orthophosphate. The total orthophosphate
test measures the orthophosphate that is
already present in the sample. The others
measure that which is already present and
that which is formed when the other forms
of phosphorus are converted to orthophos-
phate by digestion.
Sampling and equipment
considerations
Monitoring phosphorus involves two
basic steps:
• Collecting a water sample
• Analyzing it in the field or lab for
one of the types of phosphorus
described above.
This manual does not address labora-
tory methods. Refer to the references cited
at the end of this section.
Sample Containers
Sample containers made of either some
form of plastic or Pyrex® glass are accept-
able to EPA. Because phosphorus mol-
ecules have a tendency to "adsorb" (attach)
to the inside surface of sample containers, if
containers are to be reused they must be
acid-washed to remove adsorbed phospho-
rus. Therefore, the container must be able
to withstand repeated contact with hydro-
chloric acid. Plastic containers—either
high-density polyethylene or polypropy-
lene—might be preferable to glass from a
practical standpoint because they will better
withstand breakage. Some programs use
disposable, sterile, plastic Whirl-pak®
bags. The size of the container will depend
on the sample amount needed for the
phosphorus analysis method you choose
and the amount needed for other analyses
you intend to perform.
Dedicated Labware
All containers that will hold water
samples or come into contact with reagents
used in this test must be dedicated. That is,
they should not be used for other tests. This
is to eliminate the possibility that reagents
containing phosphorus will contaminate the
labware. All labware should be acid-
washed.
The only form of phosphorus this
manual recommends for field analysis is
total orthophosphate, which uses the
ascorbic acid method on an untreated
sample. Analysis of any of the other forms
requires adding potentially hazardous
reagents, heating the sample to boiling, and
using too much time and too much equip-
ment to be practical. In addition, analysis
for other forms of phosphorus is prone to
errors and inaccuracies in a field situation.
Pretreatment and analysis for these other
forms should be handled in a laboratory.
Ascorbic Acid Method
In the ascorbic acid method, a com-
bined liquid or prepackaged powder
reagent, consisting of sulfuric acid, potas-
sium antimonyl tartrate, ammonium molyb-
date, and ascorbic acid (or comparable
compounds), is added to either 50 or 25 mL
of the water sample. This colors the sample
blue in direct proportion to the amount of
orthophosphate in the sample. Absorbance
or transmittance is then measured after 10
minutes, but before 30 minutes, using a
color comparator with a scale in milligrams
per liter that increases with the increase in
color hue, or an electronic meter that
measures the amount of light absorbed or
transmitted at a wavelength of 700-880
nanometers (again depending on
manufacturer's directions).
A color comparator may be useful for
identifying heavily polluted sites with high
concentrations (greater than 0.1 mg/L).
However, matching the color of a treated
sample to a comparator can be very subjec-
-------
WATER QUALITY CONDITIONS I 161
tive, especially at low concentrations, and
can lead to variable results.
A field spectrophotometer or colorim-
eter with a 2.5-cm light path and an infrared
photocell (set for a wavelength of 700-880
nm) is recommended for accurate determi-
nation of low concentrations (between 0.2
and 0.02 mg/L). Use of a meter requires
that you prepare and analyze known
standard concentrations ahead of time in
order to convert the absorbance readings of
your stream sample to milligrams per liter,
or that your meter reads directly as milli-
grams per liter.
How to prepare standard
concentrations
Note that this step is best accomplished
in the lab before leaving for sampling.
Standards are prepared using a phosphate
standard solution of 3 mg/L as phosphate
(PO4). This is equivalent to a concentration
of 1 mg/L as Phosphorus (P). All references
to concentrations and results from this point
on in this procedure will be expressed as
mg/L as P, since this is the convention for
reporting results.
Six standard concentrations will be
prepared for every sampling date in the
range of expected results. For most
samples, the following six concentrations
should be adequate:
standard solution to each 25-mL
volumetric flask as follows:
0.00 mg/L
0.04 mg/L
0.08 mg/L
Proceed as follows:
0.12 mg/L
0.16 mg/L
0.20 mg/L
1. Set out six 25-mL volumetric
flasks—one for each standard. Label
the flasks 0.00, 0.04, 0.08, 0.12, 0.16,
. and 0.20.
2. Pour about 30 mL of the phosphate
standard solution into a 50 mL
beaker.
3. Use 1-, 2-, 3-, 4-, and 5-mL Glass A
volumetric pipets to transfer corre-
sponding volumes of phosphate
Standard
Concentration
0.00
'0.04
0.08
0.12
0.16
0.20
mL of Phosphate
Standard Solution
0
1
2
3
4
5
Note: The standard solution is calculated
based on the equation: A = (B x C) -*• D
Where:
A = mL of standard solution needed
B = desired concentration of standard
C = final volume (mL) of standard
D = concentration of standard solution
For example, to find out how much
phosphate standard solution to use to
make a 0.04-mg/L standard: •
A = (0.04 x 25) -*-1
A = 1 mL
Before transferring the solution, clear
each pipet by filling it once with the
standard solution and blowing it out. Rinse
each pipet with deionized water after use.
4. Fill the remainder of each 25 mL
volumetric flask with distilled,
deionized water to the 25 mL line.
Swirl to mix.
5. Set out and label six 50-mL Erlenm-
eyer flasks: 0.00, 0.04,0.08, 0.12,
0.16, and 0.20. Pour the standards
from the volumetric flasks, to the
Erlenmeyer flasks.
6. List the standard concentrations
(0.00, 0.04, 0.08, 0.12, 0.16, and
0.20) under "Bottle #" on the lab
sheet.
7. Analyze each of these standard
concentrations as described in the
section below.
-------
162 I WATER QUALITY CONDITIONS
How to collect and analyze
samples
The field procedures for collecting and
analyzing samples for phosphorus consist
of the following tasks:
TASK 3 | Collect the sample
TASK 1 I Prepare the sample contain-
m~mm>mmm1'^ ers
If factory-sealed, disposable Whirl-
pak® bags are used for sampling, no
preparation is needed. Reused sample
containers (and all glassware used in this
procedure) must be cleaned (including acid
rinse) before the first run and after each
sampling run by following the procedure
described in Method B on page 128.
Remember to wear latex gloves.
TASK 2
Prepare before leaving for
the sample site
Refer to page 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to sample containers and the standard
sampling apparel, you will need the follow-
ing equipment and supplies for total
reactive phosphorus analysis:
• Color comparator or field spectro-
photometer with sample tubes for
reading the absorbance of the sample
• Prepackaged reagents (combined
reagents) to turn the water blue
• Deionized or distilled water to rinse
the sample tubes between uses
• Wash bottle to hold rinse water
• Mixing container with a mark at the
recommended sample volume
(usually 25 mL) to hold and mix the
sample
• Clean, lint-free wipes to clean and
dry the sample tubes
Note that prepackaged reagents are
recommended for ease and safety.
Refer to page 128 for details on how to
collect water samples using screw-cap
bottles or Whirl-pak® bags.
TASK 4
Analyze the sample in the
field (for total orthophos-
phate only) using the
ascorbic acid method.
If using an electronic spectrophotometer or
colorimeter:
1. "Zero" the meter (if you are using
one) using a reagent blank (distilled
water plus the reagent powder) and
following the manufacturer's direc-
tions.
2. Pour the recommended sample
volume (usually 25 mL) into a
mixing container and add reagent
powder pillows. Swirl to mix. Wait
the recommended time (usually at
least 10 minutes) before proceeding.
3. Pour the first field sample into the
sample cell test tube. Wipe the tube
with a lint-free cloth to be sure it is
clean and free of smudges or water
droplets. Insert the tube into the
sample cell.
4. Record the bottle number on the field
data sheet.
5. Place the cover over the sample cell.
Read the absorbance or concentration
of this sample and record it on the
field data sheet.
6. Pour the sample back into its flask.
7. Rinse the sample cell test tube and
mixing container three times with
distilled, deionized water. Avoid
touching the lower portion of the
sample cell test tube. Wipe with a
clean, lint-free wipe. Be sure that the
lower part of the sample cell test tube
is clean and free of smudges or water
droplets.
-------
WATER QUALITY CONDITIONS I 163
Be sure to use the same sample
cell test tube for each sample. If the
test tube breaks, use a new one and
repeat step 1 to "zero" the meter.
If using a color comparator:
1. Follow the manufacturer's directions.
Be sure to pay attention to the
direction of your light source when
reading the color development. The
light source should be in the same
position relative to the color com-
parator for each sample. Otherwise,
this is a source of significant error.
As a quality check, have someone
else read the comparator after you.
2. Record the concentration on the field
data sheet.
TASKS I Return the samples (for lab
^—«^—J analysis for other tests) and
the field data sheets to the
lab/drop-off point.
Samples for different types of phospho-
rus must be analyzed within a certain time
period. For some types of phosphorus, this
is a matter of hours; for others, samples can
be preserved and held for longer periods.
Samples being tested for orthophosphate
must be analyzed within 48 hours of
collection. In any case, keep the samples
on ice and take them to the lab or drop-off
point as soon as possible.
TASK 6
Analyze the samples in the
lab.
Lab methods for other tests are de-
scribed in the references below (APHA.
1992; Hach Company, 1992; River Watch
Network, 1992; USEPA, 1983).
TASK?
Report the results and
convert to milligrams per
liter
First, absorbance values must be
converted to milligrams per liter. This is
done by constructing a "standard curve"
using the absorbance results from your
standard concentrations.
1. Make an absorbance versus concen-
tration graph on graph paper:
• Make the "y" (vertical) axis and
label it "absorbance." Mark this
axis in 0.05 increments from 0 as
high as the graph paper will allow.
• Make the "x" (horizontal) axis and
label it "concentration: mg/L as
P." Mark this axis with the
concentration of the standards: 0,
0.04,0.08,0.12, 0.16, 0.20.
2. Plot the absorbance of the standard
concentrations on the graph.
3. Draw a "best fit" straight line
through these points. The line should
touch (or almost touch) each of the
points. If it doesn't, make up new
standards and repeat the procedure.
Example: Suppose you measure the
absorbance of the six standard concentra-
tions as follows:
Concentration
0.00
0.04
0.08
0.12
0.16
0.20
Absorbance
0.000
0.039
'0.078
0.105
0.155
0.192
The resulting standard curve is dis-
played in Fig. 5.13.
4. For each sample, locate the absor-
bance on the "y" axis, read
horizontally over to the line, and
then more down to read the concen-
tration in mg/L as P.
1 5. Record the concentration on the lab
sheet in the appropriate column.
NOTE: The detection limit for this
test is 0.01 mg/L. Report any results
less than 0.01 as "<0.01." Round off
all results to the nearest hundredth of
a mg/L.
-------
164 I WATER QUALITY CONDITIONS
Figure 5.13
Absorbance of
standard
concentra-
tions, when
plotted, should
result in a
straight line
0.20
O
u
c
ra
J3
».
o
(A
ft
0.15-
0.10--
0.05-
0.00
0.00
0.08
0.12
0.16
0.20
Concentration (mg/L as P)
Results can either be reported "as P" or
"as PO4." Remember that your results are
reported as milligrams per liter—weight per
unit of volume. Since the PO4 molecule is
three times as heavy as the P atom, results
reported as PO4 are three times the concen-
tration of those reported as P. For example,
if you measure 0.06 mg/L as PO4, that's
equivalent to 0.02 mg/L as P. To convert
PO4 to P, divide by 3. To convert P to PO4,
multiply by 3, To avoid this confusion, and
since most state water quality standards are
reported as P, this manual recommends that
results always be reported as P.
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
Black, J.A. 1977. Water pollution technol-
ogy. Reston Publishing Co., Reston, VA.
Caduto, MJ. 1990. Pond and brook.
University Press of New England,
Hanover, NH.
Dates, Geoff. 1994. Monitoring for phos-
phorus or how come they don't tell you
this stuff in the manual? Volunteer
Monitor, Vol. 6(1), spring 1994.
Hach Company. 1992. Each -water analysis
handbook. 2nd ed. Loveland, CO.
River Watch Network. 1991. Total phos-
phorus test (adapted from Standard
Methods). July 17.
River Watch Network. 1992. Total phos-
phorus (persulfate digestion followed by
ascorbic acid procedure, Hach adapta-
tion of Standard Methods). July 1.
USEPA. 1983. Methods for chemical
analysis of water and wastes. 2nd ed.
Method 365.2. U.S. Environmental
Protection Agency, Washington, DC.
-------
WATER QUALITY CONDITIONS I 165
5.7
Nitrates
What are nitrates and why are
they important?
Nitrates are a form of nitrogen, which
is found in several different forms in
terrestrial and aquatic ecosystems. These
forms of nitrogen include ammonia (NH3),
nitrates (NO3), and nitrites (NO2). Nitrates
are essential plant nutrients, but in excess
amounts they can cause significant water
quality problems. Together with phospho-
rus, nitrates in excess amounts can acceler-
ate eutrophication, causing dramatic
increases in aquatic plant growth and
changes in the types of plants and animals
that live in the stream. This, in turn, affects
dissolved oxygen, temperature, and other
indicators. Excess nitrates can cause
hypoxia (low levels of dissolved oxygen)
and can become toxic to warm-blooded
animals at higher concentrations (10 mg/L)
or higher) under certain conditions. The
natural level of ammonia or nitrate in
surface water is typically low (less than 1
mg/L); in the effluent of waste water
treatment plants, it can range up to
30 mg/L.
Sources of nitrates include wastewater
treatment plants, runoff from fertilized
lawns and cropland, failing on-site septic
systems, runoff from animal manure
storage areas, and industrial discharges that
contain corrosion inhibitors.
Sampling and equipment
considerations
Nitrates from land sources end up in
rivers and streams more quickly than other
nutrients like phosphorus. This is because
they dissolve in water more readily than
phosphates, which have an attraction for
soil particles. As a result, nitrates serve as a
better indicator of the possibility of a
source of sewage or manure pollution
during dry weather.
Water that is polluted with nitrogen-
rich organic matter might show low
nitrates. Decomposition of the organic
matter lowers the dissolved oxygen level,
which in turn slows the rate at which
ammonia is oxidized to nitrite (NO2) and
then to nitrate (NO3). Under such circum-
stances, it might be necessary to also
monitor for nitrites or ammonia, which are
considerably more toxic to aquatic life than
nitrate. (See Standard Methods section
4500-NH3and 4500-NO2 for appropriate
nitrite methods; APHA, 1992):
Water samples to be tested for nitrate
should be collected in glass or polyethylene
containers that have been prepared by
using Method B in the introduction.
Volunteer monitoring programs usually
use two methods for nitrate testing: the
cadmium reduction method and the nitrate
electrode. The more commonly used
cadmium reduction method produces a
color reaction that is then measured either
by comparison to a color wheel or by use
of a spectrophotometer. A few programs
also use a nitrate electrode, which can
measure in the range of 0 to 100 mg/L
nitrate. A newer colorimetric irnmunoassay
technique for nitrate screening is also now
available and might be applicable for
volunteers.
Cadmium Reduction Method ,
The cadmium reduction method is a
colorimetric method that involves contact
of the nitrate in the sample with cadmium
particles, which cause nitrates to be con-
verted to nitrites. The nitrites then react
with another reagent to form a red color
whose intensity is proportional to the
original amount of nitrate. The red color is
then measured either by comparison to a
color wheel with a scale in milligrams per
liter that increases with the increase in
-------
166 I WATER QUALITY CONDITIONS
color hue, or by use of an electronic spec-
trophotometer that measures the amount of
light absorbed by the treated sample at a
543-nanometer wavelength. The absor-
bance value is then converted to the equiva-
lent concentration of nitrate by using a
standard curve. Methods for making
standard solutions and standard curves are
presented at the end of this section.
This curve should be created by the
program advisor before each sampling run.
The curve is developed by making a set of
standard concentrations of nitrate, reacting
them and developing the corresponding
color, and then plotting the absorbance
value for each concentration against
concentration. A standard curve could also
be generated for the color wheel.
Use of the color wheel is appropriate
only if nitrate concentrations are greater
than 1 mg/L. For concentrations below 1
mg/L, a spectrophotometer should be used.
Matching the color of a treated sample at
low concentrations to a color wheel (or
cubes) can be very subjective and can lead
to variable results. Color comparators can,
however, be effectively used to identify
sites with high nitrates.
This method requires that the samples
being treated are clear. If a sample is turbid,
it should be filtered through a 0.45-micron
filter. Be sure to test whether the filter is
nitrate-free. If copper, iron, or other metals
are present in concentrations above several
mg/L, the reaction with the cadmium will
be slowed down and the reaction time will
have to be increased.
The reagents used for this method are
often prepackaged for different ranges,
depending on the expected concentration of
nitrate in the stream. For example, the Hach
Company provides reagents for the follow-
ing ranges: low (0 to 0.40 mg/L), medium
(0 to 4.5 mg/L), and high (0 to 30 mg/L).
You should determine the appropriate range
for the stream being monitored.
Nitrate Electrode Method
A nitrate electrode (used with a meter)
is similar in function to a dissolved oxygen
meter. It consists of a probe with a sensor
that measures nitrate activity in the water;
this activity affects the electric potential of
a solution in the probe. This change is then
transmitted to the meter, which converts the
electric signal to a scale that is read in
millivolts. The millivolts are then converted
to mg/L of nitrate by plotting them from a
standard curve (see above). The accuracy of
the electrode can be affected by high
concentrations of chloride or bicarbonate
ions in the sample water. Fluctuating pH
levels can also affect the reading by the
meter.
Nitrate electrodes and meters are
expensive compared to field kits that
employ the cadmium reduction method.
(The expense is comparable, however, if a
spectrophotometer is used rather than a
color wheel.) Meter/probe combinations
run between $700 and $1,200 including a
long cable to connect the probe to the
meter. If the program has a pH meter that
displays readings in millivolts, it can be
used with a nitrate probe and no separate
nitrate meter is needed. Results are read
directly as milligrams per liter.
Although nitrate electrodes and spec-
trophotometers can be used in the field,
they have certain disadvantages. These
devices are more fragile than the color
comparators and are therefore more at risk
of breaking in the field. They must be
carefully maintained and must be calibrated
before each sample run and, if you are
doing many tests, between samplings. This
means that samples are best tested in the
lab. Note that samples to be tested with a
nitrate electrode should be at room tem-
perature, whereas color comparators can be
used in the field with samples at any
temperature.
-------
WATER QUALITY CONDITIONS I 1B7
How to collect and analyze
samples
The procedures for collecting and
analyzing samples for nitrate consist of the
following tasks:
TASK 3 | Collect the sample
TASK 1 I Prepare the sample containers
If factory-sealed, disposable Whirl-
pak® bags are used for sampling, no
preparation is needed. Reused sample
containers (and all glassware used in this
procedure) must be cleaned before the first
run and after each sampling by following
the method described on page 128 under
Method B. Remember to wear latex gloves.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, the following equipment is
needed when analyzing nitrate nitrogen in
the field:
• Color comparator or field spectro-
photometer with sample tubes (for
, reading absorbance of the sample)
• Reagent powder pillows (reagents to
turn the water red)
• Deionized or distilled water to rinse
the sample tubes between uses
• Wash bottle to hold rinse water
• Waste bottle with secure lid to hold
used cadmium particles, which
should be clearly labeled and re-
turned to the lab, where the cadmium
will be properly disposed of
• Mixing container with a mark at the
sample volume (usually 25 mL) to
hold and mix the sample
• Clean, lint-free wipes to clean and
dry the sample tubes
Refer to page 128 for details on
collecting a sample using screw-cap bottles
or Whirl-pak® bags.
TASK 4
Analyze the sample in the
field
Cadmium Reduction Method With a
Spectrophotometer
The following is the general procedure
to analyze a sample using the cadmium
reduction method with a spectrophotom-
eter. However, this should not replace the
manufacturer's directions if they differ
from the steps provided below:
1. Pour the first field sample into the
sample cell test tube and; insert it into
the sample cell of the spectropho-
tometer.
2. Record the bottle number on the lab
sheet.
i
3. Place the cover over the sample cell.
Read the absorbance or concentra-
tion of this sample and record it on
the field data sheet. :
4. Pour the sample back into the waste
bottle for disposal at the lab.
Cadmium Reduction Method With a Color
Comparator
To analyze a sample using the cad-
mium reduction method with a color
comparator, follow the manufacturer's
directions and record the concentration on
the field data sheet.
TASK 5
Return the samples and the
field data sheets to the lab/
drop-off point for analysis
Samples being sent to a lab for analysis
must be tested for nitrates within 48 hours
of collection. Keep samples in the dark and
on ice or refrigerated.
-------
168 I WATER QUALITY CONDITIONS
TASK 6
Determine results (for
spectrophotometer absor-
bance or nitrate electrode) in
lab
Preparation of Standard
Concentrations
Cadmium Reduction Method With a Spectro-
photometer
First determine the range you will be
testing (low, medium, or high). For each
range you will need to determine the lower
end, which will be determined by the
detection limit of your spectrophotometer.
The high end of the range will be the
endpoint of the range you are using. Use a
nitrate nitrogen standard solution of appro-
priate strength for the range in which you
are working. A 1-mg/L nitrate nitrogen
(NO3-N) solution would be suitable for
low-range (0 to 1.0 mg/L) tests. A 100-mg/
L standard solution would be appropriate
for medium- and high-range tests. In the
following example, it is assumed that a set
of standards for a 0 to 5.0 mg/L range is
being prepared.
Example:
1. Set out six 25-mL volumetric flasks
(one for each standard). Label the
flasks 0.0, 1.0, 2.0, 3.0, 4.0, and 5.0.
2. Pour 30 mL of a 25-mg/L nitrate
nitrogen standard solution into a 50-
mL beaker.
3. Use 1-, 2-, 3-, 4-, and 5-mL Class A
volumetric pipets to transfer corre-
sponding volumes of nitrate nitrogen
standard solution to each 25-mL
volumetric flask as follows:
Standard mL of Nitrate Nitrogen
Solution Standard Solution
0.0 0
1.0 1
2.0 2
3.0 3
4.0 4
5.0 5
Analysis of the Cadmium Reduction Method
Standard Concentrations
Use the following procedure to analyze
the standard concentrations.
1. Add reagent powder pillows to the
nitrate nitrogen standard concentra-
tions.
2. Shake each tube vigorously for at
least 3 minutes.
3. For each tube, wait at least 10
minutes but not more than 20 min-
utes to proceed.
4. "Zero" the spectrophotometer using
the 0.0 standard concentration and
following the manufacturer's direc-
tions. Record the absorbance as "0"
in the absorbance column on the lab
sheet. Rinse the sample cell three
times with distilled water.
5. Read and record the absorbance of
the 1.0-mg/L standard concentration.
6. Rinse the sample cell test tube three
times with distilled or deionized
water. Avoid touching the lower part
of the sample cell test tube. Wipe
with a clean, lint-free wipe. Be sure
that the lower part of the sample cell
test tube is clean and free of smudges
or water droplets.
7. Repeat steps 3 and 4 for each stan-
dard.
8. Prepare a calibration curve and
convert absorbance to mg/L as
follows:
• Make an absorbance versus
concentration graph on graph
paper:
(a) Make the vertical (y) axis and
label it "absorbance." Mark this
axis in 1.0 increments from 0 as
high as the graph paper will
allow.
(b) Make the horizontal (x) axis
and label it "concentration: mg/L
as nitrate nitrogen." Mark this
-------
WATER QUALITY CONDITIONS I 169
axis with the concentrations of the
standards: 0.0,1.0, 2.0, 3.0, 4.0,
and 5.0.
• Plot the absdrbance of the standard
concentrations on the graph.
• Draw a "best fit" straight line
through these points. The line
should touch (or almost touch)
each of the points. If it doesn't, the
results of this procedure are not
valid.
• For each sample, locate the
absorbance on the "y" axis, read
over horizontally to the line, and
then move down to read the
concentration in mg/L as nitrate
nitrogen.
; • Record the concentration on the
lab sheet in the appropriate
column.
For Nitrate Electrode
Standards are prepared using nitrate
standard solutions of 100 and 10 mg/L as
nitrate nitrogen (NO3-N). All references to
concentrations and results in this procedure
will be expressed as mg/L as NO3-N. Eight
standard concentrations will be prepared:
100.0 mg/L
10.0 mg/L
1.0 mg/L
0.8 mg/L
0.40 mg/L
0.32 mg/L
0.20 mg/L
0.12 mg/L
Use the following procedure:
1. Set out eight 25-mL volumetric
flasks (one for each standard). Label
the flasks 100.0, 10.0, 1.0,0.8, 0.4,
0.32,0.2, and 0.12.
2. To make the 100.0-mg/L standard,
pour 25 mL of the 100-mg/L nitrate
standard solution into the flask
labeled 100.0.
3. To make the 10.0-mg/L standard,
pour 25 mL of the 10-mg/L nitrate
standard solution into the flask
labeled 10.0.
4. To make the 1.0-mg/L standard, use
a 10- or 5-mL pipet to measure 2.5
mL of the 10-mg/L nitrate standard
solution into the flask labeled 1.0.
Fill the flask with 22.5 mL distilled,
deionized water to the fill line. Rinse
the pipet with deionized water.
5. To make the 0.8-mg/L standard, use
a 10- or 5-mL pipet or a 2-mL
volumetric pipet to measure 2 mL of
the 10-mg/L nitrate standard solution
into the flask labeled 0.8.! Fill the
flask with about 23 mL distilled,
deionized water to the fill line. Rinse
the pipet with deionized water.
6. To make the 0.4-mg/L standard, use
a 10- or 5-mL pipet or a 1-mL
volumetric pipet to measure 1 mL of
the 10-mg/L nitrate standard solution
into the flask labeled 0.4, Fill the
flask with about 24 mL distilled,
deionized water to the fill line. Rinse
the pipet with deionized water.
7. To make the 0.32-, 0.2-, and 0.12-
mg/L standards, follow step 4 to
make a 25-mL volume of 1.0 mg/L
standard solution. Transfer this to a
beaker. Pipet the following volumes
into the appropriately labeled
volumetric flasks:
Standard mL of Nitrate Nitrogen
Solution Standard Solution
0.32 8
0.20 5
0.12 3
Fill each flask up to the fill line.
Rinse pipets with deionized water.
Analysis of the Nitrate Electrode Standard
Concentrations
Use the following procedure to analyze
the standard concentrations. i
1. List the standard concentrations
(100.0, 10.0, 1.0, 0.8, 0.4, 0.32, 0.2,
and 0.12) under "bottle #" on the lab
sheet.
-------
170 I WATER QUALITY CONDITIONS
2. Prepare a calibration curve and
convert to mg/L as follows:
• Plot absorbance or mV readings
for the 100-, 10-, and 1-mg/L
standards on semi-logarithmic
graph paper, with concentration on
the logarithmic (x) axis and the
absorbance or millivolts (mV) on
the linear (y) axis.
For the nitrate electrode curve, a
straight line with a slope of 58 + 3
mV/decade at 25°C should result.
That is, measurements of 10- and
100-mg/L standard solutions
should be no more than 58 ± 3 mV
apart.
• Plot absorbance or mV readings
for the 1.0-, 0.8-, 0.4-, 0.32-, 0.2-,
and 0.12-mg/L standards on semi-
logarithmic graph paper, with
concentration on the logarithmic
(x) axis and the millivolts (mV) on
the linear (y) axis.
For the nitrate electrode, the result
here should be a curved line since
the response of the electrode at
these low concentrations is not
linear.
• For the nitrate electrode,
recalibrate the electrodes several
times daily by checking the mV
reading of the 10-mg/L and 0.4-
mg/L standards and adjusting the
calibration control on the meter
until the reading plotted on the
calibration curve is displayed
again.
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
-------
WATER QUALITY CONDITIONS I 171
5.8
Total Solids
What are total solids and why are
they important?
Total solids are dissolved solids plus
suspended and settleable solids in water. In
stream water, dissolved solids consist of
calcium, chlorides, nitrate, phosphorus,
iron, sulfur, and other ions—particles that
will pass through a filter with pores of
around 2 microns (0.002 cm) in size.
Suspended solids include silt and clay
particles, plankton, algae, fine organic
debris, and other particulate matter. These
are particles that will not pass through a 2-
micron filter.
The concentration of total dissolved
solids affects the water balance in the cells
of aquatic organisms. An organism placed
in water with a very low level of solids,
such as distilled water, will swell up
because water will tend to move into its
cells, which have a higher concentration of
solids. An organism placed in water with a
high concentration of solids will shrink
somewhat because the water in its cells will
tend to move out. This will in turn affect
the organism's ability to maintain the
proper cell density, making it difficult to
keep its position in the water column. It
might float up or sink down to a depth to
which it is not adapted, and it might not
survive.
Higher concentrations of suspended
solids can serve as carriers of toxics, which
readily cling to suspended particles. This is
particularly a concern where pesticides are
being used on irrigated crops. Where solids
are high, pesticide concentrations may
increase well beyond those of the original
application as the irrigation water travels
down irrigation ditches. Higher levels of
solids can also clog irrigation devices and
might become so high that irrigated plant
roots will lose water rather than gain it.
A high concentration of total solids
will make drinking water unpalatable and
might have an adverse effect on people
who are not used to drinking such water.
Levels of total solids that are too high or
too low can also reduce the efficiency of
wastewater treatment plants, as well as the
operation of industrial processes that use
raw water.
Total solids also affect water clarity.
Higher solids decrease the passage of light
through water, thereby slowing photosyn-
thesis by aquatic plants. Water will heat up
more rapidly and hold more heat; this, in
turn, might adversely affect aquatic life that
has adapted to a lower temperature regime.
Sources of total solids include indus-
trial discharges, sewage, fertilizers, road
runoff, and soil erosion. Total solids are
measured in milligrams per liter (mg/L).
Sampling and equipment
considerations
Total solids are important to measure
in areas where there are discharges from
sewage treatment plants, industrial plants,
or extensive crop irrigation. In particular,
streams and rivers in arid regions where
water is scarce and evaporation; is high tend
to have higher concentrations oif solids and
are more readily affected by human intro-
duction of solids from land use activities.
Total solids measurements can be
useful as an indicator of the effects of
runoff from construction, agricultural
practices, logging activities, sewage
treatment plant discharges, and other
sources. As with turbidity, concentrations
often increase sharply during rainfall,
especially in developed watersheds. They
can also rise sharply during dry weather if
earth-disturbing activities are occurring in
or near the stream without erosion control
practices in place. Regular monitoring of
total solids can help detect trends that
-------
172 I WATER QUALITY CONDITIONS
might indicate increasing erosion in devel-
oping watersheds. Total solids are related
closely to stream flow and velocity and
should be correlated with these factors. Any
change in total solids over time should be
measured at the same site at the same flow.
Total solids are measured by weighing
the amount of solids present in a known
volume of sample. This is done by weigh-
ing a beaker, filling it with a known vol-
ume, evaporating the water in an oven and
completely drying the residue, and then
weighing the beaker with the residue. The
total solids concentration is equal to the
difference between the weight of the beaker
with the residue and the weight of the
beaker without it. Since the residue is so
light in weight, the lab will need a balance
that is sensitive to weights in the range of
0.0001 gram. Balances of this type are
called analytical or Mettler balances, and
they are expensive (around $3,000). The
technique requires that the beakers be kept
in a desiccator, which is a sealed glass
container that contains material that absorbs
moisture and ensures that the weighing is
not biased by water condensing on the
beaker. Some desiccants change color to
indicate moisture content.
The measurement of total solids cannot
be done in the field. Samples must be
collected using clean glass or plastic bottles
or Whirl-pak® bags and taken to a labora-
tory where the test can be run.
How to collect and analyze
samples
The procedures for collecting and
analyzing samples for total solids consist of
the following tasks:
TASK1
Prepare the sample contain-
ers
Factory-sealed, disposable Whirl-pak®
bags are easy to use because they need no
preparation. Reused sample containers (and
all glassware used in this procedure) must
be cleaned and rinsed before the first
sampling run and after each run by follow-
ing the procedure described in Method A
on page 128.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling information. Be sure
to let someone know where you are going
and when you expect to return.
TASK 3 \ Collect the sample
Refer to page 128 for details on how to
collect water samples using screw-cap
bottles or WhirJ-pak® bags.
TASK 4
Return samples and field
sheets to the lab/drop-off
point for analysis.
Samples that are sent to a lab for total
solids analysis must be tested within seven
days of collection. Keep the samples on ice
or refrigerated.
References
APHA. 1992. Standard methods for the
examination of water and wastewater. 18th
ed. American Public Health Association,
Washington, DC.
-------
WATER QUALITY CONDITIONS I 173
5.9
Conductivity
What is conductivity and why is it
important?
Conductivity is a measure of the ability
of water to pass an electrical current.
Conductivity in water is affected by the
presence of inorganic dissolved solids such
as chloride, nitrate, sulfate, and phosphate
anions (ions that carry a negative charge) or
sodium, magnesium, calcium, iron, and
aluminum cations (ions that carry a positive
charge). Organic compounds like oil,
phenol, alcohol, and sugar do not conduct
electrical current very well and therefore
have a low conductivity when in water.
Conductivity is also affected by tempera-
ture: the warmer the water, the higher the
conductivity. For this reason, conductivity
is reported as conductivity at 25 degrees
Celsius (25 °C).
Conductivity in streams and rivers is
affected primarily by the geology of the
area through which the water flows.
Streams that run through areas with granite
bedrock tend to have lower conductivity
because granite is composed of more inert
materials that do not ionize (dissolve into
ionic components) when washed into the
water. On the other hand, streams that run
through areas with clay soils tend to have
higher conductivity because of the presence
of materials that ionize when washed into
the water. Ground water inflows can have
the same effects depending on the bedrock
they flow through.
Discharges to streams can change the
conductivity depending on their make-up.
A failing sewage system would raise the
conductivity because of the presence of
chloride, phosphate, and nitrate; an oil spill
would lower the conductivity.
The basic unit of measurement of
conductivity is the mho or Siemens. Con-
ductivity is measured in micromhos per
centimeter (|lmhos/cm) or microsiemens
per centimeter (|4,s/cm). Distilled water has
a conductivity in the range of 0.5 to 3
(imhos/cm. The conductivity of rivers in
the United States generally ranges from 50
to 1500 (Ltmhos/cm. Studies of inland fresh
waters indicate that streams supporting
good mixed fisheries have a range between
150 and 500 p-hos/cm. Conductivity outside
this range could indicate that the water is
not suitable for certain species of fish or
macroinvertebrates. Industrial waters can
range as high as 10,000 |a.mhos/cm.
Sampling and equipment
Considerations
Conductivity is useful as a general
measure of stream water quality. Each
stream tends to have a relatively constant
range of conductivity that, once estab-
lished, can be used as a baseline for
comparison with regular conductivity
measurements. Significant changes in
conductivity could then be an indicator that
a discharge or some other source of pollu-
tion has entered a stream.
Conductivity is measured with a probe
and a meter. Voltage is applied between
two electrodes in a probe immersed in the
sample water. The drop in voltage caused
by the resistance of the water is used to
calculate the conductivity per centimeter.
The meter converts the probe measurement
to micromhos per centimeter and displays
the result for the user. NOTE: Some
conductivity meters can also be used to test
for total dissolved solids and salinity. The
total dissolved solids concentration in
milligrams per liter (mg/L) can also be
calculated by multiplying the conductivity
result by a factor between 0.55 and 0.9,
which is empirically determined (see
Standard Methods #2510, APHA 1992).
-------
174 I WATER QUALITY CONDITIONS
Suitable conductivity meters cost about
$350. Meters in this price range should also
measure temperature and automatically
compensate for temperature in the conduc-
tivity reading. Conductivity can be mea-
sured in the field or the lab. In most cases,
it is probably better if the samples are
collected in the field and taken to a lab for
testing. In this way several teams of volun-
teers can collect samples simultaneously. If
it is important to test in the field, meters
designed for field use can be obtained for
around the same cost mentioned above.
If samples will be collected in the field
for later measurement, the sample bottle
should be a glass or polyethylene bottle that
has been washed in phosphate-free deter-
gent and rinsed thoroughly with both tap
and distilled water. Factory-prepared
Whirl-pak® bags may be used.
How to sample
The procedures for collecting samples
and analyzing conductivity consist of the
following tasks:
[ TASK 1 I Prepare the sample contain-
^^~^~"™"^ ers
If factory-sealed, disposable Whirl-
pak® bags are used for sampling, no
preparation is needed. Reused sample
containers (and all glassware used in this
procedure) must be cleaned before the first
run and after each sampling run by follow-
ing Method A as described on page 128.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, when sampling for conductiv-
ity, include the following equipment:
• Conductivity meter and probe (if
testing conductivity in the field)
• Conductivity standard appropriate
for the range typical of the stream
• Data sheet for conductivity to record
results
Be sure to let someone know where you
are going and when you expect to return.
TASK 3 \ Collect the sample (if
samples will be tested in the
lab)
Refer to page 128 for details on how to
collect water samples using screw-cap
bottles or Whirl-pak® bags.
TASK 4
Analyze the sample (field or
lab)
The following procedure applies to
field or lab use of the conductivity meter.
1. Prepare the conductivity meter for
use according to the manufacturer's
directions.
2. Use a conductivity standard solution
(usually potassium chloride or
sodium chloride) to calibrate the
meter for the range that you will be
measuring. The manufacturer's
directions should describe the
preparation procedures for the
standard solution.
3. Rinse the probe with distilled or
deionized water.
4. Select the appropriate range begin-
ning with the highest range and
working down. Read the conductiv-
ity of the water sample. If the reading
is in the lower 10 percent of the
range, switch to the next lower range.
If the conductivity of the sample
exceeds the range of the instrument,
you may dilute the sample. Be sure
to perform the dilution according to
the manufacturer's directions be-
cause the dilution might not have a
-------
WATER QUALITY CONDITIONS I 175
simple linear relationship to the
conductivity.
5. Rinse the probe with distilled or
deionized water and repeat step 4
until finished.
TASKS
Return the samples and the
field data sheets to the lab/
drop-off point.
Samples that are sent to a lab for
conductivity analysis must be tested within
28 days of collection. Keep the samples on
ice or refrigerated.
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
Hach Company. 1992. Each water analysis
handbook. 2nd ed. Loveland, CO.
Mississippi Headwaters River Watch. 1991.
Water quality procedures. Mississippi
Headwaters Board. March.
-------
176 I WATER QUALITY CONDITIONS
5.10
Total Alkalinity
What is total alkalinity and why is
it important?
Alkalinity is a measure of the capacity
of water to neutralize acids (see pH descrip-
tion). Alkaline compounds in the water
such as bicarbonates (baking soda is one
type), carbonates, and hydroxides remove
H+ ions and lower the acidity of the water
(which means increased pH). They usually
do this by combining with the H+ ions to
make new compounds. Without this acid-
neutralizing capacity, any acid added to a
stream would cause an immediate change in
the pH. Measuring alkalinity is important in
determining a stream's ability to neutralize
acidic pollution from rainfall or wastewater.
It's one of the best measures of the sensitiv-
ity of the stream to acid inputs.
Alkalinity in streams is influenced by
rocks and soils, salts, certain plant activi-
ties, and certain industrial wastewater
discharges.
Total alkalinity is measured by measur-
ing the amount o.f acid (e.g., sulfuric acid)
needed to bring the sample to a pH of 4.2.
At this pH all the alkaline compounds in the
sample are "used up." The result is reported
as milligrams per liter of calcium carbonate
(mg/L CaCO3).
Analytical ana equipment
considerations
For total alkalinity, a double endpoint
titration using a pH meter (or pH "pocket
pal") and a digital titrator or buret is
recommended. This can be done in the field
or in the lab. If you will analyze alkalinity
in the field, it is recommended that you use
a digital titrator instead of a buret because
the buret is fragile and more difficult to set
up and use in the field. The alkalinity
method described below was developed by
the Acid Rain Monitoring Project of the
University of Massachusetts Water Re-
sources Research Center.
Burets, titrators, and digital
titrators for measuring alkalinity
The total alkalinity analysis involves
titration. In this test, titration is the addition
of small, precise quantities of sulfuric acid
(the reagent) to the sample until the sample
reaches a certain pH (known as an end-
point). The amount of acid used corre-
sponds to the total alkalinity of the sample.
Alkalinity can be measured using a buret,
titrator, or digital titrator (described below).
• A buret is a long, graduated glass
tube with a tapered tip like a pipet
and a valve that is opened to allow
the reagent to drip out of the tube.
The amount of reagent used is
calculated by subtracting the original
volume in the buret from the volume
left after the endpoint has been
reached. Alkalinity is calculated
based on the amount used.
• Titrators forcefully expel the reagent
by using a manual or mechanical
plunger. The amount of reagent used
is calculated by subtracting the
original volume in the titrator from
the volume left after the endpoint has
been reached. Alkalinity is then
calculated based on the amount used
or is read directly from the titrator.
• Digital titrators have counters that
display numbers. A plunger is forced
into a cartridge containing the
reagent by turning a knob on the
titrator. As the knob turns, the
counter changes in proportion to the
amount of reagent used. Alkalinity is
then calculated based on the amount
used. Digital titrators cost approxi-
mately $90.
-------
WATER QUALITY CONDITIONS I 177
Digital titrators and burets allow for
much more precision and uniformity in the
amount of titrant that is used.
How to collect and analyze
samples
The field procedures for collecting and
analyzing samples for pH and total alkalin-
ity consist of the following tasks:
TASK 1 I Prepare the sample contain-
ers
Sample containers (and all glassware
used in this procedure) must be cleaned and
rinsed before the first run and after each
sampling run by following the procedure
described under Method A on page 128.
Remember to wear latex gloves.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 for details on
confirming sampling date and time, safety
considerations, checking supplies, and
checking weather and directions. In addi-
tion to the standard sampling equipment
and apparel, when sampling for pH and
alkalinity include the following equipment:
• Digital titrator
• 100-mL graduated cylinder
• 250-mL beaker
• pH meter with combination tempera-
ture and reference electrode or pH
"pocket pal"
• Sulfuric acid titration cartridge,
0.16 N
• Data sheet for pH and total alkalinity
to record results
• Alkalinity voluette ampules standard,
0.500 N, for accuracy check
• Wash bottle with deionized water to
rinse pH meter electrode
• Magnetic stirrer, if titrated in the lab
Be sure to calibrate the pH meter
before you analyze a sample. The pH meter
should be calibrated prior to sample
analysis and after every 25 samples accord-
ing to the instructions in the meter manual.
Use two pH standard buffer solutions: 4.01
and 7.0. Following are notes regarding
buffers:
• The buffer solutions should be at
room temperature when you cali-
brate the meter.
• Do not use a buffer after its expira-
tion date.
• Always cap the buffers during
storage to prevent contamination.
• Because buffer pH values change
with temperature, the meter must
have a built-in temperature sensor
that automatically standardizes the
pH when the meter is calibrated.
• Do not reuse buffer solutions!
Be sure to let someone know where
you are going and when you expect to
return.
TASK 3 j Collect the sample
Refer to page 128 for details on how to
collect water samples using screw-cap
bottles or Whirl-pak® bags.
TASK 4 j Measure total alkalinity (field
1 or lab)
The following steps are for use of a
digital titrator in the field or the lab. If you
are using a buret, consult Standard Meth-
ods (APHA, 1992).
Alkalinity is usually measured using
sulfuric acid with a digital titrator. Sulfuric
acid is added to the water sample in
measured amounts until the three main
forms of alkalinity (bicarbonate, carbonate,
and hydroxide) are converted to carbonic
acid. At pH 10, hydroxide (if present)
reacts to form water. At pH 8.3, carbonate
is converted to bicarbonate. At pH 4.5, it is
-------
178 I WATER QUALITY CONDITIONS
certain that all carbonate and bicarbonate
are converted to carbonic acid. Below this
pH, the water is unable to neutralize the
sulfuric acid and there is a linear relation-
ship between the amount of sulfuric acid
added to the sample and the change in the
pH of the sample. So, additional sulfuric
acid is added to the sample to reduce the
pH of 4.5 by exactly 0.3 pH units (which
corresponds to an exact doubling of the pH)
to a pH of 4.2. However, the exact pH at
which the conversion of these bases might
have happened, or total alkalinity, is still
unknown. This procedure uses an equation
derived from the slope of the line described
above to extrapolate back to the amount of
sulfuric acid that was added to actually
convert all the bases to carbonic acid. The
multiplier (0.1) then converts this to total
alkalinity as mg/L CaCO3. The following
steps outline the procedures necessary to
determine the alkalinity of your sample.
1. Insert a clean delivery tube into the
0.16 N sulfuric acid titration car-
tridge and attach the cartridge to the
titrator body.
2. Hold the titrator, with the cartridge
tip pointing up, over a sink. Turn the
delivery knob to eject air and a few
drops of titrant. Reset the counter to
0 and wipe the tip.
3. Measure the pH of the sample (see
pH, section, 5.4). If it is less than 4.5,
go to step 9 below.
4. Insert the delivery tube into the
beaker containing the sample. Turn
the delivery knob while magnetically
stirring the beaker until the pH meter
reads 4.5. Record the number of
digits used to achieve this pH. Do not
reset the counter.
5. Continue titrating to a pH of 4.2 and
record the number of digits.
6. Apply the following equation:
Alkalinity (as mg/L CaCO3) = (2a - b) x 0.1
Where:
a = digits of titrant to reach pH 4.5
b = digits of titrant to reach pH 4.2
(including digits required to get to
pH 4.5)
0.1 = digit multiplier for a 0.16 titration
cartridge and a 100-mL sample
Example:
Initial pH of sample is 6.5.
It takes 108 turns to get to a pH of 4.5.
It takes another 5 turns to get to pH 4.2,
for a total of 113 turns.
Alkalinity = ((2 x 108) -113) x 0.1
= 10.3 mg/L
7. Record the results as mg/L alkalinity
on the lab sheet.
8. Rinse the beaker with distilled water
before the next sample.
9. If the pH of your water sample, prior
to titration, is less than 4.5, proceed
as follows:
• Insert the delivery tube into the
beaker containing the sample.
• Turn the delivery knob while
swirling the beaker until the pH
meter reads exactly 0.3 pH units
less than the initial pH of the
sample.
• Record the number of digits used
to achieve this pH.
• Apply the equation as in step 6,
but a = 0 and b = the number of
digits required to reduce the initial
pH exactly 0.3 pH units.
Example:
Initial pH of sample is 4.3.
Enter "0" in the 4.5 column on the lab
sheet.
Titrate to a pH of 0.3 units less than the
initial pH—in this case 4.0.
It takes 10 digits to get to 4.0.
Enter this in the 4.2 column on the lab
sheet and note that the pH endpoint is
4.0.
Alkalinity = (0 -10) x 0.1 = -1.0.
-------
WATER QUALITY CONDITIONS I 179
• Record the results as mg/L alkalin-
ity on the lab sheet.
10. Perform an accuracy check on the
first field sample, halfway through
the run, and after analysis of the last
sample as described below. Check
the pH meter against pH 7.0 and 4.01
buffers after every 10 samples.
TASK 5 I Perform an accuracy check
This accuracy check should be per-
formed on the first field sample titrated,
again about halfway through the field
samples, and at the final field sample.
1. Snap the neck off an alkalinity
voluette ampule standard, 0.500 N.
Or if using a standard solution from a
bottle, pour a few milliliters of the
standard into a clean beaker.
2. Pipet 0.1 mL of the standard to the
titrated sample (see above). Resume
titration back to the pH 4.2 endpoint.
Record the number of digits needed.
3. Repeat using two more additions of
0.1 mL of standard. Titrate to the pH
4.2 after each addition.
4. Each 0.1-mL addition of standard
should require 250 additional digits
of0.16Ntitrant.
TASK 6
Return the field data sheets
and samples to the lab or
drop-off point
Alkalinity samples must be analyzed
within 24 hours of their collection. If the
samples cannot be analyzed in the field,
keep the samples on ice and take them to
the lab or drop-off point as soon as
sible.
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
Godfrey, P.J. 1988. Acid rain in Massachu-
setts. University of Massachusetts Water
Resources Research Center, Amherst,
MA.
River Watch Network. 1992. Total alkalin-
ity and pH field and laboratory proce-
dures (based on University of Massachu-
setts Acid Rain Monitoring Project). July
1.
-------
180 I WATER QUALITY CONDITIONS
5.11
Fecal Bacteria
What are fecal bacteria and why
are they important?
Members of two bacteria groups,
coliforms and fecal streptococci, are used
as indicators of possible sewage contamina-
tion because they are commonly found in
human and animal feces. Although they are
generally not harmful themselves, they
indicate the possible presence of pathogenic
(disease-causing) bacteria, viruses, and
protozoans that also live in human and
animal digestive systems. Therefore, their
presence in streams suggests that patho-
genic microorganisms might also be present
and that swimming and eating shellfish
might be a health risk. Since it is difficult,
time-consuming, and expensive to test
directly for the presence of a large variety
of pathogens, water is usually tested for
coliforms and fecal streptococci instead.
Sources of fecal contamination to surface
waters include wastewater treatment plants,
on-site septic systems, domestic and wild
animal manure, and storm runoff.
In addition to the possible health risk
associated with the presence of elevated
levels of fecal bacteria, they can also cause
cloudy water, unpleasant odors, and an
increased oxygen demand. (Refer to the
section on dissolved oxygen.)
Indicator bacteria types and what they can
tell you
The most commonly tested fecal
bacteria indicators are total coliforms, fecal
coliforms, Escherichia coli, fecal strepto-
cocci, and enterococci. All but E. coli are
composed of a number of species of
bacteria that share common characteristics
such as shape, habitat, or behavior; E. coli
is a single species in the fecal coliform
group.
Total coliforms are a group of bacteria
that are widespread in nature. All members
of the total coliform group can occur in
human feces, but some can also be present
in animal manure, soil, and submerged
wood and in other places outside the human
body. Thus, the usefulness of total
coliforms as an indicator of fecal contami-
nation depends on the extent to which the
bacteria species found are fecal and human
in origin. For recreational waters, total
coliforms are no longer recommended as an
indicator. For drinking water, total
coliforms are still the standard test because
their presence indicates contamination of a
water supply by an outside source.
Fecal coliforms, a subset of total
coliform bacteria, are more fecal-specific in
origin. However, even this group contains a
genus, Klebsiella, with species that are not
necessarily fecal in origin. Klebsiella are
commonly associated with textile and pulp
and paper mill wastes. Therefore, if these
sources discharge to your stream, you
might wish to consider monitoring more
fecal and human-specific bacteria. For
recreational waters, this group was the
primary bacteria indicator until relatively
recently, when EPA began recommending
E. coli and enterococci as better indicators
of health risk from water contact. Fecal
coliforms are still being used in many states
as the indicator bacteria.
E. coli is a species of fecal coliform
bacteria that is specific to fecal material
from humans and other warm-blooded
animals. EPA recommends E. coli as the
best indicator of health risk from water
contact in recreational waters; some states
have changed their water quality standards
and are monitoring accordingly.
Fecal streptococci generally occur in
the digestive systems of humans and other
warm-blooded animals. In the past, fecal
streptococci were monitored together with
fecal coliforms and a ratio of fecal
coliforms to streptococci was calculated.
This ratio was used to determine whether
-------
WATER QUALITY CONDITIONS I 181
the contamination was of human or nonhu-
man origin. However, this is no longer
recommended as a reliable test.
Enterococci are a subgroup within the
fecal streptococcus group. Enterococci are
distinguished by their ability to survive in
salt water, and in this respect they more
closely mimic many pathogens than do the
other indicators. Enterococci are typically
more human-specific than the larger fecal
streptococcus group. EPA recommends
enterococci as the best indicator of health
risk in salt water used for recreation and as
a useful indicator in fresh water as well.
Which Bacteria Should You Monitor?
Which bacteria you test for depends on
what you want to know. Do you want to
know whether swimming in your stream
poses a health risk? Do you want to know
whether your stream is meeting state water
quality standards?
Studies conducted by EPA to determine
the correlation between different bacterial
indicators and the occurrence of digestive
system illness at swimming beaches
suggest that the best indicators of health
risk from recreational water contact in fresh
water are E. coli and enterococci. For salt
water, enterococci are the best. Interest-
ingly, fecal coliforms as a group were
determined to be a poor indicator of the risk
of digestive system illness. However, many
states continue to use fecal coliforms as
their primary health risk indicator.
If your state is still using total or fecal
coliforms as the indicator bacteria and you
want to know whether the water meets state
water quality standards, you should monitor
fecal coliforms. However, if you want to
know the health risk from recreational
water contact, the results of EPA studies
suggest that you should consider switching
to the E. coli or enterococci method for
testing fresh water. In any case, it is best to
consult with the water quality division of
your state's environmental agency, espe-
cially if you expect them to use your data.
Sampling and equipment
considerations
Bacteria can be difficult to sample and
analyze, for many reasons. Natural bacte-
ria levels in streams can vary significantly;
bacteria conditions are strongly correlated
with rainfall, and thus comparing wet and
dry weather bacteria data can be a problem;
many analytical methods have a low level
of precision yet can be quite complex; and
absolutely sterile conditions are required to
collect and handle samples.
The primary equipment decision to
make when sampling for bacteria is what
type and size of sample container you will
use. Once you have made that decision, the
same, straightforward collection procedure
is used regardless of the type of bacteria
being monitored. Collection procedures are
described under "How to Collect Samples"
below.
It is critical when monitoring bacteria
that all containers and surfaces with which
the sample will come into contact be
sterile. Containers made of either some
form of plastic or Pyrex glass are accept-
able to EPA. However, if the containers are
to be reused, they must be sterilized using
heat and pressure. The containers can be
sterilized by using an autoclave, which is a
machine that sterilizes containers with
pressurized steam. If using an autoclave,
the container material must be able to
withstand high temperatures and pressure.
Plastic containers—either high-density
polyethylene or polypropylene—might be
preferable to glass from a practical stand-
point because they will better withstand
breakage. In any case, be sure to check the
manufacturer's specifications to see
whether the container can withstand 15
minutes in an autoclave at a temperature of
121 °C without melting. (Extreme caution
is advised when working with an auto-
clave.) Disposable, sterile, plastic Whirl-
pak® bags are used by a number of pro-
grams. The size of the container will
depend on the sample amount needed for
-------
182 I WATER QUALITY CONDITIONS
the bacteria analysis method you choose
and the amount needed for other analyses.
There are two basic methods for
analyzing water samples for bacteria:
1. The membrane filtration method
involves filtering several different-
sized portions of the sample using
filters with a standard diameter and
pore size, placing each filter on a
selective nutrient medium in a petri
plate, incubating the plates at a
specified temperature for a specified
time period, and then counting the
colonies that have grown on the
filter. This method varies for differ-
ent bacteria types (variations might
include, for example, the nutrient
medium type, the number and types
of incubations, etc.).
2. The multiple-tube fermentation
method involves adding specified
quantities of the sample to tubes
containing a nutrient broth, incubat-
ing the tubes at a specified
temperature for a specified time
period, and then looking for the
development of gas and/or turbidity
that the bacteria produce. The
presence or absence of gas in each
tube is used to calculate an index
known as the Most Probable Number
(MPN).
Given the complexity of the analysis
procedures and the equipment required,
field analysis of bacteria is not recom-
mended. Bacteria can either be analyzed by
the volunteer at a well-equipped lab or sent
to a state-certified lab for analysis. If you
send a bacteria sample to a private lab,
make sure that it is certified by the state for
bacteria analysis. Consider state water
quality labs, university and college labs,
private labs, wastewater treatment plant
labs, and hospitals. You might need to pay
these labs for analysis.
This manual does not address labora-
tory methods because several bacteria types
are commonly monitored and the methods
are different for each type. For more
information on laboratory methods, refer to
the references at the end of this section.
If you decide to analyze your samples
in your own lab, be sure to carry out a
quality assurance/quality control program.
Specific procedures are recommended in
the section below.
How to Collect Samples
The procedures for collecting and
analyzing samples for bacteria consist of
the following tasks:
TASK 1 | Prepare sample containers
If factory-sealed, presterilized, dispos-
able Whirl-pak® bags are used to sample,
no preparation is needed. Any reused
sample containers (and all glassware used
in this procedure) must be rinsed and
sterilized at 121 °C for 15 minutes using an
autoclave before being used again for
sampling.
TASK 2
Prepare before leaving for
the sampling site
Refer to pages 19-21 of the introduc-
tion for details on confirming sampling data
and time, picking up equipment, reviewing
safety considerations, and checking weather
and directions. In addition, to sample for
coliforms you should check your equipment
as follows:
• Whirl-pak® bags are factory-sealed
and sterilized. Check to be sure that
the seal has not been removed.
• Bottles should have tape over the cap
or some seal or marking to indicate
that they have been sterilized. If any
of the sample bottles are not num-
bered, ask the lab coordinator how to
number them. Unless sample con-
tainers are to be marked with the site
number, do not number them your-
self.
-------
WATER QUALITY CONDITIONS I 183
TASK 3 | Collect the sample
Refer to page 128 for details on collect-
ing a sample using screw-cap bottles or
Whirl-pak® bags. Remember to wash your
hands thoroughly after collecting samples
suspected of containing fecal contamina-
tion. Also, be careful not to touch your
eyes, ears, nose, or mouth until you've
washed your hands.
Recommended field quality assurance/
quality control procedures include:
• Field Blanks. These should be
collected at 10 percent of your
sample sites along with the regular
samples. Sterile water in sterilized
containers should be sent out with
selected samplers. At a predeter-
mined sample site, the sampler fills
the usual sample container with this
sterile water. This is labeled as a
regular sample, but with a special
notation (such as a "B") that indi-
cates it is a field blank. It is then
analyzed with the regular samples.
Lab analysis should result in "0"
bacteria counts for all blanks. Blanks
are used to identify errors or con-
tamination in sample collection and
analysis.
• Internal Field Duplicates. These
should be collected at 10 percent of
your sampling sites along with the
regular samples. A field duplicate is
a duplicate stream sample collected
at the same time and at the same
place either by the same sampler or
by another sampler. This is labeled
as a regular sample, but with a
special notation (such as a "D") that
indicates it is a duplicate. It is then
analyzed with the regular samples.
Lab analysis should result in compa-
rable bacteria counts per 100 mL for
duplicates and regular samples
collected at the same site. Duplicates
are used to estimate sampling and
laboratory analysis precision.
External Field Duplicates. An
external field duplicate is a duplicate
stream sample collected and pro-
cessed by an independent (e.g.,
professional) sampler or team at the
same place at the same time as
regular stream samples. It is used to
estimate sampling and laboratory
analysis precision.
TASK 4 | Return the field data sheets
<"""""^~~^ and the samples to the lab or
drop-off point
Samples for bacteria must be analyzed
within 6 hours of collection. Keep the
samples on ice and take them to the lab or
drop-off point as soon as possible.
TASK 5 I Analyze the samples in the
1 lab
This manual does not address labora-
tory analysis of water samples. Lab meth-
ods are described in the references below
(APHA, 1992; River Watch Network,
1991; USEPA, 1985). However, the lab
you work with should carry out the follow-
ing recommended laboratory quality
assurance/quality control procedures:
• Negative Plates result when the
buffered rinse water (the water used
to rinse down the sides of the filter
funnel during filtration) has been
filtered the same way as a sample.
This is different from a field blank in
that it contains reagents used in the
rinse water. There should be no
bacteria growth on the filter after
incubation. It is used to detect
laboratory bacteria contamination of
the sample.
• Positive Plates result when water
known to contain bacteria (such as
wastewater treatment plant influent)
is filtered the same way as a sample.
There should be plenty of bacteria
growth on the filter after incubation.
-------
184 I WATER QUALITY CONDITIONS
Positive plates are used to detect
procedural errors or the presence of
contaminants in the laboratory
analysis that might inhibit bacteria
growth.
Lab Replicates. A lab replicate is a
sample that is split into subsamples
at the lab. Each subsample is then
filtered and analyzed. Lab replicates
are used to obtain an optimal number
of bacteria colonies on filters for
counting purposes. Usually,
subsamples of 100, 10, and 1 millili-
ter (mL) are filtered to obtain
bacteria colonies on the filter that can
be reliably and accurately counted
(usually between 20 and 80 colo-
nies). The plate with the count
between 20 and 80 colonies is
selected for reporting the results, and
the count is converted to colonies per
lOOmL.
Knowns. A predetermined quantity
of dehydrated bacteria is added to the
reagent water, which should result in
a known result, within an acceptable
margin of error.
Outside Lab Analysis of Duplicate
Samples. Either internal or external
field duplicates can be analyzed at an
independent lab. The results should
be comparable to those obtained by
the project lab.
River Watch Network. 1991. Escherichia
coli (E. coli) membrane filter procedure
(adapted from USEPA Method 1103.1,
1985). Montpelier, VT. October.
USEPA. 1985. Test methods for Escheri-
chia coli and enterococci in water by the
membrane filter procedure (Method
#1103.1). EPA 600/4-85-076. U.S.
Environmental Protection Agency,
Environmental Monitoring and Support
Laboratory, Cincinnati, OH.
USEPA. 1986. Bacteriological ambient
water quality criteria for marine and
fresh recreational waters. EPA 440/5-84-
002. U.S. Environmental Protection
Agency, Office of Research and Devel-
opment, Cincinnati, OH.
References
APHA. 1992. Standard methods for the
examination of water and wastewater.
18th ed. American Public Health Asso-
ciation, Washington, DC.
Hogeboom, T. Microbiologist, Vermont
Environmental Conservation Laboratory,,
Waterbury, VT. Personal communica-
tion.
-------
WATER QUALITY CONDITIONS I 185
LU
Ul
CO
g
Q
Q
LJJ
LL
O
-1
i*
Q) oJ
0) "**
' c
CO
-I
a> 5
U. M
(0
i
CD
CO O
0
U
0>
El
CO ^
75
£
n
Q.
,
o
DC
LJJ
i
|
£
i=
OT
0)
to
o
O
£
g
f
i
•s
Si
"w
0)
8
a.
<
!
c
-a
|
0)
I
o
2 1.
I "5
fB 0)
•i -8
.*•
0)
•S
o
S«
2
E
OT
2
I
o
I
f
>^2
OO
aa
-------
186 I WATER QUALITY CONDITIONS
-------
MANAGING AND PRESENTING VOLUNTEER DATA I 187
-------
188 I MANAGING AND PRESENTING VOLUNTEER DATA
It is hard to overemphasize the impor-
tance of having established methods of
handling volunteer data, analyzing that
data, and presenting results effectively to
volunteers, the public, and water resource
decision-makers. Without these tools and
processes, the data that volunteers and
program managers have labored hard to
collect are virtually useless, and the pro-
gram will surely fail to meet its goals.
This chapter addresses data manage-
ment and data presentation. Members of the
program planning committee will need to
make many decisions on these issues before
the first field data sheet is filled out by the
program's first volunteer. In particular, they
should consult any potential data users such
as state water quality agencies or county
planning boards regarding their own data
needs. Data users will be particularly
concerned about:
• Procedures used to verify and check
the raw volunteer data.
• Databases and software used to
manage the data.
• Analytical procedures used to
convert the raw data into findings
and conclusions.
• Reporting formats.
Data users may, for example, be able to
offer concrete suggestions about databases
and presentation formats that will make the
data more accessible to them. To ensure
that all questions about the validity of the
data can be answered, the program planning
committee should develop and implement a
quality assurance/quality control plan
designed to minimize data collection errors,
weed out data that fail to meet the
program's standards, and effectively
analyze and present the results. This plan
should identify key personnel with respon-
sibilities for data management and data
analysis and clearly indicate all the steps
the program will take to handle the data.
Unfortunately, volunteers and program
coordinators seldom recognize the impor-
tance of this aspect of a volunteer monitor-
ing program. It tends to be considered
"drudge" work assigned to one or two
technically- inclined people. However, that
attitude is seriously out of date. Program
organizers should make every effort to
involve a range of volunteers and program
staff in all aspects of data management and
presentation. Sufficient time should be
budgeted to the tasks that are involved.
People who produce the reports should be
acknowledged. After all, it is the final
reports that will be reviewed by stream
management decision-makers, not the field
data sheets. No other tasks are more
important to the success of the volunteer
stream monitoring program.
-------
MANAGING AND PRESENTING VOLUNTEER DATA I 189
6.1
Managing Volunteer
Data
The following steps will help ensure
that the data collected by volunteers are
well managed, credible, and of value to
potential data users.
Review Field Data Sheets
The volunteer program coordinator or
designated analyst should screen and
review the field data sheets as they are
received. This involves some basic "reality
checks." Questions that should be kept in
mind include the following:
• Are the results as might be antici-
pated, or are they highly
unexpected? If unexpected, are they
still within the realm of possibility?
For example, can the kit or technique
the volunteer used actually produce
results like that? Does the volunteer
offer any possible explanations for
the results (e.g., a sewage treatment
plant malfunction had been recently
reported) or corollary information
(e.g., a fish kill has been observed
along with the extremely low dis-
solved oxygen readings)? Also check
for consistency between similar
parameters. For example, total
dissolved solids and conductivity
should track together—if one goes
up, so should the other. So should
total solids and turbidity.
• Are there outliers? (Findings that
differ radically from past data or
other data from similar sites.)
Values that are off by a factor of 10
or 100 should be questioned. Follow
up on any data that seems suspect. If
you can't come up with an explana-
tion for why the results are so
unusual, but they are still within the
realm of possibility, you may want
to flag the data as questionable. Ask
an experienced volunteer or program
staffer to sample at that site as a
backup until uncertainties are
resolved, or work with the volunteer
to verify that proper sampling and
analytical protocols are being
followed.
• Are the field data sheets complete?
If a volunteer is consistently leaving
a section of the sheet incomplete,
follow up and ask why. Instructions
may not always be easily under-
stood. All sheets should include site
location and identification, name of
the volunteer, date, time, and
weather conditions.
• Are all measurements reported in the
correct units ?
You should minimize the chance for
error by including on the data form
itself any equations needed to
convert measurements, and specify
on the form what units should be
used. Check the math. All field data
sheets should be kept on .file in the
event that findings are brought into
question at a later date.
Review Information in Your
Database
Once volunteer data enters a computer-
ized database, it can take on a life of its
own. It is a phenomenon of human nature
that data suddenly seem more believable
once computerized. Therefore, be sure to
carefully screen information as soon as you
enter it into a database. Then review a
printout (preferably with a fresh pair of
eyes) against the original field data sheets.
One way to minimize transcription errors is
to design the computer input screens to
look like the field data forms.
-------
190 I MANAGING AND PRESENTING VOLUNTEER DATA
As a further check, you can run some
simple calculations like determining
medians and means to make sure no errors
have slipped through. (If the median and
the mean are very different, an outlier may
be skewing the results.) Again, if you
uncover unusual data points that cannot be
explained by backup information on the
field data sheets or the comment field in the
database, flag the data as questionable until
it can be verified.
Review Your Final Results
Once volunteer monitoring data has
been entered into a database, the next step
is to generate reports on the findings of the
data. Even at this stage you should continue
to look for inconsistencies and problems.
For example, you should:
• Review findings against previous
years' data.
• Look for outliers on graphs and
maps.
• Not remove data just because you
don't like it, but do investigate
findings that are unusual or can't be
explained.
By the time you present your final
results to your volunteers or other data
users, you should feel fully confident that
you have assembled the best possible
picture of water quality conditions in your
study streams.
Develop a Coding System
A coding system will help simplify the
tracking and recording of data. Make sure,
however, that the system you create is
easily understood and simple to use. Codes
developed for sample sites, parameters, and
other information on field and lab sheets
should parallel the codes you use in your
database. If you will be sharing your
information with a state or local natural
resource agency, you may want your
coding system to match or complement the
agency system.
Sample Sites: Because sample sites tend
to change over time, it is important to have
a site numbering system that accommodates
change. A good convention to follow is to
use a site coding system that includes an
abbreviation of the waterbody and a site
number (e.g., CtR020 for a site on the
Connecticut River). For consistency, you
might choose to start the site numbers at the
downstream end of the stream and increase
them as you move upstream (e.g., the first
Connecticut River site would be CtROlO,
the second CtR020, etc.). Leave extra
numbers between sites to allow for your
program's future expansion.
Water Quality Parameters: It is also
important to develop a coding system for
each of the water quality parameters you
are testing. These are the codes you will use
in the database to identify and extract
results. To keep the amount of clerical work
to a minimum, abbreviate without losing
the ability to distinguish parameters from
one another. For example, EC could
represent E. coli bacteria and FC fecal
coliform bacteria.
Spreadsheets, Databases, and
Mapping Software
Today's computer software includes a
variety of spreadsheet and database pack-
ages that allow you to sort, manipulate, and
perform statistical analyses on the data you
have entered into the computer. For most
applications, spreadsheets are adequate and
have the advantage of being relatively
simple to use. Most spreadsheet packages
have graphics capabilities that will allow
you to plot your data onto a graph of your
choice (i.e., bar, line, or pie chart). Ex-
amples of common spreadsheet software
packages are Lotus 1-2-3, Excel, and
Quattro Pro.
Database software may be more
difficult to master and usually lack the
graphics capabilities of spreadsheet soft-
ware. If you manage large amounts of data,
however, a database is almost a necessity.
-------
MANAGING AND PRESENTING VOLUNTEER DATA I 191
Using a database, you can store and ma-
nipulate very large data sets without
sacrificing speed. The database can also
relate records in one file to records in
another file. This allows you to break your
data up into smaller, more easily managed
files that can work together as though they
were one.
If you use database software for storage
and retrieval, you may still want to use a
spreadsheet or other program with graphics
capabilities. Many spreadsheet and data-
base software packages are compatible and
will allow you to transport sets of data back
and forth with relative ease. Very large data
sets can be organized and manipulated in a
database. Specific parts of the data (such as
results for a particular metric from all
stations and all sampling events) can then
be transported into the spreadsheet, statisti-
cally analyzed, and graphically displayed.
Examples of popular database software
packages are dBase, FileMaker Pro, and
FoxPro.
An effective way to display your data is
on a map of the stream or watershed. This
clearly illustrates the relationship between
land uses and the quality of water, habitat,
and biological communities. This type of
graphic display can be used to effectively
show the correlation between specific
activities or land uses and the impacts they
have on the ecosystem. Simple personal
computer-based mapping packages are
available. They allow you to enter layers of
data and conduct spatial analysis of that
data.
Systems that allow you to map and
manipulate various layers of information
(such as water quality data, land use
information, county boundaries, or geologic
conditions) are known as Geographic
Information Systems (GIS). They can vary
from simple systems run on personal
computers to sophisticated and very power-
ful systems that run on large main frames.
For any GIS application, you need to know
the coordinates of your sample sites—either
their latitude and longitude, or some
alternate system such as an EPA River
Reach File identifier. You can also locate
your sites on a topographic map that can be
digitized on to an electronic map of the
watershed. Once these points have been
established, you can link your database to
the points on the map, query your database,
and create graphic displays of the data.
Powerful GIS applications typically
require expensive hardware, software, and
technical training. Any volunteer program
interested in GIS applications should
consider working in partnership with other
organizations such as universities, natural
resource agencies, or large nonprofit
groups that can provide access to a GIS.
Many people are capable of writing
their own programs to manipulate and
display data. The disadvantage of using a
"homegrown" software program, however,
is that if its author leaves, so too does all
knowledge about how the program works.
Commercial software, on the other hand,
comes with consumer services that provide
over-the-phone help and instructions,
user's guides, replacement guarantees, and
updates as the company improves its
product. Also, most commercial programs
are developed to easily import and export
data in standard formats. This feature is
important if you want to share data with
other programs or organizations—all you
need are compatible software programs.
STORET
EPA's national water and biological data storage and retrieval
system, STORET, is being modernized and will be available in
1998-1999. Volunteer programs are encouraged to enter their data
into the modernized STORET. Individual systems will "feed" data
to a centralized file server which will permit national data analyses
and through which data can be shared among organizations. A
specific set of quality control measures will be required for any
data entered into the system to aid in data sharing. For more
information, see the EPA web page at www.epa.gov/owow/
STORET.
-------
192 I MANAGING AND PRESENTING VOLUNTEER DATA
Fig. 6.1
6.2
Presenting the Data
When presenting numerical data, one of
your chief goals should be to maintain the
attention and interest of your audience. This
is very difficult using tables filled with
numbers. Most people will not be interested
in the absolute values of each parameter at
each sampling site. Rather, they will want
to know the bottom line for each site (e.g.,
is it good or bad) and seasonal and year to
year trends.
Graphs and charts, therefore, are
typically the best way to present volunteer
data. Take care, however, that your graphs
"fit" your audience and are neither too
technical nor too simplistic.
Example of a
bar graph
displaying
biological data
Habitat scores as a percent of reference
condition at sites #1 and #2 for 1992-1994
0)
u
c
£
0)
W-»
O
CC
OT
CO
O
o
en
100 -
80-
60-
2 20-
40-
Site #1
Site #2
Graphs and Charts
Graphs can be used to display the
summarized results of large data sets and to
simplify complicated issues and findings.
The three basic types of graphs that are
typically used to present volunteer monitor-
ing data are:
• Bar graph
• Line graph
• Pie chart
Bar and line graphs are typically used
to show results, such as bioassessment
scores, along a vertical or y-axis for a
corresponding variable (such as sampling
date or site) which is marked along the
horizontal or x-axis. These types of graphs
can also have two vertical axes, one on
each side, with two sets of results shown in
relation to each other and to the variable
along the x-axis.
Bar Graph
A bar graph uses columns with heights
that represent the value of the data point for
the parameter being plotted. Fig. 6.1 is an
example using fictional data from Volun-
teer Creek.
Line graph
A line graph is constructed by connect-
ing the data points with a line. It can be
effectively used for depicting changes over
time or space. This type of graph places
more emphasis on trends and the relation-
ship among data points and less emphasis
on any particular data point.
Fig. 6.2 is an example of a line graph
again using fictional data from Volunteer
Creek.
Pie chart
Pie charts are used to compare catego-
ries within the data set to the whole. The
proportion of each category is represented
by the size of the wedge. Pie charts are
popular due to their simplicity and clarity.
(See Fig. 6.3)
-------
MANAGING AND PRESENTING VOLUNTEER DATA I 193
June phosphorus concentrations
at Sites #1 and #2 from 1991 - 1997
o>
E
(0
(0
(0
o
.c
Q.
(0
O
0.20
0.15- •
0.10
0.05---
0.00
Fig. 6.2
mmmmi^m
Example of a
line graph
depicting
trends in
phosphorus
data
6/91 6/92 6/93 6/94 6/95 6/96 6/97
Graphing Tips
Regardless of which graphic style you
choose, follow these rules to ensure you use
them most effectively.
• Each graph should have a clear
purpose. The graph should be easy to
interpret and should relate directly to
the content of the text of a document
or the script of a presentation.
• The data points on a graph should be
proportional to the actual values so
as not to distort the meaning of the
graph. Labeling should be clear and
accurate and the data values should
be easily interpreted from the scales.
Do not overcrowd the points or
values along the axes. If there is a
possibility of misinterpretation,
accompany the graph with a table of
the data.
• Keep it simple. The more complex
the graph, the greater the possibility
for misinterpretation.
Fig. 6.3
Summary of water quality ratings
for Volunteer Creek
Example of a
pie chart
summarizing
water quality
ratings
(Total no. ofstations= 52)
-------
194 I MANAGING AND PRESENTING VOLUNTEER DATA
• Limit the number of elements. Pie
charts should be limited to five or six
wedges, the bars in a bar graph
should fit easily, and the lines in a
line graph should be limited to three
or less.
• Consider the proportions of the
graph and expand the elements to fill
the dimensions, thereby creating a
balanced effect. Often, a horizontal
format is more visually appealing
and makes labeling easier. Try not to
use abbreviations that are not obvi-
ous to someone who is unfamiliar
with the program.
• Create titles that are simple, yet
adequately describe the information
portrayed in the graph.
• Use a legend if one is necessary to
describe the categories within the
graph. Accompanying captions may
also be needed to provide an ad-
equate description of the elements.
Summary Statistics
Summary statistics can reduce a very
large data set to a few numerical values that
can then be easily described and analyzed.
Such statistics include the mean and
standard deviation—two of the most
frequently used descriptors of environmen-
tal data.
Textbook statistics commonly assume
that if a parameter is measured many times
under the same conditions, then the mea-.
surement values will be randomly distrib-
uted around the average with more values
clustering near the average than further
away. In this ideal situation, a graph of the
frequency of each measure plotted against
its magnitude should yield a bell-shaped or
normal curve. The mean and the standard
deviation determine the height and breadth
of this curve, respectively.
The mean is simply the sum of all the
measurement values divided by the number
of measurements. This statistic is a measure
of location and in a normal curve marks the
highest point at the center of the bell.
The standard deviation, on the other
hand, describes the variability of the data
points around the mean. Very similar
measurement values will have a small
standard deviation while widely scattered
data will have a much larger standard
deviation.
While both the mean and standard
deviation are quite useful in describing
stream data, often the actual measures do
not fit a normal distribution. Other statistics
often come into play to describe the data.
Some data are skewed in one direction or
the other. Other data may have a flattened
bell shape.
It is important to note that biological
information often does not follow normal,
bell-shaped distribution. This is because
biological communities are dynamic,
complex, and interdependent systems;
many factors influence them, and these
cannot be statistically predicted. For
example, bioassessment scores plotted
against habitat assessment scores will be at
their best when habitat quality is at its best.
For data that is nonnormally distributed, the
mean and the standard deviation are not
appropriate summary statistics.
For describing nonnormally distributed
data, it is best to use statistics that can
convey the information for a variety of
conditions and which are not overly influ-
enced by the data points at the extremes of
the distribution. The median and the
interquartile range are two statistics that are
commonly used to describe the central
tendency and the spread around the median,
respectively. These statistics are derived by
placing the data points in order of value
from lowest to highest. The median is
simply the value that is in the middle of the
data set. The interquartile range is the
difference between the value at the 75
percent level and the value at the 25 percent
level.
-------
MANAGING AND PRESENTING VOLUNTEER DATA
The best method for presenting this
type of data is called a box and whisker
plot. One simple box and whisker plot will
graphically display the following informa-
tion:
Median
• Variability of the data around the
median
• Skew of the data
• Range of the data
• Size of the data set
Statistical software packages for
computers will easily construct box and
whisker plots. You can construct these plots
by following procedure shown below:
1. Order the data from the lowest to the
highest.
2. Plot the lowest and highest values on
the graph as short horizontal lines.
These are the extreme values of the
data set and represent the data range.
3. Determine the 75 percent value and
the 25 percent value of the data set.
These values define the interquartile
range and are represented by the
location of the top and bottom lines
of the box.
4. The horizontal length of the lines that
define the top and bottom lines of the
box (the box width) can be used as a
relative indication of the size of the
data set. For example, the box width
that describes a data set of 20 values
can be displayed twice as wide as a
data set of 10 values. Any propor-
tional scheme can be used as long as
it is consistently applied.
5. Close the box by drawing vertical
lines that connect to the ends of the
horizontal lines.
6. Plot the median inside the box.
Fig. 6.4 is an example depicting the
extreme values, interquartile range, and
median of biosurvey metric scores from 52
sites sampled in Volunteer Creek in June,
1995.
Maps
Displaying the results of your monitor-
ing data on a map can be a very effective
way of showing the data and helping
people understand what it means. A map
shows the location of sample sites in
relation to land features, such as cities,
wastewater treatment plants, farmland, and
tributaries that may have an effect on water
quality. Because a map also displays the
stream's relationship to neighborhoods,
parks and recreational areas, it can help to
develop concern for the stream and
strengthens interest in protecting it.
Choosing a Map
It is best to have two types of maps.
One should be a working map with a lot of
detail. The other should be used for display
Box Plot of Total Metric
Scores from June, 1995
(No. of sites = 52)
25
o
o
V)
o
4->
O
20-
15-
10-
5-
Fig. 6.4
•^••M
Example of a
box plot
Maximum value
(24)
75% value
(20)
Median (50%)
value
(14)
25% value
(8)
Minimum value
(2)
-------
196 I MANAGING AND PRESENTING VOLUNTEER DATA
purposes. The working map should include
important features such as:
• Stream and its tributaries
• Wetlands
• Lakes and ponds
• Cultural features such as roads
• Rail and power lines; municipal
boundaries
• Some indication of land use patterns
and vegetation.
The map should be of a scale large
enough to add the location of sample sites.
U.S. Geological Survey (USGS) 7.5
minute quads (scale of 1:24,000; 1 in. =
2,000 ft) are available with and without
topographic contours (elevation markings).
These maps are available for the most of
the United States.
The USGS maps are particularly useful
if your information will be incorporated
into a geographic information system
(GIS), since many of these systems use the
USGS maps as base maps. For your data to
be used in a GIS, it is likely that you will
have to provide the latitude and longitude
of your sample sites, which can be obtained
by using the grid markings on the USGS
topographic maps. Several different coordi-
nate systems are marked, including stan-
dard latitude/longitude and the Universal
Transmercator coordinates. For assistance
in learning how to use these coordinate
markings, talk to the local USGS office or
someone in the geography department at a
university. It may also be possible for the
GIS office you work with you to "digitize"
the maps, thus saving you the trouble of
trying to calculate the coordinates.
The display map is best used to illus-
trate your program results at public meet-
ings or in reports. This map should be
simpler than the detailed map and show
only principal features such as roads,
municipal boundaries, and waterways. It
should have sufficient detail and scale to
show the location of sample sites, and have
space for summary information about each
of the sample sites. Commercial road
atlases and county or town road maps
available from state transportation depart-
ments are examples of the types of maps
that can be used for display purposes (See
Fig. 6.5).
Creating a Display Map
Some suggestions for using a map to
display your data include:
• Keep the amount of information
presented on each map to a mini-
mum. Do not try to put so much on
one map that it becomes visually
complicated and difficult to read or
understand. Use another map to
display a different layer or "view" of
the data. For example, if there are
several dates for which you wish to
display sampling results, use one
map for each date.
• Clearly label the map and provide an
explanation of how to interpret it. If
you need a long and complicated
explanation, you may want to present
the data differently. If you have
reached a clear conclusion, state the
conclusion on the map. For example,
if a map shows that tributaries are
cleaner than the mainstem, use that
information as the subtitle of the
map.
• Provide a key to the symbols that are
used on the map.
• Rather than packing lots of informa-
tion into a small area of the map, use
a "blow-up" or enlargement of the
area elsewhere on the map to ad-
equately display the information.
• Use symbols that vary in size and
pattern to represent the magnitude of
results. For example, a site with a
fecal coliform level of 10 per 100
milliliters could be a light gray circle
one-sixteenth inch diameter while a
site with a level of 200 per 100
-------
MANAGING AND PRESENTING VOLUNTEER DATA I 197
BtiRKE LAKE
CLUSTER
Figure 6.5
A road map
is useful for
displaying
station loca-
tions.
milliliters would be a dark gray circle
one-quarter inch diameter. Start by
finding the highest and lowest
values, assign diameters and patterns
to those and then fill in steps along
the way. For the above example you
might have four ranges: 0 to 99, 100
to 199, 200 to 500 and 500 +.
Maps on Demand
EPA provides a World Wide Web service known as Maps on
Demand that allows users to generate maps displaying environ-
mental information for anywhere in the U.S. (except Hawaii, Puerto
Rico, and the Virgin Islands). Types of information that can be
mapped include EPA-regulated facilities, demographic information,
roads, streams, and drinking water sources. Maps of varying
scales can be generated on the site (latitude and longitude), zip
code, county, and basin levels. Submit your request and email
address, and after a brief wait, you will be able to view your map
on-line or download it. Maps on Demand can be reached through
EPA's Surf Your Watershed homepage at www.epa.gov/surf/
info.html.
-------
198 I MANAGING AND PRESENTING VOLUNTEER DATA
6.3
Producing Reports
On a regular basis, a successful stream
volunteer monitoring program should
produce reports that summarize key find-
ings to volunteers; data users such as state
water quality agencies, and local planning
boards; and/or the general public, including
the media. State water quality agencies will
require detailed reports, whereas shorter
and less technical summaries are more
appropriate for the general public. All
reports should be subjected to the review
process prescribed by your Quality Assur-
ance Project Plan.
Professional Report
In a report designed for water quality or
planning professionals, you should go into
detail about:
• The purpose of the study
• Who conducted it
• How it was funded
• The methods used
• The quality control measures taken
• Your interpretation of the results
• Your conclusions and recommenda-
tions
• Further questions that have arisen as
a result of the study.
Graphics, tables and maps may be
fairly sophisticated. Be sure to include the
raw data in an appendix and note any
problems encountered.
Lay Report
A report for the general public should
be short and direct. It is very important to
write in a non-technical style and to include
definitions for terms and concepts that may
be unfamiliar to the lay person. Simple
charts, summary tables, and maps with
accompanying explanations can be espe-
cially useful. This type of report should
include a brief description of the program,
the purpose of the monitoring, an explana-
tion of the parameters that were monitored,
the location of sample sites, a summary of
the results, and any recommendations that
may have been made.
Both types of reports should acknowl-
edge the volunteers and the sources of
funding.
Publicizing the Report
Develop a strategy for distributing and
publicizing your report before it is com-
pleted. Be sure the planning committee is
confident about the data and comfortable
with the statements and conclusions that
have been included in the document. When
the report is released to the public, you will
need to be prepared to respond to questions
regarding the data and your interpretation
of that data.
Some ideas for distributing the results
and informing the public include the
following:
• Mailing the report. If you have
access to a mailing list of people who
are interested in your stream, mail
the report with a cover letter that
summarizes the major findings of the
study. The cover letter should be
brief and enticing so that the recipi-
ent will be curious enough to read
the report. If you want people to take
some kind of action, such as support-
ing the expenditure of public funds to
upgrade a sewage treatment plant,
you may want to ask for their support
in the cover letter. If you do not have
an extensive mailing list, perhaps
other organizations that share your
goals would be willing to supply you
with their list. Be sure to also send
the report to the newspapers, radio
and television stations, and state and
federal agencies.
-------
MANAGING AND PRESENTING VOLUNTEER DATA I 199
Speaking tour. You may also want to
develop an oral presentation (with
slides, overheads, etc.) that could be
offered to groups such as the Cham-
ber of Commerce, Rotary clubs,
conservation organizations, schools,
and government entities. Your
presentation could even be video-
taped for distribution to a wider
audience.
Public meetings. You may want to
schedule a series of public meetings
that highlight the program and its
findings and recommendations. At
the meetings, distribute the report,
answer questions and tell your
audience how they can get involved.
These meetings can also help you
recruit more volunteers.
Be sure to schedule the meetings
at times when people are more likely
to attend (i.e., weekday evenings,
weekend days) and avoid periods
when people are normally busy or on
vacation. Invite the media and
publicize the meetings in newspaper
calendars, send press releases to
newspapers, radio and television
stations and other organizations, and
ask volunteers to distribute flyers at
grocery stores, city hall, etc.
News releases. Writing and distribut-
ing a news release is a cost effective
means of informing the public about
the results and accomplishments of
your program. Develop a mailing list
of newspapers, radio and television
stations, and organizations that
solicit articles for publication. Send
the news release to volunteers and
others who are interested in publiciz-
ing the monitoring program.
The first page of your news
release should feature the sponsoring
organization's name and logo to
clearly designate the source of the
news. Include a headline, the date, a
contact name and number, and
whether the story is for release
immediately or a later date. The first
paragraph should begin with a
dateline (the city of origin for the
event or story described in the
release) and include the essentials:
who, what, where, when, and why
and a synopsis of the most important
elements of the story. The second
paragraph should contain the second
most important facts, the third
paragraph the third most important
points and so on. Editors tend to
chop off the last paragraphs if short
on space. Therefore, be sure to state
your major points early in the press
release.
News conferences. If your report
contains some real news, or if it has
led to a significant event, (e.g., the
mayor or city council has recognized
the value of the report and issued a
statement of support) hold a news
conference. Timing and location are
important. Early in the day, but after
10 a.m. is good (most camera crews
start their workday at 9'a.m.) be-
cause it allows plenty of time to edit
the tape before the noon news
broadcast. You may want to consider
timing the conference so that a TV
station could broadcast it live at the
noon or the evening news show. For
the conference, choose a place that
has good visuals, such as location
along the river or water body that
you have been studying, at your
headquarters where volunteers can
be shown working in the background
or at a recognition gathering for
volunteers.
Other publicity. Be creative in
getting your report and message out.
Try writing op-ed articles for local
or statewide papers, writing letters to
the editor, producing radio feeds (a
-------
200 I MANAGING AND PRESENTING VOLUNTEER DATA
recording of the group's leader
played over the phone to a radio
station), issuing media advisories,
and even advertising in publications.
For more help on getting your
message across, consult the refer-
ences cited below.
References and Further Reading
Byrnes, J. 1994. How Citizen Monitoring
Data Became a Part of Community Life.
Volunteer Monitor. 6(1):17.
Ely, E. 1992. (ed.) Monitoring for Advo-
cacy. Volunteer Monitor. 4(1) Spring
1992.
Ely, E. 1992. (ed.) Building Credibility.
Volunteer Monitor. 4(2) Fall 1992.
Ely, E. 1994. Putting Data to Use. Volun-
teer Monitor. 6(1):11.
Ely, E. 1995. (ed.) Managing and Present-
ing Your Data. Volunteer Monitor. 7(1)
Spring 1995.
Sweeney, K. 1989. The Media Director:
Patagonia's Guide for Environmental
Groups, Ventura, CA.
Tufte, E.R. 1991. The Visual Display of
Quantitative Information, Graphics
Press, Cheshire, Connecticut.
-------
APPENDICES I 201
-------
202 I APPENDICES
Appendix A:
Glossary
accuracy - a measure of how close re-
peated trials are to the desired target.
acidity - a measure of the number of free
hydrogen ions (H+) in a solution that
can chemically react with other sub-
stances.
alkalinity - a measure of the negative ions
that are available to react and neutralize
free hydrogen ions. Some of most
common of these include hydroxide
(OH"), sulfate (SO4), phosphate (PO4),
bicarbonate (HCO3) and carbonate
(C03)
ambient - pertaining to the current environ-
mental condition.
assemblage - the set of related organisms
that represent a portion of a biological
community (e.g., benthic macroinverte-
brates).
benthic - pertaining to the bottom (bed) of
a water body.
biochemical oxygen demand (BOD) - the
amount of oxygen consumed by
microorganisms as they decompose
organic materials in water.
biological criteria - numerical values or
narrative descriptions that depict the
biological integrity of aquatic commu-
nities in that state. May be listed in
state water quality standards.
buret - a graduated glass tube used for
measuring and releasing small and
precise amounts of liquid.
channel - the section of the stream that
contains the main flow.
channelization - the straightening of a
stream; this often is a result of human
activity.
chemical constituents - chemical compo-
nents that are part of a whole.
cobble - medium-sized rocks (2-10 inches)
that are found in a stream bed.
combined sewer overflow (CSO) - sewer
systems in which sanitary waste and
stormwater are combined in heavy
rains; this is especially common in
older cities. The discharge from CSOs
is typically untreated.
community - the whole of the plant and
animal population inhabiting a given
area.
culvert - man-made construction that
diverts the natural flow of water.
d-frame net - a fine mesh net that is
attached to a pole and used for sam-
pling. It resembles a butterfly net.
deionized water - water that has had all of
the ions (atoms or molecules) other
than hydrogen and oxygen removed.
designated uses - state-established desir-
able uses that waters should support,
such as fishing, swimming, and aquatic
life. Listed in state water quality
standards.
dissolved oxygen (DO) - oxygen dissolved
in water and available for living
organisms to use for respiration.
distilled water - water that has had most of
its impurities removed.
effluent - wastewater discharge.
dredge - to remove sediments from the
stream bed to deepen or widen the
channel.
ecoregion - geographic areas that are
distinguished from others by ecological
characteristics such as climate, soils,
geology, and vegetation.
embeddedness - the degree to which rocks
in the streambed are surrounded by
sediment.
emergent plants - plants rooted underwa-
ter, but with their tops extending above
the water.
Erlenmeyer flask - a flask having a wide
bottom and a smaller neck and mouth
that is used to mix liquids.
-------
APPENDICES I 203
eutrophication - the natural and artificial
addition of nutrients to a waterbody,
which may lead to depleted oxygen
concentrations. Eutrophication is a
natural process that is frequently
accelerated and intensified by human
activities.
floating plants - plants that grow free
floating, rather that being attached to
the stream bed.
flocculent (floe) - a mass of particles that
form into a clump as a result of a
chemical reaction.
glide/run - section of a stream with a
relatively high velocity and with little
or no turbulence on the surface of the
water.
graduated cylinder - a cylinder used to
measure liquids that is marked in units.
gross morphological features - large
obvious identifying physical character-
istics of an organism.
headwaters - the origins of a stream.
hypoxia - depletion of dissolved oxygen in
an aquatic system.
impairment - degradation.
impoundment - a body of water contained
by a barrier, such as a dam.
inert - not chemically or physically active.
kick-net - a fine mesh net used to collect
organisms. Kick-nets vary in size, but
generally are about three feet long and
are attached to two wooden poles at
each end.
land uses - activities that take place on the
land, such as construction, farming, or
tree clearing.
macroinvertebrate - organisms that lack a
backbone and can be seen with the
naked eye.
NPDES - National Pollutant Discharge
Elimination System, a national program
in which pollution dischargers such as
factories and sewage treatment plants
are given permits to discharge. These
permits contain limits on the pollutants
they are allowed to discharge.
orthophosphate - inorganic phosphorus
dissolved in water.
outfall - the pipe through which industrial
facilities and wastewater treatment
plants discharge their effluent (waste-
water) into a waterbody.
permeable - porous
pH - a numerical measure of the hydrogen
ion concentration used to indicate the
alkalinity or acidity of a substance.
Measured on a scale of 1.0 (acidic) to
14.0 (basic); 7.0 is neutral.
phosphorus - a nutrient that is essential for
plants and animals.
photosynthesis - the chemical reaction in
plants that utilizes light energy from
the sun to convert water and carbon
dioxide into simple sugars. This
reaction is facilitated by chlorophyll.
pipet - an eye dropper-like instrument that
can measure very small amounts of a
liquid.
pool - deeper portion of a stream where
water flows slower than in neighbor-
ing, shallower portions.
precision - a measure of how close re-
peated trials are to each other.
protocol - defined procedure.
reagent - a substance or chemical used to
indicate the presence of a chemical or
to induce a chemical reaction to
determine the chemical characteristics
of a solution.
riffle - shallow area in a stream where
water flows swiftly over gravel and
rock.
riparian - of or pertaining to the banks of a
body of water.
riparian zone - the vegetative area on each
bank of a body of water.
riprap - rocks used on an embankment to
protect against bank erosion.
run/glide - see glide/run.
saturated - inundated; filled to the point of
capacity or beyond.
sheen - the glimmering effect that oil has
on water as light is reflected more
sharply off of the surface.
-------
204 I APPENDICES
sieve bucket - a bucket with a screen
bottom that is used to wash macroin-
vertebrate samples and to remove
excess silt and mud.
silviculture - forestry and the commercial
farming of trees.
submergent plants - plants that live and
grow fully submerged under the water.
substrate - refers to a surface. This
includes the material comprising the
stream bed or the surfaces which plants
or animals may attach or live upon.
taxon (plural taxa) - a level of classification
within a scientific system that catego-
rizes living organisms based on their
physical characteristics.
taxonomic key - a quick reference guide
used to identify organisms. They are
available in varying degrees of com-
plexity and detail.
titration - the addition of small, precise
quantities of a reagent to a sample until
the sample reaches a certain endpoint.
Reaching the endpoint is usually
indicated by a color change.
tolerance - the ability to withstand a
particular condition - e.g. pollution
tolerant indicates that ability to live in
polluted waters.
tributaries - a body of water that drains
into another, typically larger, body of
water.
turbidity - murkiness or cloudiness of
water, indicating the presence of some
suspended sediments, dissolved solids,
natural or man-made chemicals, algae,
etc.
volumetric flask - a flask that holds a
predetermined amount of liquid.
water quality criteria - maximum concen-
trations of pollutants that are accept-
able, if those waters are to meet water
quality standards. Listed in state water
quality standards.
water quality standards - written goals for
state waters, established by each state
and approved by EPA.
watershed - the area of land drained by a
particular river or stream system.
-------
APPENDICES I 205
Appendix B:
Scientific Supply
Houses
This is a partial list of chemical and
scientific equipment supply companies
from which to purchase equipment for a
volunteer monitoring program.
Aquatic Research Instruments
P.O. Box 2214
Seattle, WA 98111
(206) 789-0138
Water samplers, plankton nets, Surber
samplers, Hess samplers, drift nets,
calibrated lines, armored thermom-
eters, BOD bottles.
Ben Meadows
3589 Broad Street
Atlanta, GA 30341
(800)241-6401
Waders, rubber boots, field water test
equipment, kick nets, dip nets, wash
buckets, forceps.
Carolina Biological Supply
Company
2700 York Court
Burlington, NC 27215-3398
(800) 334-5551
Flexible arm magnifiers, hand lenses,
forceps, kick nets, microscopes,
reagents, educational materials, live
and mounted specimens for instruc-
tion.
Cole Palmer Instruments, Inc.
625 East Bunker Court
Vernon Hills, IL 60061
(800) 323-4340
Lab equipment, field water test equip-
ment, microscopes.
Chemetrics
Route 28
Calverton, VA 22016-0214
(800) 356-3072
Water testing mini-kits for field
analysis of dissolved oxygen, nitrate,
nitrite, ammonia, phosphates, chlo-
rine, sulfur, manganese, etc.
Consolidated Plastics
8181 DarrowRoad
Twinsburg, OH 44087
(800)362-1000
Sampling trays, buckets, nalgene
bottles, garbage bags, Whirl Paks ®.
Dazor Manufacturing Corp.
4483 Duncan Ave.
St. Louis, MO 63110
(800) 245-9103 .
Illuminated magnifiers.
Fisher Scientific
711 Forbes Ave.
Pittsburgh, PA 15219-4785
(800)766-7000
Lab equipment, sample bottles, sieves,
reagents, incubators, water test
equipment, Whirl Paks ®.
Hach Equipment Company
P.O. Box 329
Loveland, CO 80539-0389
(800)227-4224 .
Field and lab water testing equipment,
spectrophotometers, incubators, water
sampling kits, fecal coliform sampling
supplies, reagents, educational
materials.
Hydrolab Corporation
P.O. Box 50116
Austin, TX 78763
(800) 949-3766
Water monitoring equipment and
supplies.
-------
206 I APPENDICES
LaMotte
P.O. Box 329
Chestertown, MD 21620
(800) 344-3100
Water sampling kits, field and lab
water testing equipment, Secchi disks,
water samplers, armored thermom-
eters, calibrated lines, plankton nets,
kicknets, educational materials.
Lawrence Enterprises
P.O. Box 344
Seal Harbor, ME 04675
(207) 276-5746
Transparency tubes, view scopes,
Secchi disks, water samplers, kick
nets, sieve buckets.
Millipore Corporation
397 Williams Street
Marlborough, MA 01752
(800) 645-5476
Fecal coliform testing supplies (com-
plete sterile water filtration system),
membrane filters, sterile pipette, petri
dishes, sterile media, other water
sampling equipment and lab supplies,
incubators, Whirl Paks ®.
Nalge Company
P.O. Box 20365
Rochester, NY 14602
Fecal coliform testing supplies,
disposal fecal coliform filtration
systems, membrane filters, sterile
pipettes, petri dishes, incubators,
Whirl Paks®.
Nichols Net and Twine, Inc.
200 Highway 111
Granite City, IL 62040
(618)797-0211
Kick nets.
Ohmicron
375 Pheasant Run
Newtown, PA 18940
(800) 544-8881
Immunoassay kits for pesticides, other
contaminants.
Thomas Scientific Company
99 High Hill Road at 1-295
P.O. Box 99
Swedesboro, NJ 08085-0099
(609) 345-2100
Lab equipment, sample bottles, sieves,
reagents, incubators, water test
equipment, Whirl Paks ®.
VWR Scientific
1230 Kennestone Circle
Marietta, GA 30066
(800) 932-5000
Glassware, labeling tape, sample
vials, lab equipment, incubators,
reagents, Whirl Paks ®.
Wards Biological and Lab Supplies
P.O. Box 92912
Rochester, NY 14692-9012
(800) 635-8439
Alcohol lamps, balances, microscopes,
sample trays, goggles, rubber stop-
pers, autoclaves, spectrophotometers,
incubators, petri dishes, sterile pi-
pettes, glassware, educational materi-
als, live and mounted specimens for
instruction.
Wildco Wildlife Supply Company
301 Cass Street
Saginaw, MI 48602
(517) 799-8100
Kick nets, wash buckets, field biologi-
cal sampling equipment.
YSI Incorporated
1725 Brannum Lane
Yellow Springs, Ohio 45387
(513) 767-7241
Water quality monitoring equipment
supplies.
-------
APPENDICES I 207
Appendix C:
Determining Latitude
and Longitude
There are many ways that monitoring
groups identify and describe the location of
sampling sites. Commonly, monitoring
sites are described by stream name and
geographic location, such as Volunteer
Creek at Oak Road or Volunteer Creek
behind the picnic area in Volunteer Park.
Often these description are accompanied by
an assigned station number (i.e. VC001,
VC002). Some programs use river miles—
the distance from the sampling station to
the stream's mouth—as an additional
identifier.
Maps, in many forms, are also typically
used to help identify sites. These include
road maps, state/county maps, aerial maps,
hand-drawn site maps, and topographic
maps. Section 3.1 in Chapter 3, Watershed
Survey Methods, discusses the various types
of maps used by monitoring programs and
provides information on obtaining topo-
graphic maps from the U.S. Geological
Survey (USGS).
The most accurate way to identify
sampling locations is by determining their
latitude and longitude. Any volunteer
program that wishes to have its data used
by state, local, or federal agencies, or that
plans to enter its data into a Geographic
Information Systems (GIS) either now or in
the future, must provide latitudes and
longitudes for its sampling locations. EPA's
STORET water quality database, for
example, requires latitude/longitude
information before any data can be entered.
Section 4.1 in Chapter 4, Macroinverte-
brates and Habitat, briefly describes using
a global positioning system (GPS) to
Latitude and Longitude
Latitude and longitude are defined and measured in degrees (°),
minutes ('), and seconds ("). There are 60 seconds in a minute and
60 minutes in a degree of latitude and longitude.
Latitude (lat) is the angular distance of a particular location
north or south from the equator. Latitude lines are called parallels.
Longitude (long) is the angular distance of a particular location
east or west of some prime meridian (usually Greenwich, En-
gland). Longitude lines are called meridians.
determine latitude and longitude. This
hand-held tool is used in the field and
receives signals from orbiting satellites to
calculate the lat/long coordinates of the
user.
New tools are continuously developing
to help you locate your sites. For example,
EPA's Surf Your Watershed web page ties
in with the U.S. Geological Survey's
Names Information System to provide
latitude and longitude information for
locations throughout the U.S. These
locations include bridges, schools, rivers,
parks, and more. Visit this feature of Surf
Your Watershed at www.epa.gov/surf/
surf_search.html for more information.
Latitude and longitude can also be
calculated manually. To do this, you will
need a topographic map, a metric ruler, and
a calculator. A worksheet for calculating
latitude and longitude based on the EPA
Region 10 Streamwalk protocol is pre-
sented below.
-------
208 I APPENDICES
Worksheet for Calculating Latitude and Longitude
7.5 x 15 Minute Series
57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75
47° 37' 30"
72
71
70
68
67
66
64
63
62
122° 15' 12'
7130" 5'
Read Longitude
To Determine Latitude:
1. Look at the right side (upper or lower corner) under the map name,
or the second of two numbers separated by "x" to find the height
scale (latitude) of the topo map.
If "7.5 Minute Series," enter 450
If "15 Minute Series," enter 900
If "7.5 x 15 Minute Series," enter 450
Your
Work
Example
450
-------
APPENDICES I 209
Your Calculation Example
2. Using the ruler, measure the length of your map (exclude the map
borders) north to south in centimeters (cm).
3. Divide #1 by #2 to the nearest whole number.
4. Enter the latitude located in the map's edge closest to your site.
10cm
450 - 10 = 45
47° 30'
5. Using the ruler, measure from your site straight down, to the
bottom of the map (in centimeters).
4.8cm
6. Multiply #5 by #3 to the nearest whole number.
4.8x45 = 216
7. Determine how many times 60 goes into #6 completely and what
is left as the remainder (don't use a calculator for this). These
answers will become the minutes and seconds of the latitude.
60 goes into 216
completely 3 times
with 36 left over.
(3 x 60 = 180;
216-180 = 36).
8. Convert these numbers to minutes and seconds. Minutes are
equal to the whole number determined in #7, or the number of
times 60 goes into #6 completely. In other words, your whole
number after the division in the previous step is the number of
minutes. Seconds are equal to what is left (remainder) after the
division in #7.
3 minutes and
36 seconds =
3' 36"
9. Determine the latitude of your site by adding #4 to #8.
47° 30' + 3' 36"
= 47° 33' 36"
To Determine Longitude:
1. Look at the right side (upper or lower corner) under the map name,
or the second of two numbers separated by "x" to find the width
scale (longitude) of the topo map.
If "7.5 Minute Series," enter 450
If "15 Minute Series," enter 900
If "7.5 x 15 Minute Series," enter 900
900
-------
210 I APPENDICES
Your Calculation Example
2. Using the ruler, measure the width of your map (exclude the map
borders) east to west in centimeters (cm).
10cm
3. Divide #1 by #2 to the nearest whole number.
900 ^ 10 = 90
4. Enter the longitude located in the map's lower right hand corner.
122° 00'
5. Using the ruler, measure from your site straight across, to the right
hand side of the map (in centimeters).
3.7cm
6. Multiply #5 by #3 to the nearest whole number.
3.7 x 90 = 333
7. Determine how many times 60 goes into #6 completely and what
is left as the remainder (don't use a calculator for this). These
answers will become the minutes and seconds of the longitude.
(The longitude degrees are #4.)
60 goes into 333
completely 5 times
with 33 left over.
(5 x 60 = 300;
333 - 300 = 33).
8. Convert to these numbers to minutes and seconds. Minutes are
equal to the whole number determined in #7, or the number of
times 60 goes into #6 completely. In other words, your whole
number after the division in the previous step is the number of
minutes. Seconds are equal to what is left (remainder) after the
division in #7.
5 minutes and
33 seconds
= 5' 33"
9. Determine the longitude of your site by adding #4 to #8.
122° 00'
+ 5' 33"
== 122° 5' 33"
------- |