&EPA
United States
Environmental Protection
Agency
Field Operations Manual for
Assessing the Hydrologic
Permanence and Ecological
Condition of Headwater Streams
RESEARCH AND DEVELOPMENT
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EPA600/R-06/126
October 2006
www.epa.gov
Field Operations Manual for
Assessing the Hydrologic
Permanence and Ecological
Condition of Headwater Streams
Prepared by
Ken M. Fritz
Brent R. Johnson
David M. Walters
Ecosystems Research Branch
Ecological Exposure Research Division
National Exposure Research Laboratory
Office of Research and Development
U.S. Environmental Protection Agency
Notice:
Although this work was reviewed by EPA and approved for publication, it may not
necessarily reflect official Agency policy. Mention of trade names and commercial
products does not constitute endorsement or recommendation for use.
U.S. Environmental Protection Agency
Office of Research and Development
Washington, DC 20460
11
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NOTICE
The information in this document has been subjected to the Agency's peer and administrative
review requirements and has been approved for publication as an EPA document.
Mention of trade names or commercial products does not constitute endorsement or
recommendation for use.
The correct citation for this document is:
Fritz, K.M., Johnson, B.R., and Walters, D.M. 2006. Field Operations Manual for
Assessing the Hydrologic Permanence and Ecological Condition of Headwater Streams.
EP A/600/R-06/126. U.S. Environmental Protection Agency, Office of Research and
Development, Washington DC.
in
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PREFACE
The mission of the Ecological Exposure Research Division (EERD), National Exposure
Research Laboratory (NERL), United States Environmental Protection Agency (USEPA) is to
improve the scientific basis for understanding, measuring, and protecting biological integrity so
that USEPA and other resource agencies can make sound, defensible environmental decisions.
Our research is primarily focused on the development, evaluation, and implementation of new
methods to assess ecosystem condition, to evaluate biotic responses to environmental stressors,
and to predict future vulnerability of natural populations, communities and ecosystems.
This document originated from a research project, the Headwater Intermittent Streams Study
(HISS), funded through the USEPA's Regional Methods (RM) Program (overseen by the
Biological Advisory Committee and supported by the USEPA, Office of Science and Policy).
The purpose of RM is to support development of methods needed by EPA regions, states and
tribes to meet their monitoring and enforcement objectives. The widespread need for
standardized methods for assessing headwater streams is apparent from the sponsorship and
participation by USEPA Regional offices (1, 2, 3, 4, 5, 8, 9 and 10) and several state offices
therein. The initial development of the methods was in forested headwater streams located in
Indiana, Kentucky, and Ohio over 2003 and 2004. Following training workshops, state and
regional teams used the methods to collect data from forested headwater streams in Illinois, New
Hampshire, New York, Vermont, West Virginia, and Washington. This manual is a product of
the working collaboration among EERD, regional, and state scientists. We hope that the
methods described in this manual will be useful to individuals and organizations interested in
monitoring and protecting headwater streams.
Florence Fulk
Acting Director
Ecological Exposure Research Division
IV
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TABLE OF CONTENTS
Notice iii
Preface iv
Table of Contents v
List of Tables vii
List of Figures viii
Acknowledgements xiv
Acronyms and Abbreviations xv
1 Introduction 1
1.1 Purpose 1
2 Factors Influencing Study Design 10
2.1 Study design for comparing across stream reaches with varying hydrologic
permanence 18
3 Physical Habitat Characterization 25
3.1 Designating hydrologic condition for stream reaches 29
3.2 Continuous monitoring of hydrologic condition 35
3.3 Identifying the channel head 40
3.4 Identifying channel headcuts 43
3.5 Measuring channel sinuosity 46
3.6 Designating habitat units 48
3.7 Measuring channel slope 51
3.8 Measuring water depth 54
3.9 Measuring wetted width 58
3.10 Measuring basic channel geomorphology 60
3.11 Measuring water velocity 64
3.12 Measuring discharge 70
3.13 Measuring depth to bedrock and groundwater table 75
3.14 Gravimetrically measuring streambed sediment moisture 79
3.15 Characterizing the size distribution of streambed sediments 82
3.16 In situ water chemistry measurements 87
3.17 Measuring riparian canopy cover 92
4 Biological Sampling 96
4.1 Sampling the bryophyte assemblage 99
4.2 Sampling the epilithic algal assemblage 105
4.3 Visual and tactile assessment of algal cover Ill
4.4 Sampling the benthic invertebrate assemblage 114
4.5 Surveying the amphibian assemblage 123
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5 Appendix Field Forms 130
VI
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LIST OF TABLES
Table 3-1 Modified Wentworth scale for sediment particle size classes. Bold-faced
numbers indicate values to be entered on field forms 85
Table 3-2 Data Quality Objectives (DQO) for in situ water chemistry
measurements 89
Table 4-1 Algal Cover Index (ACT) scores and their associated characteristics 112
vn
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LIST OF FIGURES
Figure 1-1 Portions of a 1:100 000 (A; Ironton 30 x 60 minute quadrangle) and a
1:24 000 (B; Gallia 7.5 minute quadrangle) scale United States
Geological Survey (USGS) topographic maps illustrating the upper
reaches of Buffalo Creek in Wayne National Forest (Lawrence and
Gallia Counties, OH). Black circles and associated letters mark
corresponding points on both maps. Black horizontal bars represent 1
km. Buffalo Creek at "a" is a second-order stream on the 1:100 000
map, but is a third-order stream on the 1:24 000 map. Likewise, Buffalo
Creek at "b" is considered a first-order stream on the 1:100 000 map, but
is a second-order stream on the 1:24 000 map. The point marked "c" is
shown as a first-order stream on the 1:24 000 map, but is not designated
as a stream on the 1:100 000 map. The number of first-order streams
shown upstream of "a" on the 1:100 000 map is two, whereas the 1:24
000 map has five. Field surveys of this drainage would likely find >
10X first-order streams upstream of "a" 2
Figure 1-2 Portions of 1:15 840 United States Department of Agriculture (USD A)
National Resources Conservation Service (NRCS) maps from Lawrence
and Gallia Counties, OH illustrate the upper reaches of Buffalo Creek in
Wayne National Forest (McCleary and Hamilton 1998). Green circles
and associated letters mark corresponding points on maps in Figure 1-1.
The yellow circles highlight the delineated stream origins. Black
horizontal bars represent 0.5 mi (0.805 km). Buffalo Creek at "a" is a
fourth-order stream, at "b" it is considered a third-order stream, and at
"c" it is shown as a second-order. The number of first order streams
shown upstream of "a" is 41 3
Figure 2-1 Relationship between sample size and standard error estimations
assuming proportions are equal among populations 11
Figure 2-2 Types of disturbance (solid) and responses (dashed) in streams: pulse
(a), press (b), ramp (c) and stepped (d). Based on figures from Lake
(2003) and Boulton (2003) 14
Figure 2-3 Map highlighting position of headwater channels within the watershed
of Falling Rock Branch, KY. Yellow represents boundary of watershed,
blue represents "blue line" designation on the 1:24 000 USGS
topographic map (Noble 7.5 minute quadrangle, Breathitt County, KY),
and red represent headwater channels not shown on the topographic
map 20
Figure 2-4 Schematic showing suboptimal and preferred longitudinal positioning of
sites along headwater channels to maximize the range of hydrologic
permanence across study sites. Hypothetical drainage areas are shown
to further illustrate spatial hierarchy 21
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Figure 2-5 Map showing positioning of sites along two Indiana headwater streams
where the downstream perennial site (P) is "shared" between two
tributaries. DI = downstream intermittent; UI = upstream intermittent;
and E = ephemeral. Shading shows cumulative drainage area in
downstream direction 22
Figure 2-6 Schematic of headwater channels showing numerical designation and
position of study sites relative to origins of intermittent and perennial
flow 23
Figure 3-1 Appropriate location for recording hydrologic condition on page 1 of
field forms 29
Figure 3-2 Primary components of a water sensor used to continuously monitor
hydrologic condition 35
Figure 3-3 Launch dialog box for Onset Boxcar Pro 36
Figure 3-4 Desiccant packs and Onset Hobo® State data logger with jacks and
LEDs shown 36
Figure 3-5 Water sensor with 2.5 mm cable, O-ring and seat shown 37
Figure 3-6 Schematic showing assembly of stilling well and contact end of water
sensor 37
Figure 3-7 Water sensor securely attached to rebar above and below stilling well 38
Figure 3-8 Water sensor positioned for continuous monitoring of hydrologic
condition. Meter stick shown for scale 38
Figure 3-9 Drawing showing a valley hillslope (swale or hollow) relative to
channel. Valley head (A), gradual (B) and abrupt (C) channel heads are
identified. Gray areas indicate zero-order basins draining into channel
heads. Redrawn from Dietrich and Dunne (1993) 40
Figure 3-10 An abrupt channel head in Wayne National Forest, OH 41
Figure 3-11 Views from gradual channel heads in east-central Kentucky. A)
Looking upslope toward the valley head from the channel head position.
B) Looking downslope at the cascade structure of the transitional
channel 42
Figure 3-12 Longitudinal view of a headcut, (A.) Blue arrows illustrate flowpaths
that lead to undercutting, failure of the headwall, and eventually
upstream migration of the headcut; (B.) Abrupt change in summer
baseflow hydrology at a headcut 44
Figure 3-13 Portion of page 1 of field forms showing the cell for recording
presence of channel headcuts 45
Figure 3-14 Subtle headcut in Falling Rock Creek in east-central KY (looking
upstream) 45
IX
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Figure 3-15 Huge headcut (~2 m change in bed elevation) in an unnamed stream in
Athens, GA (looking upstream) 45
Figure 3-16 Headcut in Taylor Branch in south-central IN (looking downstream),
where streambed elevation at the arrow was ~ 1 m higher than
streambed below headcut at the yellow circle 45
Figure 3-17 Examples of stream channels varying in sinuosity (number of bends)
along 30-m study reaches 47
Figure 3-18 Portion of page 1 of field forms showing the cell for recording channel
sinuosity 47
Figure 3-19 Plan view of study reach (top) and picture showing series of
alternating erosional and deposit!onal habitats along a headwater stream 49
Figure 3-20 Appropriate location for recording habitat units and notes on Page 2 of
the Field Forms 50
Figure 3-21 Longitudinal section of channel 51
Figure 3-22 Plan view of study reach showing measurement locations (vertical
black tick marks) for channel slope. Flow is from right to left and the
dotted line represents the thalweg 52
Figure 3-23 Crew members measuring slope of intermittent stream 52
Figure 3-25 Portion of page 1 of field forms showing cells for percent slope values 53
Figure 3-26 Longitudinal section of channel showing position of manometer and
points of measurement to calculate slope (redrawn fro Gordan et al.
1992). Blue arrow shows direction of flow. L = horizontal length, hi =
height at the upstream end and Ii2 = height at downstream end 54
Figure 3-27 Overhead view of study reach showing locations for water depth
measurement (vertical black tick marks) along the reach thalweg (dotted
line). Water is flowing from right to left. (A.) overhead view of study
reach (B.) channel cross-section, and (C.) lateral close-up of depth 55
Figure 3-28 Appropriate location for recording longitudinal water depth
measurements on page 2 of the field forms 56
Figure 3-29 Schematic showing appropriate reading of water depth where water
surface is turbulent 57
Figure 3-30 Appropriate location for recording maximum pool depth measurement
on page 1 of the field forms 57
Figure 3-31 Channel cross-section illustrating wetted width 58
Figure 3-32 Overhead view of study reach showing measurement locations
(vertical black tick marks for wetted width. Flow is from right to left
and the dotted line represents the thalweg 58
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Figure 3-33 Appropriate location for recording wetted width measurement on page
2 of the field forms 59
Figure 3-34 Channel cross-sections showing wetted width measurements where
there is emergent cobble (A.), island (B.), and side-pool (C.) 59
Figure 3-35 Headwater stream channel showing the location of the streambed and
the banks (white arrows) 60
Figure 3-36 Plan view of study reach showing 5-m intervals. Direction of arrows
shows direction of flow, and the dotted line represents the thalweg 61
Figure 3-37 Photograph shows measurement of bankfull (BF) width and bankfull
depth 62
Figure 3-38 Appropriate location for recording bankfull (BF) width (red), bankfull
depth (blue), and flood prone area (FPA) width (black) measurements on
page 2 of the field forms 63
Figure 3-39 Photograph illustrating flood-prone area (FPA) width 63
Figure 3-40 Plan view of study reach showing measurement locations (vertical
black tick marks) for current velocity measurements. Flow is from right
to left and the dotted line represents the thalweg 65
Figure 3-41 Longitudinal section across the channel thalweg showing orientation of
the velocity probe for measurements 66
Figure 3-42 Appropriate location for recording water velocity on page 2 of the field
forms 66
Figure 3-43 Bag meter used to measure water velocity 67
Figure 3-44 Overview of study reach showing measurement locations (black tick
marks crossing the thalweg, shown as dotted line), upstream (dashed
blue lines) and downstream segment boundaries (solid red lines) for the
neutrally-buoyant procedure to measure water velocity 68
Figure 3-45 Overhead view of study reach showing leading and trailing edges of
fluoroscene plume 69
Figure 3-46 Plan view of study reach (top) showing discharge measurement cross-
section (red dashed line). Cross-section for discharge measurement
(bottom) showing measurement cells 71
Figure 3-47 Appropriate location for recording discharge and procedures used on
page 3 of field forms. Example values shown in red 72
Figure 3-48 Bag meter used to measure discharge 72
Figure 3-49 Example of a concentration curve from a slug injection. Discharge
(mV1) is the hatched area under the curve._ 74
Figure 3-50 Example of a concentration curve from a continuous injection.
Discharge (mV1) is the hatched area under the curve 74
XI
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Figure 3-51 Using sounding rod and hand sledge hammer to estimate depth to
bedrock and the groundwater table 77
Figure 3-52 Cross-section of a dry channel illustrating depth to underlying bedrock
(A) and depth to the groundwater table (B) 78
Figure 3-53 Appropriate location for recording depth to bedrock (example values
in blue) and depth to groundwater (example values in red) on page 1 of
field forms 78
Figure 3-54 Sampling sediment moisture 80
Figure 3-55 Tapping core vertically into streambed 80
Figure 3-56 Appropriate location for recording the number of sediment moisture
cores collected on page 1 of field forms 80
Figure 3-57 Example of the sediment moisture data sheet 81
Figure 3-58 Schematic of study reach illustrating thalweg (dotted line) and patch
locations for determining modal sediment particle size class. Inset
provides a close-up of a patch (overlaid) with measuring tape used in
designating patch locations longitudinally along the study reach) 84
Figure 3-59 Appropriate location for recording modal particle size data on page 2
of field forms (example from Figure 3-58 highlighted) 87
Figure 3-60 An example of an instrument inspection and calibration log sheet 89
Figure 3-61 Appropriate locations for recording in situ water quality measurements
on page 1 of field forms, example values shown in red 90
Figure 3-62 Plan view of a convex spherical densitometer, showing percent cover
values associated with intersections. Values are equivalent to the
number of squares meeting at each intersection 93
Figure 3-63 Appropriate location for recording percent canopy cover on page 1 of
field forms 93
Figure 3-64 Plan view of a convex spherical densitometer, modified for measuring
over 17 intersections (open circles) that are delimited by a "V" taped to
the convex mirror 94
Figure 4-1 Examples of a species-area curve (A) and a species gained-area curve
(B) for benthic invertebrates samples (sample area = 0.053 m2) collected
from a perennial site on Falling Rock Branch, Robinson Forest, KY.
Each point represents the mean (± 1 SE) of 100 permutations 98
Figure 4-2 Sporophyte and gametophyte generations of a moss. (Photo by Michael
Liith) 99
Figure 4-3 An epilithic moss (Musci) growing in a headwater stream 100
Figure 4-4 An epilithic liverwort (Hepaticae) growing in a headwater stream 100
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Figure 4-5 Appropriate location for recording bryophyte sample information on
page 1 of field forms 101
Figure 4-6 Appropriate location for recording algal sample information on page 1
of field forms. Example information is shown in red 106
Figure 4-7 Collecting epilithic algae from a stone within the sample delimiter 107
Figure 4-8 Equipment used to collect and preserve algal assemblage samples.
Numbers correspond to the Equipment and Supplies list Ill
Figure 4-10 Appropriate location for recording the dominant reach score for the
Algal Cover Index on page 1 of field forms 113
Figure 4-12 Coarse surface substrate set aside in basin for scrubbing 116
Figure 4-13 Sweep the hand net through the water column to collect suspended
invertebrates within the bucket area 117
Figure 4-14 Scrubbing attached invertebrates off the coarse surface substrate in the
washbasin (or sieve) 117
Figure 4-15 Carefully adding water to the wash basin before sample elutriation 117
Figure 4-16 Sample elutriation in the wash basin and pouring invertebrates and fine
detritus into the sieve 117
Figure 4-17 Carefully search the basin for heavy-bodied invertebrates that were not
transferred to the sieve 118
Figure 4-18 Washing sieve contents to one side by gentle agitation while sieve is
partially submerged 118
Figure 4-19 Sieve contents condensed for transfer to sample bag 118
Figure 4-20 Sieve contents rinsed into sample bag (over basin) using ethanol squirt
bottle 118
Figure 4-21 Appropriate location for recording invertebrate sampling information
on page 1 of field forms. Example sample information shown in red 119
Figure 4-22 Equipment used to collect and preserve benthic invertebrate samples.
Numbers correspond to the equipment and supplies list above 123
Figure 4-23 Northern tow-lined salamander, Eurycea cirrigera, from Robinson
Forest, KY: A) egg clutch; B) larva; and C) adult 124
Figure 4-24 Larval spring salamander, Gyrinophilus porphyriticus, from Robinson
Forest, KY 125
Figure 4-25 Amphibian survey field form 127
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ACKNOWLEDGEMENTS
We gratefully acknowledge the following USEPA and state agency personnel for input, support,
and encouragement throughout the development of this document: Pete Nolan, Dave McDonald,
and Tom Faber (Region 1, West Chelmsford, MA); Jim Kurtenbach (Region 2, Edison, NJ);
Maggie Passmore, Greg Pond, Frank Borsuk, and Lou Reynolds (Region 3, Wheeling, WV);
Pete Kalla (Region 4, Athens, GA); Jim Harrison (Region 4, Atlanta, GA); Ed Hammer, Dave
Pfeifer, Kerry Gerard, Holly Arrigoni, and Jonathan Burian (Region 5, Chicago, IL); Bill
Schroeder (Region 8, Denver, CO); Tina Laidlaw (Region 8, Helena, MT); Bobbi Smith (Region
9, San Francisco, CA); Pete Husby (Region 9, Richmond, CA); Gretchen Hayslip, Lil Herger,
Lorraine Edmond, Peter Leinenbach, Denise Clark, and Ben Cope (Region 10, Seattle, WA); Jeff
Bailey (WVDEP, Wheeling, WV), Bob Bode (NYSDEC, Albany, NY), Heather Pembrook and
Jim Kellogg (VTDEC, Waterbury, VT), Dan Dudley (Ohio EPA, Columbus, OH), Jeff DeShon,
Edward Moore, and Mike Bolton (Ohio EPA, Groveport, OH), Robert Davic and Paul Anderson
(Ohio EPA, Twinsburg, OH); Larry Eaton and Dave Penrose (NCDNR, Raleigh, NC); Diane
Regas, Craig Hooks, Donna Downing, Traci Nadeau, Laura Gabanski, Rose Kwok (USEPA
Office of Water (OW), Office of Wetlands, Oceans and Watersheds (OWOW), Washington,
DC); Richard Sumner (OW, OWOW, Corvallis, OR); Bill Swietlik OW, Office of Science and
Technology, Washington, DC); Allison Roy, Bill Shuster, Chris Nietch, and Keith Taulbee
(Office of Research and Development, National Risk Management Research Laboratory
(NRMRL), Cincinnati, OH).
We would also like thank following people and their associated management for facilitating the
project at various locations: Jason Taylor (The Nature Conservancy (TNC), Logan, OH); Peter
Whan (TNC, West Union, OH); Rebecca Ewing (USFS Wayne National Forest, Nelsonville,
OH); Anne Timm (USFS Hoosier National Forest, Tell City, IN); Mike Welker (USFS Shawnee
National Forest, Harrisburg, IL); Dr. Chris Barton (University of Kentucky, Lexington, KY); and
Mr. Will Marshall (University of Kentucky, Robinson Forest, Clayhole, KY).
We appreciate the critical peer-review of the manual by the following USEPA ORD scientists:
Ted Angradi and Brian Hill (National Health and Environmental Effects Laboratory (NHEERL),
Duluth, MN); Michael Griffith (National Center for Environmental Assessment, Cincinnati,
OH), Steve Reynolds (NRMRL, Ada, OK); Allison Roy (NRMRL, Cincinnati, OH); Jim
Wigington (NHEERL, Corvallis, OR); and Paul Wagner (National Exposure Research
Laboratory, Research Triangle Park, NC). We also thank the following scientists for
commenting on earlier versions of the manual: Kyle Hartman (West Virginia University), Jeff
Jack and Art Parola (University of Louisville), Frank McCormick (U.S. Forest Service), and Ben
Stout (Wheeling Jesuit University).
Special thanks to the other members of the Ecosystem Research Branch who have helped with
field work and other technical aspects: Brad Autrey, Karen Blocksom, Randy Bruins, Joe
Flotemersch, Florence Fulk, Jennifer Greenwood, Don Klemm, Chuck Lane, Pat Lewis, Michael
Moeykens, Marie Nieman, Mary Sullivan, and Lori Winters.
xiv
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ACRONYMS AND ABBREVIATIONS
°C Degrees Centigrade
|j,m Micrometer
|j,S/cm Micro-Siemens per Centimeter
ACT Algal Cover Index
AFDM Ash-Free Dry Mass
BACI Before/After and Control/Impact
BF Bankfull
cm Centimeter
cm2 Square Centimeters
Cond Conductivity
DEM Digital Elevation Model
DI Downstream Intermittent Site
DO Dissolved Oxygen
DQO Data Quality Objectives
E Ephemeral Site
EERD Ecosystem Exposure Research Division
EMAP Environmental Monitoring and Assessment Program
FCSPD Fairfax County Stormwater Planning Division
FPA Flood Prone Area
GPS Global Positioning System
HISS Headwater Intermittent Streams Study
IEI Intermountain Environmental, Inc
in Inch
km Kilometer
km2 Square Kilometers
LIDAR Light Detection and Ranging
m Meter
m2 Square Meters
mV1 Cubic Meters per Second
m/s Meters per Second
mi Miles
mi2 Square Miles
mg/1 Milligrams per Liter
ml Milliliter
mm Millimeter
NaCl Sodium Chloride
NAWQA National Water Quality Assessment
NCDNR North Carolina Department of Natural Resources
NCDWQ North Carolina Division of Water Quality
NHEERL National Health and Environmental Effects Laboratory
NRCS National Resources Conservation Service
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NRMRL National Risk Management Research Laboratory
NYSDEC New York State Department of Environmental Conservation
Ohio EPA Ohio Environmental Protection Agency
OW Office of Water
OWOW Office of Wetlands, Oceans and Watersheds
P Perennial Site
PC Personal Computer
PDA Personal Digital Assistant
PVC Polyvinyl chloride
RHAF Rapid Habitat Assessment Form
RM Regional Methods
SE Standard Error
SVL Snout-Vent Length
Temp Temperature
TNC The Nature Conservancy
UI Upstream Intermittent Site
USDA United States Department of Agriculture
USEPA United States Environmental Protection Agency
USFS United States Forest Service
USGS United States Geological Survey
UTM Universal Transverse Mercator
VTDEC Vermont Department of Environmental Conservation
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1 INTRODUCTION
1.1 Purpose
The purposes of this manual are to: 1)
document procedures that were developed and
used by the United States Environmental
Protection Agency's (USEPA) Ecosystem
Exposure Research Division (EERD) for the
assessment of the physical and biological
characteristics of headwater streams; and 2)
provide a catalog of procedures to other
groups with an interest in headwater stream
assessment. Earlier EPA field operations
manuals for running waters have focused on
larger systems, including wadeable streams,
non-wadeable rivers, and Great Rivers (e.g.,
Barbour et al. 1999, Lazorchak et al. 1998,
2000, Angradi 2006). There is a growing
interest in headwater streams because human
activities (e.g., road building, stormwater
management) frequently intersect these
widespread waterbodies. There is also
considerable legal debate regarding extent of
jurisdictional waters under the Clean Water
Act and the role or nexus of various types of
headwater streams to the integrity of
downstream interstate waters (Nadeau and
Rains in press). Some states, like North
Carolina and Ohio, have already began to
initiate headwater stream classification
methods for regulatory purposes (Ohio
Environmental Protection Agency 2002, N.C.
Division of Water Quality 2005).
This document provides methods specifically
designed for assessing the hydrologic
permanence and ecological condition of
headwater streams. A universal, spatially-
explicit definition of a headwater stream is
lacking because stream size and drainage area
varies with surrounding topography and
geographic location. Regardless, headwater
streams are important because they are the
origins of the stream network and have unique
ecological characteristics that separate them
from larger, downstream waterbodies.
What are headwater streams?
Stream order is a measure of stream position
within a drainage network system (Horton
1932, Strahler 1945, Shreve 1966).
Headwater streams are typically considered to
be first- and second-order streams (Gomi et al.
2002, Meyer and Wallace 2001), meaning
streams that have no upstream tributaries (i.e.,
"branches") and those that have only first-
order tributaries, respectively. Use of stream
order to define headwater streams is
problematic because stream-order
designations vary depending upon the
accuracy and resolution of the stream
delineation (Mark 1983, Hanson 2001). Lack
of agreement among maps with different
mapping resolution is common when
identifying headwater stream, determining
stream order, and determining total stream
(Figures 1-1 and 1-2). The designation of the
mainstem (central tributary) origin is typically
similar between the 1:100 000 and 1:24 000
scale maps. However, the 1:24 000 maps
delineate more lateral tributaries (Figures 1-
1A and 1-1B) and this can result in substantial
differences of headwater extent. The total
stream length within the Coweeta Creek
watershed (16.3 km2) in western North
Carolina on a 1:500 000 scale map was only
3% of the length shown on a 1:24 000 scale
map (Meyer and Wallace 2001). The smallest
headwater streams are not designated as
channels on topographic maps and may be
difficult to discern in aerial photographs.
Thus, stream-order designations based on
maps are typically underestimated (Hughes
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^
Figure 1-1 Portions of a 1:100 000 (A; Ironton 30 x 60 minute quadrangle) and a 1:24 000 (B; Gallia 7.5 minute quadrangle)
scale United States Geological Survey (USGS) topographic maps illustrating the upper reaches of Buffalo Creek in Wayne
National Forest (Lawrence and Gallia Counties, OH). Black circles and associated letters mark corresponding points on both
maps. Black horizontal bars represent 1 km. Buffalo Creek at "a" is a second-order stream on the 1:100 000 map, but is a
third-order stream on the 1:24 000 map. Likewise, Buffalo Creek at "b" is considered a first-order stream on the 1:100 000
map, but is a second-order stream on the 1:24 000 map. The point marked "c" is shown as a first-order stream on the 1:24
000 map, but is not designated as a stream on the 1:100 000 map. The number of first-order streams shown upstream of "a"
on the 1:100 000 map is two, whereas the 1:24 000 map has five. Field surveys of this drainage would likely find > 10X first-
order streams upstream of "a".
2
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2\ ¥
Figure 1-2 Portions of 1:15 840 United States Department of Agriculture (USDA) National
Resources Conservation Service (NRCS) maps from Lawrence and Gallia Counties, OH
illustrate the upper reaches of Buffalo Creek in Wayne National Forest (McCleary and
Hamilton 1998). Green circles and associated letters mark corresponding points on maps
in Figure 1-1. The yellow circles highlight the delineated stream origins. Black horizontal
bars represent 0.5 mi (0.805 km). Buffalo Creek at "a" is a fourth-order stream, at "b" it is
considered a third-order stream, and at "c" it is shown as a second-order. The number of
first order streams shown upstream of "a" is 41.
and Omernik 1983), prompting some
investigators to characterize such streams as
zero-order streams (e.g., Brown et al. 1997).
Most "blue line" designations on topographic
maps are not based on field studies, but are
"drawn to fit a rather personalized aesthetic"
of the cartographer (Leopold 1994) or drawn
with standards that exclude a proportion of
headwater channels (Drummond 1974).
Moreover, large scale aerial and satellite
image databases (e.g., 30-m DEM) are
typically too coarse to accurately identify
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most headwater channels, particularly in
forested regions. Further development and
more affordable application of Light Detection
and Ranging (LIDAR) mapping technology
provides the most promise for remotely
recording the location and extent of headwater
streams (e.g., Jarnagin and Jennings 2005). In
addition, further work in understanding factors
contributing to the evolution of stream
channels will be useful for predicting the
spatial distribution of headwater streams
across the landscape (e.g., Montgomery and
Dietrich 1988, 1992).
Headwater streams as monitoring units
Headwater streams are useful monitoring units
owing to their extent (i.e., widespread and
abundant), spatial scale and landscape
position. Replicate streams of given
treatments (e.g., types of land use/cover) and
reference conditions, are more available for
headwater streams because of their abundance
across the landscape and relatively small
watershed areas. Experimental studies are
also more feasible (and ethically acceptable)
in headwater streams and watersheds because
they are easier to modify or perturb than
downstream waterbodies (e.g., Likens et al.
1970, Wallace etal. 1999). Assessments of
headwater streams can provide better
resolution to diagnose cause and effect
because they drain smaller areas with less land
use heterogeneity than their larger
counterparts. Flow of water from land to
headwater channels is relatively short
compared to larger rivers; therefore responses
to land changes may be more rapidly detected.
Because headwater streams have narrower
widths and shallower depths than larger
streams and rivers, a larger proportion of
water flowing through headwater channels is
directly contacting (and exchanging water and
solutes with) the stream bed and banks at a
given moment. Biogeochemical processes
(e.g., denitrification) and biotic densities are
often higher in the saturated sediments of beds
and banks than in the water column. This
increased wetted area to water volume ratio
therefore suggests that headwater channels
may strongly influence downstream water
quality. Lastly, because headwater streams
represent the dominant interface between
surrounding landscapes and downstream
surface waters, further understanding of the
structure and function of headwater streams
will improve our ability to protect all water
bodies.
Headwater streams and drying
One of the most distinctive and ecologically
influential characteristics of many headwater
streams is natural drying. In contrast to
perennial or permanent streams that maintain
continuous surface flow throughout most
years, temporary streams (e.g., intermittent,
ephemeral) have a recurrent dry phase(s)
(Comin and Williams 1994, Uys and O'Keefe
1997, Williams 2006). Not to be confused
with temporary waters are aestival water
bodies (more commonly used to describe
ponds than streams, but see Johansson and
Nilsson 1994). Aestival habitats are
characterized by being shallow and
permanent, but freeze completely during the
winter (Daborn and Clifford 1974).
Temporary streams are the dominant form of
running waters in arid and semiarid regions
(Zale et al. 1989, Dodds 1997, Gasith and
Resh 1999. Nanson et al. 2002), but are also
common in temperate and tropical areas (e.g.,
Clifford 1966, Chapman and Kramer 1991,
Delucchi 1988, Feminella 1996). Regardless
of climatic region, headwater streams are
more prone to drying than larger streams
because they have smaller drainage areas for
capturing recharge and generally have higher
topographic elevation (McMahon and
Finlayson 2003, Rivenbark and Jackson 2004,
Svec et al. 2005). The rate of drying, and
predictability, duration, and frequency of dry
-------
periods vary with geographic setting and
annual precipitation.
Variation in the temporal aspects of drying has
been categorized by various classification
schemes of temporary streams (Abell 1984,
Poff and Ward 1989, Uys and O'Keefe 1997).
Intermittent streams are typically identified as
those that dry seasonally. During the dry
season(s), frequently compounded by high
evapotranspiration of watershed vegetation,
the groundwater table may drop below the
elevation of the streambed causing the stream
to dry (Williams 2006). Ephemeral (or
episodic) streams are usually dry except for
several days immediately following
precipitation. Surface flow in ephemeral
channels is derived from surface runoff and
shallow throughflow. Rather than having
distinct, rigid boundaries, stream reaches
classified as perennial, intermittent, and
ephemeral may more accurately be described
as dynamic zones within stream networks.
The length or extent of these zones may be
highly variable and is dictated by multiple
factors (e.g., annual precipitation,
evapotranspiration, land-use practices). The
variable source area concept describes the
dynamic zones as the expansion and
contraction of flow within forested headwater
systems (Hewlett and Hibbert 1967).
Increases in discharge within small watersheds
following a rain storm are rarely equivalent to
the volume of rain fallen on the watershed.
Much of the rain infiltrates into the soil and
displaces subsurface water (already saturating
the watershed) downslope into channels (i.e.,
throughflow or translatory flow). When this
subsurface flow exceeds the capacity of the
soil to transmit it downslope, water will be
seen at the streambed surface and the wetted
channel will extend upslope. Using a
conservative tracer (NaCl) Genereux et al.
(1993) measured the spatial and temporal
variation in flow generation within a small
watershed in Tennessee. They determined
that two downstream, perennial springs
generated most of the flow during late
summer, but as discharged increased, flow
was predominantly generated from upstream,
temporary reaches.
The natural process of drying causes changes
in physical and chemical conditions (e.g., loss
of wetted habitat, reduced dissolved oxygen),
which can exclude some species while
allowing others to thrive (Boulton et al. 2000).
Temporary streams may, therefore, harbor
communities containing mixtures of unique
endemics (i.e., locally distributed species) and
opportunist cosmopolitans (i.e., widespread
species). The biotic community will vary
among temporary waters with duration of
hydroperiod (Williams 1996) and timing of
the hydrologic cycle (Boulton and Lake 1992,
Fritz and Dodds 2002). The hydrologic
permanence (duration and frequency of
continuous surface flow) of headwater streams
must be understood to avoid confounding
effects of natural drying when assessing the
ecological integrity or condition. Different
ecological expectations are likely needed
when assessing condition of perennial and
temporary streams. Although the methods
described in this manual were used to identify
hydrologic regimes primarily in forested
headwater streams, some of the methods can
also serve to quantify the ecological integrity
of non-forested headwater streams.
Organization of the manual
This manual is divided into three sections: 1)
Assessment Design and Site Selection, 2)
Physical Habitat Characterization, and 3)
Biological Sampling. Sections are further
divided into subsections, covering relevancy
of a measure, detailed steps to collect data,
lists of equipment and supplies, and
alternative ways of quantifying measures
(where applicable). References are provided
-------
at the end of each subsection to aid the reader.
We refer to example field sheets for recording
data throughout the manual. Complete copies
of these field sheets are provided at end of the
manual in Appendix 1. The procedures
described in this document are intended to
maximize the information gained for amount
of resources expended. The initial intent of
most procedures described is to collect
information that characterizes the hydrologic
permanence of stream reaches (i.e.,
indicators); however, most measures are also
commonly used in stream condition
assessments (e.g., macroinvertebrates,
substrate size).
References
Abell, D. L. 1984. Benthic invertebrates of
some California intermittent streams. Pages
46-60 in S. Jain and P. Moyle (editors).
Vernal pools and intermittent streams.
University of California Press, Davis
Institute of Ecology, Davis, California.
Angradi, T. R. (editor). 2006. Environmental
Monitoring and Assessment Program: Great
River Ecosystems field operations manual.
EPA/620/R-06/002, U.S Environmental
Protection Agency, Duluth, Minnesota.
Barbour, M. T., J. Gerritsen, B. D. Snyder,
and J. B. Stribling. 1999. Rapid
bioassessment protocols for use in streams
andwadeable rivers: periphy'ton, benthic
macroinvertebrtes and fish. 2nd edition,
EPA/841/B/98-010. Office of Water, U.S.
Environmental Protection Agency,
Washington, D.C.
Boulton, A. J., F. Sheldon, M. C. Thorns, and
E. H. Stanley. 2000. Problems and
constraints in managing rivers with variable
flow regimes. Pages 415-430 in P. J. Boon,
B. R. Davies, and G. E. Petts (editors).
Global perspectives on river conservation:
science, policy and practice. John Wiley &
Sons, Chichester, United Kingdom.
Brown, A. V., Y. Aguila, K. B. Brown, and
W. P. Fowler. 1997. Responses of benthic
macroinvertebrates in small intermittent
streams to silvicultural practices.
Hydrobiologia 347:119-125.
Clifford, H. F. 1966. The ecology of
invertebrates in an intermittent stream.
Investigations of Indiana Lakes and Streams
7(2):57-98.
Comin, F. A. and W. D. Williams. 1994.
Parched continents: our common future?
Pages 473-527 in R. Margalef (editor).
Limnology now: a paradigm of planetary
problems. Elsevier Science, Amsterdam,
Netherlands.
Daborn, G. R. and H. F. Clifford. 1974.
Physical and chemical features of an aestival
pond in western Canada. Hydrobiologia
44:43-59.
Delucchi, C. M. 1988. Comparison of
community structure among streams with
different temporal flow regimes. Canadian
Journal of Zoology 66:579-586.
Dodds, W. K. 1997. Distribution of runoff
and rivers related to vegetative
characteristics, latitude, and slope: a global
perspective. Journal of the North American
Benthological Society 16:162-168.
Drummond, R. R. 1974. When is a stream a
stream? The Professional Geographer
26:34-37.
Chapman L. J. and D. L. Kramer.
Limnological observations of an intermittent
tropical dry forest stream. Hydrobiologia
226:153-166.
-------
Feminella, J. W. 1996. Comparison of benthic
macroinvertebrate assemblages in small
streams along a gradient of flow
permanence. Journal of the North American
Benthological Society 15:651-669.
Fritz, K. M. and W. K. Dodds. 2002.
Macroinvertebrate assemblage structure
across a tallgrass prairie stream landscape.
Archivfur Hydrobiologie 154:79-102.
Gasith, A. and V. H. Resh. 1999. Streams in
Mediterranean climate regions: abiotic
influences and biotic response to predictable
seasonal events. Annual Review of Ecology
and Systematics 30:51-81.
Genereux, D. P., H. F. Hemond, and P. J.
Mulholland. 1993. Spatial and temporal
variability in streamflow generation on the
West Fork of Walker Branch Watershed.
Journal of Hydrology 142:13 7-166.
Gomi, T., R. C. Sidle, and J. S. Richardson.
2002. Understanding processes and
downstream linkages of headwater systems.
BioScience 52:905-916.
Hansen, W. F. 2001. Identifying stream types
and management implications. Forest
Ecology and Management 143:39-46.
Hewlett, J. D. and A. R. Hibbert. 1967.
Factors affecting the response of small
watersheds to precipitation in humid areas.
Pages 275-290 in W. E. Sopper and H. W.
Lull (editors). Forest Hydrology. Pergamon
Press, New York.
Hughes, R. M. and J. M. Omernik. 1983. An
alternative for characterizing stream size.
Pages 87-102 in T. D. Fontaine and S. M.
Bartell (editors). Dynamics oflotic
ecosystems. Ann Arbor Science, Ann Arbor,
Michigan.
Jarnagin, S. T. and D. B. Jennings. 2005.
Collaborative science for environmental
solutions: Collaborative hydrologic research
in the Clarksburg special protection area.
(Ab stract) US Environmental Protection
Agency Science Forum., Washington, DC
May 16-18,2005.
Johansson, A. and A. N. Nilsson. 1994.
Insects of a small aestival stream in northern
Sweden. Hydrobiologia 294:17-22.
Lazorchak, J. M., D. J. Klemm, and D. V.
Peck (editors). 1998. Environmental
Monitoring and Assessment Program -
Surface Waters: field operations and
methods for measuring the ecological
condition ofwadeable streams. EPA/620/R-
94/004F. U.S. Environmental Protection
Agency, Washington, D.C.
Lazorchak, J. M., B. H. Hill, D. K. Averill, D.
V. Peck, and D. J. Klemm (editors). 2000.
Environmental Monitoring and Assessment
Program - Surface Waters: field operations
and methods for measuring the ecological
condition ofnon-wadeable rivers and
streams. EPA/620/R-00/007. U.S.
Environmental Protection Agency,
Washington, D.C.
Leopold, L. B. 1994. A View of the River.
Harvard University Press, Cambridge,
Massachusetts.
Likens, G. E., F. H. Bormann, N. M. Johnson,
D. W. Fisher, and R. S. Pierce. 1970.
Effects of forest cutting and herbicide
treatment on nutrient budgets in the Hubbard
Brook watershed-ecosystem. Ecological
Monographs 40:23-47.
-------
Mark, D. M. 1983. Relations between field-
surveyed channel networks and map-based
geomorphometric measures, Inez, Kentucky.
Annals of the Association of American
Geographers 73:358-372.
McCleary, F. E. and S. J. Hamilton. 1998.
Soil Survey of Lawrence County, Ohio. U.S.
Department of Agriculture, Natural
Resources Conservation Service, 231 p., 70
folded plates.
McMahon, T. A. and B. L. Finlayson. 2003.
Droughts and anti-droughts: the low flow
hydrology of Australian rivers. Freshwater
Biology 48:1147-1160.
Meyer, J. L. and J. B. Wallace. 2001. Lost
linkages and lotic ecology: rediscovering
small streams. Pages 295-317 in M. C.
Press, N. J. Huntly, and S. Levin (editors).
Ecology: achievement and challenge.
Blackwell Science, Oxford, United
Kingdom.
Montgomery, D. R. and W. E. Dietrich. 1988.
Where do channels begin? Nature 336:232-
234.
Montgomery, D. R. and W. E. Dietrich. 1992.
Channel initiation and the problem of
landscape scale. Science 255:826-830.
Nadeau, T.-L. and M. R. Rains. In press.
Hydrological Connectivity Between
Headwater Streams and Downstream
Waters: How Science Can Inform Policy.
Journal of the American Water Resources
Association.
Nanson, G. C., S. Tooth, and A. D. Knighton.
2002. A global perspective on dryland
rivers: perceptions, misconceptions and
distinctions. Pages 17-54 in L. J. Bull and
M. J. Kirkby (editors) Dryland rivers:
hydrology and geomorphology of semi-arid
channels. John Wiley & Sons, Chichester,
United Kingdom.
NC Division of Water Quality. 2005.
Identification Methods for the Origins of
Intermittent and Perennial streams, Version
3.1. North Carolina Department of
Environment and Natural Resources,
Division of Water Quality. Raleigh, NC.
http://h2o.enr.state.nc.us/ncwetlands/docum
ents/NC_Stream_ID_Manual. pdf
Ohio Environmental Protection Agency.
2002. Field Evaluation Manual for Ohio's
Primary Headwater Habitat Streams, Final
Version 1.0. Ohio Environmental Protection
Agency, Division of Surface Water,
Columbus, OH.
http://www.epa.state.oh.us/dsw/wqs/headwa
ters/PHWHManual 2002 102402.pdf
Poff, N. L. and J. V. Ward. 1989.
Implications of streamflow variability and
predictability for lotic community structure:
a regional analysis of streamflow patterns.
Canadian Journal of Fisheries and Aquatic
Sciences 46:1805-1818.
Rivenbark, B. L. and C. R. Jackson. 2004.
Average discharge, perennial flow initiation,
and channel initiation - small southern
Appalachian basins. Journal of the
American Water Resources Association
40:639-646.
Sprules, W. M. 1947. An ecological
investigation of stream insects in Algonquin
Park, Ontario. University of Toronto Studies
Biological Series No. 56. Publication of the
Ontario Fisheries Research Laboratory 69:1-
81.
Svec, J. R., R. K. Kolka, and J. W. Stringer.
2005. Defining perennial, intermittent, and
ephemeral channels in eastern Kentucky:
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application to forestry best management
practices. Forest Ecology and Management
214:170-182.
Uys, M. C. and J. H. O'Keefe. 1997. Simple
words and fuzzy zones: early directions for
temporary river research in South Africa.
Environmental Management 21:517-531.
Wallace, J. B., S. L. Eggert, J. L. Meyer, and
J.R.Webster. 1999. Effects of resource
limitation on a detrital-based ecosystem.
Ecological Monographs 69:409-442.
Williams, D. D. 2006. The Biology of
Temporary Waters. Oxford University Press,
Oxford, United Kingdom.
Zale, A.V., D. M. Leslie, W.L. Fisher, and
S.G. Merrifield. 1989. The
physicochemistry, flora, and fauna of
intermittent prairie streams: a review of the
literature. Biological Report 89(5), US
Department of Interior, Fish and Wildlife
Service. Washington, DC.
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2 FACTORS INFLUENCING STUDY DESIGN
This section discusses some initial
considerations for planning a study of
headwater streams. This section is not
intended to cover all possible issues when
preparing a study or assessment. Rather,
general options and some unique
considerations for headwater streams are
discussed.
Clearly stated objectives (and associated
hypotheses) are important to any scientific
study and should be decided before moving
forward. The objectives should set the initial
stage for what and how much will be
measured. Therefore, the spatial and temporal
scales (sampling resolution) and scope (range
or extent of the study) should be determined
by the data needed to meet the objectives or
test the hypotheses. Logistical and economic
constraints also influence the scale and scope
of studies. Norris et al. (1992) point out that
the objectives of most studies fall into two
general categories: 1) determining values at a
single location and time; and 2) comparing
values from multiple locations or time periods.
In the first case the goal is to provide an
accurate estimate (e.g., total density), whereas
the second focuses on comparing the
difference of values between locations or time
periods. Downes et al. (2002) identified four
general objectives for assessment studies: 1)
assess the ecological state of ecosystems; 2)
determine if regulated criteria have been
exceeded; 3) detect and quantify impacts
generated by anthropogenic disturbance(s);
and 4) assess the effectiveness of restoration
projects. In any case, the objectives should
guide the design, implementation, and analysis
of the study.
Field sampling designs
After identifying the specific objectives,
decisions are made regarding the study design
(i.e., how, what, when and where to sample).
There are two major categories for study
designs: comparative and manipulative.
Comparative (also called measurative) studies
have location or time period as the primary
treatment(s) being investigated, where the
treatment exists without the intervention of the
scientists. An example of a comparative study
is comparing biological and physiochemical
measures among streams with different land
uses or an intensity gradient of a land practice.
The primary treatment of manipulative (or
experimental) studies is an intervention or
perturbation by the investigators. An example
of a manipulative study is measuring the
biological characteristics in one set of streams
where large woody debris has been removed
by the investigators and in another where large
woody debris is left intact. Manipulative
studies generally offer more control over the
independent variables (and therefore greater
potential to identify cause-effect relationships)
than comparative studies. On the other hand,
comparative studies typically offer greater
realism and generality than manipulative
studies. The main effects (or treatment
differences) and associated variation of effects
over the study duration of comparative studies
are directly relevant to the systems studied.
Investigators designing experimental studies
should strive to apply realistic manipulations
(i.e., relevant to real world situations) to
experimental units. Both categories have
merits and limitations that should be
considered when planning a study (see
Diamond 1986 for a detailed discussion).
Spatial and temporal scales of a study should
match the objectives and be relevant to the
10
-------
organisms and environments studied. Gotelli
and Ellison (2004) identified two aspects of
spatial scale that should be addressed when
designing a study: the grain (size of the
smallest unit of study) and the extent (total
area encompassed by all units in the study).
Investigators need to efficiently balance the
size of the grain and extent of study with
logistics and cost to effectively achieve the
scope of the objectives. Temporal scale
includes the time needed to collect a sample,
the frequency of sampling, and the duration of
the study. Hierarchical or nested designs can
be used to identify variation associated with
different spatial scales, and repeated measures
designs assess interaction among sampling
periods and treatments. Stratification of
sampling by habitat type can account for
variation that would otherwise be considered
in the error.
A critical aspect of a field study is the sample
size needed to effectively test a hypothesis or
to provide an acceptable level of confidence
around estimates of resource condition. Often
the emphasis for condition surveys is to
estimate the proportion of a resource among
classes of condition (e.g., Diaz-Ramos et al.
1996). Condition classes reflect categories of
ecological integrity and are measured with
indicators representing various physical and
biological parameters. Thresholds separating
condition classes are typically set by
regulatory standards. The formula for
estimating the standard error for a proportion
is:
P(1-P)
n
where/? is the proportion of a population
representative of a class and n is the total
population size (i.e., sample size). By
assuming a proportion that results in the
largest estimate of the standard error of the
proportion (p =0.5), one can visualize that
standard error decreases asymptotically with
increasing sample size (Figure 2-1).
0.6
0.5
c
o
IS
E
to
UJ
I
UJ
1 0.2
•o
c
to
« 0.1
0.4
0.3
20 40 60 80
Number of Samples
100
Figure 2-1 Relationship between sample
size and standard error estimations
assuming proportions are equal among
populations.
Therefore, confidence around estimates also
increases with higher sample size, but
investigators need to balance sampling cost
and acceptable level of confidence when
designing surveys.
In hypothesis testing, power analysis can be
useful for determining the appropriate number
of replicates to provide sufficient statistical
power for an expected effect size (the
detectable difference between treatments) and
natural variation (Peterman 1990, Fairweather
1991, Foster 2001). Statistical power
measures the probability of correctly rejecting
the null hypothesis when in fact it is false
(converse of the probability of Type II error).
Power is generally described as:
ES • a • 4n
rower cc
s
where ES is effect size, a is the a priori
significance level (Type I error probability), n
is the sample size, and s is the standard
deviation among replicate units. This
relationship indicates that for a given effect
11
-------
size and level of variability, power increases
with higher study unit replication; however,
with that in mind, increasing sample size can
enable detection of very small effects that may
not be ecologically significant. Larger effect
sizes are more likely to be detected than
smaller ones with the same sample size and
level of variability. The actual formulae for
calculating power or deriving appropriate
sample size or minimum detectable effect size
will vary with statistical test and test statistic
(see Cohen 1988, Zar 1998). Effect size may
be derived from previous studies, regulatory
thresholds, or convention (e.g., order of
magnitude). Expected variation can be taken
from the literature or pilot studies. An
appropriate a priori level of statistical power
will vary depending upon the objectives of the
study. For example, failing to detect an
environmental impact where one exists (i.e.,
Type II error) may have greater consequences
than detecting an impact that does not exist
(Type I error), therefore greater power may be
desired to protect against a Type II error (see
Peterman 1990, Di Stefano 2003). Frequently,
cost (time and money) is a critical factor
governing sample size. Mapping the study
beforehand (estimating time and costs) will
help determine the feasibility of the study
design. Designing an effective study is
balancing effect size, sample size, and cost to
meet the study objective.
Randomization should be used whenever
feasible to ensure unbiased data collection.
Random or probabilistic site selection
produces a representative sample of the
population(s) targeted under the study
objectives, so that results can be more
confidently extrapolated to the overall
population from which the selected sites were
randomly chosen. In contrast, targeted
sampling focuses the effort toward a specific
problem. The difficulty with randomized site
selection is the a priori knowledge of the
entire population of possible sites or sampling
points within the bounds of the study
objective. If the scope of the objectives is
narrow and the population is known (e.g.,
water bodies within Central Park),
probabilistic sampling is more feasible
compared to broader scales where the
population is uncertain (e.g., spring seeps of
Kentucky). The scope of a study will be
narrowed under most circumstances because
of the inability to account for the entire
population of potential sites. Time of data
collection is rarely randomly selected because
of the stochastic nature of streams; however,
seasonal sampling is usually desired. Index
periods are typically determined by the
logistics of sampling and the life history of
targeted biota.
There are practical difficulties associated with
large scale experiments, including the need for
a large number of independent replicates to
overcome natural variability among replicate
study units. A study design that is
increasingly used in stream research is the
Before/After and Control/Impact (BACI;
Stewart-Oaten et al. 1986, Carpenter et al.
1989, Downes et al. 2002). In this design, one
or more control sites and one or more impact
sites are simultaneously sampled multiple
times, both before and after the manipulation
to the impact site(s). The difference in
parameters measured between the control and
impact at each time period represents a
replicate unit for the Before and After
treatments. Underwood (1991, 1992) strongly
advocated the incorporation multiple
randomly selected control sites in the design
to overcome the possibility that the control
and impact sites may have naturally different
trends in the measured parameters. Further
issues and concerns about BACI designs are
reviewed by Smith et al. 1993, Osenberg et al.
1994, and Downes et al. 2002. An in-depth
discussion of specific statistical designs is
12
-------
outside the scope of this manual. A few
relevant texts for ecological studies include:
Clarke and Warwick (2001), Gotelli and
Ellison (2004), Scheiner and Gurevitch
(1993), Quinn and Keough (2002), and
Underwood (1997).
Special considerations for headwater streams
Headwater streams are narrower, shallower,
have higher drainage density, and are more
likely to dry than larger streams and rivers.
Their position in the network also makes
many headwater streams more responsive to
precipitation, so lag time is shorter between
precipitation and peak discharge. Notable
exceptions to this are spring-fed streams,
where deep and more stable groundwater
discharge can dominate the hydrologic
regime. Depending upon the geographic
location, headwater streams may have higher
gradients and therefore the repeating habitat
units are typically more closely spaced than
wadable streams. Reach lengths for
ecological assessment are typically scaled to
the channel width (e.g., Barbour et al. 1999,
Lazorchak et al. 1998, Moulton et al. 2002).
Following this convention, reach lengths of
headwaters are shorter than those needed for
larger perennial streams and rivers. Multiple
reaches or longer reaches may be required for
studies using multiple indicators or assessment
approaches (i.e. amphibian surveys, tracer
additions, etc.). If multiple reaches are used,
they should be as close as possible given the
sampling or logistical limitations. They
should have similar channel dimensions and
levels of permanence, avoiding influences by
intervening tributary confluences. Higher
drainage density affords the opportunity to
have nearby replicate streams for studies, but
also may result in frequent discontinuities
(e.g., abrupt changes in substrate size) at
tributary confluences (Rice et al. 2001, Brenda
et al. 2004). Unique sampling methods are
often required for headwaters because the low
flows prevent use of many conventional
sampling devices. For example, core samples
are preferred for headwater invertebrate
sampling rather than Surber or other net
samplers that require sufficient flow to carry
dislodged debris into the net. Estimates of
flow permanence are critical and may be the
master variable influencing headwater
communities. Measures of channel dimension
and substrate size may provide critical insight
into the typical flow regime or degree of
permanence at a site and should be included in
any headwater assessment.
Time of year for sampling is critical in
temporary headwaters because precipitation
and evapotranspiration has a relatively strong
influence on stream discharge. Historic
hydrological data are rare for headwater
streams because most gauges are positioned
on wadeable streams and large rivers.
Discharge data from downstream gauges can
provide an integrated measure of precipitation
and evapotranspiration for a basin. The utility
of gauging data from downstream locations
will depend upon their distance from
headwater sites, their position relative to
reservoirs (where levels may reflect not solely
precipitation, but recreational and
socioeconomic use), and changes in watershed
land cover. In addition, many gauges on
intermediate size streams and rivers have been
retired, and therefore problematic for
developing stage relationships with headwater
sites. However, long-term precipitation
records may serve as surrogate for flow. The
seasonal and interannual variation in
precipitation and hydrologic observations
provide the likelihood of flowing conditions.
While year-round sampling (both dry and wet
seasons) over several years may be optimal for
categorizing or assessing a headwater site,
researchers are rarely afforded such
opportunities. For shorter-term studies,
sampling should take place during the driest
13
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and wettest periods of the year to assess
extreme conditions. If sampling is restricted
to one visit, headwater index periods will
typically be during the spring when flow is
higher, and most aquatic organisms can be
collected.
The gradual change in environmental
conditions (e.g., lower dissolved oxygen,
higher temperatures) as temporary habitats dry
can be as critical to understanding
mechanisms influencing biotic response as the
duration and frequency of drying.
Disturbances (disrupting force) or
perturbations (sequence of disrupting force
and system response) have been classified as
either pulse or press events (Bender et al.
1984, Glasby and Underwood 1996). A pulse
disturbance is characterized by a short and
sharply delineated event (relative to the time
scale of the response measure, Figure 2-la),
whereas a press disturbance has a continuous
and constant level that is relative long-lasting
(Figure 2-lb). In contrast to pulse and press
disturbances, environmental conditions for
many organisms worsen over time as streams
dry (Slack and Feltz 1968, Towns 1985,
Ostrand and Wilde 2004). Lake (2000, 2003)
characterized this difference by
conceptualizing that drying or drought was a
"ramp" disturbance (Figure 2-lc). As the
sequence of physicochemical changes
progresses, greater stress is placed upon
inhabitants, causing more taxa to succumb or
emigrate over time. Rather than a steady
sequence of physicochemical changes of a
"ramp", Boulton (2003) argues that the
sequence of changes may be better
characterized as a series of "steps" (Figure 2-
Id), wherein critical thresholds cause
substantial shifts in wetted habitat (e.g., drying
of riffles, subsurface habitat). Differences
between the ramp and stepped models may be
to some extent dependent upon the
hierarchical scale through which the drying
process is approached (Stanley et al. 1997).
Some changes may be more apparent at small
spatial or temporal scales, but undetectable at
larger scales.
O)
c
(U
CO
Time
Figure 2-2 Types of disturbance (solid) and
responses (dashed) in streams: pulse (a),
press (b), ramp (c) and stepped (d). Based
on figures from Lake (2003) and Boulton
(2003).
Wetted area and volume are reduced initially
in the drying sequence that leads to increased
isolation of the wetted area from stream banks
(contraction toward the deeper flowpaths in
the channel) and between habitat units
(contraction to pools). As discharge declines,
flow may at first become braided between
larger emergent substrates, then become
limited to strong upwelling zones along the
channel, and then finally cease altogether,
leaving surface water to remain only in deep
pools. These remaining pools shrink by
evaporation and the hyporheic habitat
(subsurface zone between the surface water
and groundwater) dries if water deficit
continues. The rate of channel drying varies
with channel gradient, degree of exposure (to
wind and sun), evapotranspiration by
14
-------
watershed vegetation, soil moisture status, and
permeability or infiltration capacity from the
surrounding watershed. Vegetation cover,
type, and succession stage can also influence
headwater stream hydrology (Bosch and
Hewlett 1982). For example, annual stream
flow is typically lower in streams draining
conifers because of higher annual interception
(and subsequently evaporation) of
precipitation and higher transpiration loss at
the beginning and end of the growing season
than hardwoods (e.g., Swank et al. 1988).
Streams draining limestone or "karst" geology
retain surface water for shorter periods than
streams draining geologic materials with
lower hydraulic conductivity and effective
porosity (e.g., sandstone and clay). Many of
these factors will also influence the timing of
flow commencement following precipitation
(Blyth and Rodda 1973, Day 1978, de Vries
1995) or leaf abscission (Doyle 1991).
As previously mentioned, headwaters have a
distinct bioassessment advantage because the
small watershed areas make stressor
identification more straight-forward.
However, the timing of sample collection
relative to the resumption of flow or start of
drying is critical. The diversity, abundance,
and biomass of benthic organisms increase
and community composition shifts with time
following the resumption of flow (Peterson
1987, Boulton and Lake 1992, Fritz and
Dodds 2002, 2004). The rate of assemblage
recovery varies with magnitude, duration, and
extent of drying, particularly in relation to the
permanence history (i.e., flow predictability)
of streams. Resilience will likely vary among
assemblage types and biological parameters
(e.g., abundance, biomass) because of
differences in the recovery mechanism (i.e.,
resistance vs. colonization), vagility, and
growth rates. For purposes of bioassessments,
samples should be collected near the peak of
recovery from drying to maximize the number
of indicator taxa present and biotic index or
metric discrimination among condition
categories.
Minimizing impacts associated with sampling
The potential for impacting streams during
sampling is higher for headwater streams
compared to larger streams and rivers, and
therefore requires special consideration.
Small wetted areas mean that sample
collection and geomorphic measurements can
potentially disturb a large portion of the local
channel with potential adverse effects
downstream. Individual substrates (e.g.,
cobble, small woody debris) that are
inconsequential in larger streams and rivers
may provide important geomorphic functions
in headwater streams. Channel alteration
caused by sampling may be more persistent in
small streams than in larger channels because
the power associated with flood events that
resets channels is typically lower. Sampling
in an upstream direction is typical in larger
streams, and it is especially important when
working in headwaters to minimize trampling
the stream reach during assessments. Because
headwater streams are small and positioned at
the tips of stream networks, oversampling of
unique populations and species is a concern.
Headwater streams, particularly those that are
spring-fed, often contain endemic taxa (Hubbs
1995, Ferrington 1995, Myers et al. 2001).
Rather than further the endangerment of these
unique communities, sampling protocols
should provide information for their
conservation.
References
Barbour, M.T., J. Gerritsen, B.D. Snyder, and
J.B. Stribling. 1999. Rapid Bioassessment
Protocols for Use in Streams and Wadeable
Rivers: Periphyton, Benthic
Macroinvertebrates and Fish. Second
edition, EPA/841/B/98-010. Office of
15
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Water, U.S. Environmental Protection
Agency, Washington, D.C.
Benda, L., N. L. Poff, D. Miller, T. Dunne, G.
Reeves, G. Pess, and M. Pollock. 2004.
The network dynamics hypothesis: how
channel networks structure riverine habitats.
BioScience 54:413-427.
Bender, A., T. J. Case, M. E. Gilpin. 1984.
Perturbation experiments in community
ecology: theory and practice. Ecology 65:1-
13.
Blyth, K. and J. C. Rodda. 1973. A stream
length study. Water Resources Research
9:1454-1461.
Bosch, J. M. and J. D. Hewlett. 1982. A
review of catchment experiments to
determine the effect of vegetation changes
on water yield and evapotranspiration.
Journal of Hydrology 55:3-23.
Boulton, A. J. 2003. Parallels and contrasts
in the effects of drought on stream
macroinvertebrate assemblages. Freshwater
Biology 48:1173-1185.
Boulton, A. J. and P. S. Lake. 1992. The
ecology of two intermittent streams in
Victoria, Australia. III. Temporal changes
in faunal composition. Freshwater Biology
27:123-138.
Clarke, K. R. and R. W. Warwick. 2001.
Change in Marine Communities: An
Approach to Statistical Analysis and
Interpretation, 2nd edition. PRIMER-E Ltd.,
Plymouth, United Kingdom.
Cohen, J. 1988. Statistical Power Analysis
for the Behavioral Sciences, 2nd edition.
Lawrence Erlbaum, New Jersey.
Day, D. G. 1978. Drainage density changes
during rainfall. Earth Surface Processes
3:319-326.
de Vries, J. J. 1995. Seasonal expansion and
contraction of stream networks in shallow
groundwater systems. Journal of Hydrology
170:15-26.
Diamond, J. 1986. Overview: laboratory
experiments, field experiments, and natural
experiments. Pages 3-22 in J. Diamond and
T. J. Case (editors). Community ecology.
Harper & Row, New York.
Di Steffano, J. 2003. How much power is
enough? Against the development of an
arbitrary convention for statistical power
calculations. Functional Ecology 17:707-
709.
Diaz-Ramos, S., D. L. Stevens, Jr and A. R.
Olsen, 1996. EMAP Statistical Methods
Manual. EPA/620/R-96/002, U.S.
Environmental Protection Agency, Office of
Research and Development, NHEERL-
WED, Corvallis, Oregon.
Downes, B. J., L. A. Barmuta, P. G.
Fairweather, D. P. Faith, M. J. Keough, P. S.
Lake, B. D. Mapstone, and G. P. Quinn.
2002. Monitoring ecological impacts -
concepts and practice in flowing waters.
Cambridge University Press, Cambridge,
UK.
Doyle, P. F. 1991. Documented autumnal
streamflow increase without measurable
precipitation. Water Resources Bulletin
27:915-923.
Fairweather, P. G. 1991. Statistical power
and design requirements for environmental
monitoring. Australian Journal of Marine
and Freshwater Research 42:555-567.
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Ferrington, L. C. (editor) 1995. Biodiversity
of aquatic insects and other invertebrates in
springs. Journal of the Kansas
Entomological Society 68:1-165.
Foster, J. R. 2001. Statistical power in forest
monitoring. Forest Ecology and
Management 151:211 -222.
Fritz, K. M. and W. K. Dodds. 2002.
Macroinvertebrate assemblage structure
across a tallgrass prairie stream landscape.
Archivfur Hydrobiologie 154:79-102.
Fritz, K. M. and W. K. Dodds. 2004.
Resistance and resilience of
macroinvertebrate assemblages to drying
and flood in a tallgrass prairie stream
system. Hydrobiologia 527:99-112.
Glasby, T. M. and A. J. Underwood. 1996.
Sampling to differentiate between pulse and
press perturbations. Environmental
Monitoring and Assessment 42:241-252.
Gotelli, N. J. and A. M. Ellison. 2004. A
Primer of Ecological Statistics. Sinauer,
Sunderland, Massachusetts.
Hubbs, C. 1995. Springs and spring runs as
unique aquatic systems. Copeia 1995:989-
991.
Lake, P. S. 2000. Disturbance, patchiness,
and diversity in streams. Journal of the
North American Benthological Society
19:573-592.
Lake, P. S. 2003. Ecological effects of
perturbation by drought in flowing waters.
Freshwater Biology 48:1161-1172.
Lazorchak, J. M., D. J. Klemm, and D. V.
Peck (editors). 1998. Environmental
Monitoring and Assessment Program -
Surface Waters: Field Operations and
Methods for Measuring the Ecological
Condition ofWadeable Streams.
EPA/620/R-94/004F. U.S. Environmental
Protection Agency, Washington, D.C.
Moulton, S. R. II, J. G. Kennen, R. M.
Goldstein, and J. A. Hambrook. 2002.
Revised Protocols for Sampling Algal,
Invertebrate, and Fish Communities As Part
of the National Water-Quality Assessment
Program. Department of Interior, U.S.
Geological Survey Open-File Report 02-
150, 75 p.
Myers, M. J., F. A. H. Sperling, and V. H.
Resh. 2001. Dispersal of two species of
Trichoptera from desert springs:
conservation implications for isolated vs
connected populations. Journal of Insect
Conservation 5:207-215.
Norris, R. H., E. P. McElravy, and V. H. Resh.
1992. The sampling problem. Pages 282-
306 in P. Calow and G. E. Petts (editors)
The rivers handbook: hydrological and
ecological principles Volume 1. Blackwell
Scientific, Oxford, United Kingdom.
Osenberg, C. W., R. J. Schmitt, S. J.
Holbrook, K. E. Abu-Saba, and A. R. Flegal.
1994. Detection of environmental impacts:
natural variability, effect size, and power
analysis. Ecological Applications 4:16-30.
Ostrand, K. G. and G. R. Wilde 2004.
Changes in prairie stream fish assemblages
restricted to isolated streambed pools.
Transactions of the American Fisheries
Society 133:1329-1338.
Peterman, R. M. 1990. Statistical power
analysis can improve fisheries research and
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management. Canadian Journal of Fisheries
and Aquatic Sciences 47:2-15.
Peterson, C. G. 1987. Influence of flow
regime on development and desiccation
response of lotic diatom communities.
Eco/ogy 68:946-954.
Quinn, G. P. and M. J. Keough. 2002.
Experimental Design and Data Analysis for
Biologists. Cambridge University Press,
Cambridge, United Kingdom.
Rice, S. P., M. T. Greenwood, and C. B.
Joyce. 2001. Tributaries, sediment sources,
and the longitudinal organisation of
macroinvertebrate fauna along river
systems. Canadian Journal of Fisheries and
Aquatic Sciences 58:824-840.
Scheiner, S. M. and J. Gurevitch (editors).
1993. Design and Analysis of Ecological
Experiments. Chapman & Hall, New York.
Slack, K. V. and H. R. Feltz. 1968. Tree leaf
control on low flow water quality in a small
Virginia stream. Environmental Science and
Technology 2:126-131.
Smith, E. P., D. R. Orvos, J. Cairns. 1993.
Impact assessment using the before-after-
control-impact (BACI) model: concerns and
comments. Canadian Journal of Fisheries
and Aquatic Sciences 50:627-637.
Stanley, E. H., S. G. Fisher, and N. B. Grimm.
1997. Ecosystem expansion and contraction
in streams BioScience 47:427-435.
Stewart-Oaten, A., W. W. Murdoch, and K. R.
Parker. 1986. Environmental impact
assessment: "pseudoreplication" in time?
Ecology 67:929-940.
Swank, W. T., L. W. Swift, and J. E. Douglas.
1988. Streamflow changes associated with
forest cutting, species conversions and
natural disturbance. Pages 297-312 in W. T.
Swank and D. A. Crossley (editors) Forest
hydrology and ecology at Coweeta.
Ecological Studies Volume 66, Springer-
Verlag, New York.
Towns, D. R. 1985. Limnological
characteristics of a South Australian
intermittent stream, Brown Hill Creek.
Australian Journal of Marine and
Freshwater Research 36:821-837.
Underwood, A. J. 1997. Experiments in
Ecology. Cambridge University Press,
Cambridge, United Kingdom.
Zar, J. H. 1998. Biostatistical Analysis, 4th
edition. Prentice Hall, Englewood Cliffs,
New Jersey.
2.1 Study design for comparing across
stream reaches with varying hydrologic
permanence.
This section describes a specific study design
used for comparing among headwater stream
reaches varying in hydrologic permanence.
The objectives were: 1) to characterize
biological and physical features of reference
headwater streams across a gradient of
hydrologic permanence (frequency and
duration of drying) and 2) to identify
indicators of hydrologic permanence. The
study focused on headwater streams in intact
forests to limit potentially confounding effects
of land use on hydrology. Streams were
sampled in Indiana, Illinois, Kentucky, Ohio,
New Hampshire, New York, Vermont,
Washington, and West Virginia. The drainage
area of study sites was restricted to basins
< 2.92 km2 (1 mi2) that corresponded to the
upper boundary of streams measured. For
assessment purposes, Ohio EPA is using this
18
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drainage area size to distinguish "Primary
Headwater Habitat Streams" from the rest of
the stream network (Ohio Environmental
Protection Agency 2002).
Selecting study units that incorporate the
range of hydrologic permanence (i.e., from
ephemeral to perennial) was critical to meet
the goals of the study. No data for annual
hydrologic patterns were available prior to
sampling, so the general positive relationship
between drainage area and flow permanence
was used to select sites (i.e., drainage area was
used as a surrogate for flow permanence). As
drainage area increases, groundwater storage
increases and approaches the level of the
streambed. Exceptions to this general pattern
include perched aquifers and artesian springs
in upper reaches that sustain year-round
surface flow (Dunne and Leopold 1978) or
substantial storage in swale soils above the
channel head that sustains patches with
perennial surface water (Hunter et al. 2005).
These characteristics can result in fragmented
longitudinal patterns of flow permanence
along headwater streams (Lake 2003).
Likewise, local changes in streambed
topography along a stream influence the
spatial pattern of hydrologic permanence.
Sediments and woody debris originating from
landslides, debris flows, and windthrow are
transported downstream and deposited in
reaches with lower gradient (Benda and
Dunne 1987, Grizzel and Wolff 1998,
Montgomery 1999). These deposits (i.e.,
sediment wedges) locally elevate the
streambed above the dry season water table,
causing reaches with such deposits to
seasonally dry (May and Lee 2004, Harvey et
al. 2005). Recognizing this, the study design
incorporated multiple study units along
multiple headwater streams. This design
included a broad range of hydrologic regimes
and capable of detecting repeating
associations between stream features
(biological and physical) and hydrology.
The study units were 30-m long reaches of
stream channel (land form with bed and bank
features). This length is on average 40X the
headwater channel width and is consistent
with study units used by USEPA in the
Environmental Monitoring and Assessment
Program (EMAP). Adjustment of the reach
length may be needed to incorporate repeating
geomorphic channel units. Three or four 30-
m study reaches were selected within each
stream. The aim was to include 1 reach with
perennial flow, 2 reaches with varying degrees
of intermittent flow and 1 reach with
ephemeral flow. This design ensures
sampling across a sufficient range of
hydrologic conditions within a stream, while
also allowing for multiple streams to be
assessed. This sampling regime of the study
required at least 2 sampling periods for each
site within a year. These periods included
visits in spring (wet season) and late summer
(dry season), but not necessarily in that order.
An initial visit to the streams during the dry
season helps ensure that a perennial site is
sampled, but it may be difficult to determine if
a dry reach is intermittent or ephemeral at that
time. Field visits during wet and dry seasons
prior to selecting sites, where possible, may
provide greater confidence in the distribution
of sites across the flow permanence gradient.
Desktop selection procedure
In most cases the upstream study reaches
along the streams were not marked with "blue
lines", but appeared as "depressions" on 1:24
000 scale topographic maps (Figure 2-2). Red
lines have been added to Figure 2-2 to show a
more realistic and complete network of stream
channels within the Falling Rock Branch
watershed. The yellow line represents the
approximate watershed boundary. Maps
(typically 1:15 840 scale) published by USDA
19
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NRCS (formerly Soil Conservation Service)
provided better resolution of the headwater
channel network, but still underestimated the
extent of channels. Likewise, orthophotos
(e.g., 1:12 000 scale) aided planning, but the
ability to discern headwater channels varied
with photo resolution and vegetation cover.
Both types of maps and photos were used in
the planning stage, but the topographic maps
were more useful while in the field. The
definition of the upstream extent of headwater
channels is discussed in detail in Section 3.3.
Figure 2-3 Map highlighting position of headwater channels within the watershed of
Falling Rock Branch, KY. Yellow represents boundary of watershed, blue represents
"blue line" designation on the 1:24 000 USGS topographic map (Noble 7.5 minute
quadrangle, Breathitt County, KY), and red represent headwater channels not shown on
the topographic map.
The channel network configuration,
particularly dendritic or reticulate networks,
creates a nonlinear relationship between
distance downstream and drainage area.
Drainage area does not increase gradually
downstream, but rather increases in steps with
each tributary confluence. This is an
important consideration when selecting study
reaches to maximize the range of hydrologic
permanence. Tributary confluences are also
useful landmarks for returning to study
reaches for subsequent visits. Where possible,
a more gradual drainage area transition
between study reaches is preferred (Figure 2-
20
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3). In some cases the entire drainage area of
headwater stream may not be sufficient to
supply perennial flow. In this situation a
stream may need to be paired with an
adjacent, larger tributary so there is a shared
perennial site (Figure 2-4).
2 ha
SUBOPTIMAL
90 ha
PREFERRED
90 ha
Figure 2-4 Schematic showing suboptimal and preferred longitudinal positioning of sites
along headwater channels to maximize the range of hydrologic permanence across study
sites. Hypothetical drainage areas are shown to further illustrate spatial hierarchy.
Properties that contained intact forest were
identified and we obtained USGS 7.5 minute
quadrangle maps (1:24 000 scale) for the
selected areas. Land owners or managers of
the properties were contacted. We described
to them the objectives and design of the study
and provided background material that they
may need (e.g. Quality Assurance Project
Plan, research proposal). We also inquired
about headwater streams draining the
property, especially pertaining to their flow
permanence, current land use, ongoing or
previous research downstream, and
accessibility by roads and hiking trails. Being
able to quickly travel between study reaches
helped ensure that a sufficient number of
study units were assessed and that sampling
was done over a reasonable timeframe (i.e.,
within the same index period).
Field study reach - general selection
guidelines
In the field we located the stream reaches
preliminarily selected from the map. Final
selection was adjusted to ensure that the study
reaches were entirely upstream or downstream
of tributary confluences. We also selected
reaches having multiple habitat units
(erosional and depositional habitats).
Although large woody debris dams are
characteristic features of intact forested
streams, reaches with excessively large woody
debris dams (prevented access to >50% of the
wetted channel in the study reach) were
avoided when possible. These structures are
likely to 1) complicate the association
between reach properties (physical and
biological) and hydrology and 2) impede data
collection. With this in mind, debris dams are
common in some regions and will be
unavoidable when designating study reaches.
21
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Figure 2-5 Map showing positioning of sites along two Indiana headwater streams where
the downstream perennial site (P) is "shared" between two tributaries. DI = downstream
intermittent; UI = upstream intermittent; and E = ephemeral. Shading shows cumulative
drainage area in downstream direction.
Using a measuring tape, we marked the 30-m
study reach from the downstream boundary
(located at 0 m) to the upstream boundary
(located at 30 m). The tape was positioned to
follow the thalweg. The thalweg is the
deepest flow path in a channel. The study
reaches were designated using flagging tape or
other clearly visible markers attached to trees
near each boundary. The location of the study
reach was identified on the topographic map
or a PDA with electronic topographic maps,
and a written description of the study reach
location and appropriate locality information
(e.g., topographic map, county, state) was
entered on the field forms. Photographs of the
study reach were taken and coordinates from a
GPS unit were recorded. Study reaches were
consistently identified by site numbers that
increased in an upstream direction starting
with 1 at the downstream-most reach (Figure
2-5).
Field selection - initial visit in spring (wet
season)
When the initial field visit to a study region
was in the spring (wet season), then sites were
located as follows. The ephemeral site for
each headwater stream studied was designated
22
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just upstream of the origin of intermittent flow
(Paybins 2003; upstream-most location of
spatially-continuous surface flow in the spring
or wet season; Figure 2-5). The upstream
intermittent site was positioned downstream of
the origin of intermittent flow. The drainage
area of the downstream intermittent site often
incorporated at least an additional ephemeral
drainage. Similarly, the perennial site
frequently incorporated at least twice the
drainage area of the downstream intermittent
site (Figure 2-5).
Downstream
intermittent
site
Origin of perennial flow
rigin of intermittent flow
Upstream intermittent site
Figure 2-6 Schematic of headwater channels showing numerical designation and position
of study sites relative to origins of intermittent and perennial flow.
The spatial pattern of hydrologic permanence
may not reflect a downstream progression
from ephemeral to intermittent to perennial
reaches along headwater channels for reasons
discussed earlier (e.g., perched aquifers,
artesian springs). Incorporating multiple
streams into the design may provide support
for alternative longitudinal patterns of flow
permanence within headwater drainages.
Depending upon the precipitation and
geographic setting the prevalence of some
permanence categories and therefore variation
in flow permanence among study sites is more
subtle.
Field selection - initial visit in summer (dry
season)
When the initial field visit to a study region
was in the summer (dry season), then sites
were located as follows. The origin of
perennial flow (Paybins 2003; upstream-most
location of spatially continuous surface flow
in the summer or dry season; Figure 2-5) was
located. The perennial site was positioned just
downstream of the origin of perennial flow.
The other three study reaches often did not
have continuous surface flow during the
summer. The next site upstream frequently
23
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drained approximately half the drainage area
of the perennial site. The upstream
intermittent site (Site 3) was often positioned
at least one confluence upstream of Site 2.
The ephemeral site was designated near the
top of the watershed, but where there was a
defined streambed and banks. Terrestrial
herbaceous vegetation was common within the
channel of the ephemeral study reach.
References
Benda, L. and T. Dunne. 1987. Sediment
routing by debris flow. Pages 213-223. in R.
L. Beschta, T. Blinn, G. E. Grant, F. J.
Swanson, and G. G. Ice (editors)
International symposium on erosion and
sedimentation in the Pacific rim.
International Association of Hydrological
Sciences Press, Institute of Hydrology,
Wallingford, United Kingdom.
Dunne, T. and L. B. Leopold. 1978. Water in
Environmental Planning. W. H. Freemann
and Company, New York.
Grizzel, J. D. and N. Wolff. 1998.
Occurrence of windthrow in forest buffer
strips and its effect on small streams in
northwest Washington. Northwest Science
72:214-223.
Harvey, B. C., J. L. White, and R. J.
Nakamoto. 2005. Habitat-specific biomass,
survival, and growth of rainbow trout
(Oncorhynchus mykiss) during summer in a
small coastal stream. Canadian Journal of
Fisheries and Aquatic Sciences 62:650-658.
Hunter, M. A., T. Quinn, and M. P. Hayes.
2005. Low flow spatial characteristics in
forested headwater channels of southwest
Washington. Journal of the American
Water Resources Association 41:503-516.
Lake, P. S. 2003. Ecological effects of
perturbation by drought in flowing waters.
Freshwater Biology 48:1161 -1172.
May, C. L. and D. C. Lee. 2004. The
relationships among in-channel sediment
storage, pool depth, and summer survival of
juvenile salmonids in Oregon Coast Range
streams. North American Journal of
Fisheries Management 24:761 -774.
Montgomery, D. R. 1999. Process domains
and the river continuum. Journal of the
American Water Resources Association
36:397-410.
Ohio Environmental Protection Agency.
2002. Field Evaluation Manual for Ohio's
Primary Headwater Habitat Streams, Final
Version 1.0. Ohio Environmental Protection
Agency, Division of Surface Water,
Columbus, OH.
http://www.epa.state.oh.us/dsw/wqs/headwa
ters/PHWHManual 2002 102402.pdf
Paybins, K. S. 2003. Flow origin, drainage
area, and hydrologic characteristics for
headwater streams in the mountaintop coal-
mining region of southern West Virginia,
2000-01. USGS Water-Resources
Investigations Report 02-4300.
Equipment and supplies
USGS 7.5 minute quadrangle map(s)
NRCS soil survey map(s)
Measuring tape (50 m)
Flagging or other marker
Camera
GPS unit or Handheld personal computer or
Personal Digital Assistant (PDA) with digital
maps and GPS card
24
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3 PHYSICAL HABITAT CHARACTERIZATION
Physical habitat, typically refers to the
structural attributes of the stream channel. For
convenience of organization, we also discuss
the measurement of physicochemical
attributes of the stream water in this section.
Habitat degradation from land-use change is
the greatest threat to streams and their
inhabitants (Allen and Flecker 1993, Sala et
al. 2000, USEPA 2001). Although stream
scientists generally agree that habitat
degradation is a serious threat, no universally
accepted index or procedure exists to rate
physical habitat condition for streams. The
complexity and natural variation of stream
habitat, the need for rapid field protocols, and
objectivity must be balanced before such a
measure is accepted. While the development
of such a universal tool is beyond the scope of
this document, this work has modified existing
procedures and developing new ones
specifically for headwater streams. We
believe that these procedures will contribute
toward effectively quantifying condition,
identifying causes of degradation, and
restoring stream habitat.
Hierarchical classification across spatial and
temporal scales is useful for delimiting
sources of natural variability within and
among complex systems and provides a
framework for integrating information from
different levels of resolution (O'Neill et al.
1986). Such a framework for streams ranges
spatially from whole drainage networks down
to microhabitats (Frissell et al. 1986). Implicit
in this framework is an understanding that
absolute linear dimensions for spatial scales
across all streams (Brussock et al. 1985) or
even longitudinally within a stream (Vannote
et al. 1980, Montgomery 1999) are
unattainable due to variation in geology,
climate, and topography.
The stream reach is the most commonly used
and practical spatial scale for study units. The
spacing of distinctive features (e.g., pools,
riffles) within streams is partly driven by
channel width. The length of study reaches
should be sufficient to incorporate multiple
features of the same type to prevent
evaluations based solely on potentially
anomalous features. As discussed in Section
2.1, study reaches that are 30-m long are
sufficient in most cases where streams are 1-to
2-m wide.
Transect sampling (i.e., line-intercept
technique) is a commonly used method to
quantify physical habitat at the reach scale
(e.g., Platts et al. 1983, Fitzpatrick et al. 1998,
Lazorchak et al. 1998). Transect sampling
uses a series of lines (transects) that are
positioned perpendicular to flow and cross the
channel. Measurements are taken along these
transects to characterize the stream reach, and
thus, provide the investigator with mean
estimations and a degree of variation along
stream reaches. Transects can continue
beyond the stream channel where
measurements of the adjacent riparian zone,
floodplain, and terraces are of interest. The
number and positioning of transects should be
sufficient to characterize the spatial scale of
interest. Physical parameters that vary little
along a stream reach will require fewer
measurements (e.g., discharge) to arrive at
representative values than those that can vary
substantially (e.g., water depth). The
positioning of transects can be done
systematically (e.g., every meter), randomly or
stratified random (e.g., stratified by habitat
type). Systematic selection ensures that the
measurements span the entire study reach and
may be logistically easier; however, random
25
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selection may be preferred because all cross-
sections have an equal chance of being
measured (see Section 2, Field sampling
designs).
As streams dry, surface water will gradually
become constricted to the channel thalweg.
Therefore, the thalweg will often be the last
area to dry for a given channel cross-section.
The thalweg is an important location for
measuring many physical parameters because
this can be a consistent and conservative target
when comparing across sites with varying
hydrologic permanence and ecological
condition. Where transect sampling is used,
the thalweg (rather than the banks) is the
central axis along the stream where the
transects should be perpendicularly spaced.
Many of the measures described in the
following sections are centered on the thalweg
at sampling transects. Because of the narrow
widths of headwater channels, these sampling
points represent most of the channel width and
the portion of the channel width that is
inundated longest.
Characterization of physical habitat is widely
used in stream assessments (see Somerville
and Pruitt 2004); however, assessment
protocols vary in purpose, breadth, and
targeted stream type (Montgomery and
McDonald 2002). Few protocols specifically
target headwater streams, but several region-
specific assessment protocols are potentially
available. The associated objectives of these
protocols vary somewhat. For instance, the
Ohio Environmental Protection Agency's
Primary Headwater Habitat Evaluation Form
(Ohio EPA 2002) was developed to
differentiated among 1) coldwater perennial
streams, 2) warmwater perennial and
intermittent streams, and 3) ephemeral
streams. The North Carolina Division of
Water Quality's Classification Method
(NCDWQ 2005) and Fairfax County (VA)
Stormwater Planning Division's Perennial
Streams Field Identification Protocol (FCSPD
2003), were designed to classify streams based
on hydrologic permanence (i.e., ephemeral,
intermittent and perennial flow). Some
agencies like the Louisville District of the U.
S. Army Corps of Engineers (Sparks et al.
2003a, b) have adopted protocols developed
for wadable streams (USEPA Rapid Habitat
Assessment Form (RHAF); Barbour et al.
1999). The Louisville District uses RHAF, in
conjunction with specific conductivity and
macroinvertebrate bioassessment index scores
(Pond and McMurray 2002), to assess the
ecological integrity of headwater streams in
the Eastern Coalbelt Region of Kentucky.
Ideally, all three components are then used by
district personnel when reviewing Clean
Water Act Section 404 permit applications for
dredging and filling headwater streams and
determining appropriate mitigation or in lieu
of fees for impacted streams.
Habitat assessment protocols vary in level of
subjectivity; some use visually-based
qualitative attributes across categories such as
absent, weak, and strong, whereas others rely
on quantitative measures. Qualitative
protocols are advantageous under high
workloads with limited resources and training
because they often require less expertise and
time to complete than quantitative
assessments. However, the versatility,
applicability, and rigor of qualitative
assessments are more limited. For instance,
the attribute scoring of individual measures or
questions in qualitative assessments are
weighted based on regionally derived
investigations that may not be applicable
outside the original region. The data for
qualitative assessments are often categorical
or discrete (i.e., integer values) over a limited
range, whereas quantitative data can be
distributed continuously or categorized for
analyses. Lastly, data sets from sources that
26
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use the same quantitative measures are more
feasible to combine for broader assessments
than qualitatively collected parameters. Many
habitat characteristics, however, are currently
limited to only qualitative or semi-quantitative
methods for assessments (e.g., habitat unit
designations, substrate embeddedness,
instream fish cover). Wang et al. (1996) noted
that among 27 habitat characteristics evaluated
for among-observer precision, those that were
scored quantitatively (directly measured,
rather than visually scored across categories)
were more precise than qualitatively scored
characteristics. In their review of physical
stream protocols used by regulatory agencies,
Somerville and Pruitt (2004) recommended
the use of quantitative measures in physical
habitat assessments, where practicable, to
limit observer bias as much as possible.
The following subsections provide methods
for measuring physical habitat parameters in
headwater streams. We have attempted to
explain the ecological relevance of each
parameter and keep the methods as
straightforward as possible. Headwater
streams may be remote from roads or even
hiking trails, so many of the methods
described in the following sections use
minimal equipment. Rather than providing a
single method for measuring a parameter, we
have attempted to include multiple methods
from which the reader can choose based on
her/his particular needs and situations.
References
Allen, J. D. and A. S. Flecker. 1993.
Biodiversity conservation in running waters.
BioScience 43:32-43.
Barbour, M.T., J. Gerritsen, B.D. Snyder, and
J.B. Stribling. 1999. RapidBioassessment
Protocols for Use in Streams and Wadeable
Rivers: Periphyton, Benthic
Macroinvertebrates and Fish. 2nd edition,
EPA/841/B/98-010. Office of Water, U.S.
Environmental Protection Agency,
Washington, D.C.
Brussock, P. P., A. V. Brown, J.C. Dixon.
1985. Channel form and stream ecosystem
model s. Water Resources Bulletin 21:859-
866.
Fairfax County Stormwater Planning Division.
2003. Perennial Stream Field Identification
Protocol. Fairfax County Department of
Public Works and Environmental Services,
Stormwater Planning Division, Watershed
Planning and Assessment Branch. Fairfax,
Virginia.
Fitzpatrick, F. A., I. R. Waite, P. J. D'Arconte,
M. R. Meador, M. A. Maupin, and M. E.
Gurtz. 1998. Revised Methods for
Characterizing Stream Habitat in the
National Water-Quality Assessment
Program. U.S. Geological Survey Water-
Resources Investigations Report 98-4052,
Raleigh, North Carolina.
Lazorchak, J.M., DJ. Klemm, and D.V. Peck
(editors). 1998. Environmental Monitoring
and Assessment Program - Surface Waters:
Field Operations and Methods for
Measuring the Ecological Condition of
Wadeable Streams. EPA/620/R-94/004F.
U.S. Environmental Protection Agency,
Washington, D.C.
Montgomery, D.R. 1999. Process domains
and the river continuum. Journal of the
American Water Resources Association
36:397-410.
Montgomery, D.R. and L. H. MacDonald.
2002. Diagnostic approach to stream
channel assessment and monitoring. Journal
of the American Water Resources
Association 38:1-16.
27
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NC Division of Water Quality. 2005.
Identification Methods for the Origins of
Intermittent and Perennial streams, Version
3.1. North Carolina Department of
Environment and Natural Resources,
Division of Water Quality. Raleigh, NC.
http://h2o.enr.state.nc.us/ncwetlands/docum
ents/NC_Stream_ID_Manual .pdf
Ohio Environmental Protection Agency.
2002. Field Evaluation Manual for Ohio's
Primary Headwater Habitat Streams, Final
Version 1.0. Ohio Environmental Protection
Agency, Division of Surface Water,
Columbus, OH.
http ://www. epa. state, oh.us/dsw/wqs/headwa
ters/PHWHManual 2002 102402.pdf
O'Neill, R.V., D.L. DeAngelis, J.B. Waide,
and T.F.N. Allen. A Hierarchical Concept
of Ecosystems. Princeton Press, Princeton,
New Jersey, USA.
Platts, W.S., W.F. Megahan, and G.W.
Minshall. 1983. Methods for Evaluating
Stream, Riparian, andBiotic Conditions.
USDA Forest Service General Technical
Report INT-183.
Pond, G. J. and S. E. McMurray. 2002. A
Macroinvertebrate Bioassessment Index for
Headwater Streams of the Eastern Coalfield
Region, Kentucky. Kentucky Department for
Environmental Protection, Division of
Water, Water Quality Branch, Frankfort,
KY.
Sala, O. E., F. S. Chapin III, J. J. Armesto, E.
Berlow, J. Bloomfield, R. Dirzo, E. Huber-
Sanwald, L. F. Huenneke, R. B. Jackson, A.
Kinzig, R. Leemans, D. M. Lodge, H. A.
Mooney, M. Oesterheld, N. L. Poff, M. T.
Sykes, B. H. Walker, M. Walker, and D. H.
Wall. 2000. Global biodiversity scenarios
for the year 2100. Science 287:1770-1774.
Somerville, D.E. and B.A. Pruitt. 2004.
Physical Stream Assessment: A Review of
Selected Protocols for Use in the Clean
Water Act Section 404 Program. Prepared
for the U.S. Environmental Protection
Agency, Office of Wetlands, Oceans, and
Watersheds, Wetlands Division (Order No.
3W-0503-NATX). Washington, D.C. 213
pp.
Sparks, J., T. Hagman, D. Messer, and J.
Townsend. 2003 a. Eastern Kentucky
stream assessment protocol: utility in
making mitigation decisions. Aquatic
Resources News: A Regulatory Newsletter
2(2):4-10.
Sparks, J., J. Townsend, T. Hagman, and D.
Messer. 2003b. Stream assessment protocol
for headwater streams in the Eastern
Kentucky Coalfield Region. Aquatic
Resources News: A Regulatory Newsletter
USEPA. 2001. Protecting and Restoring
America 's Watersheds - Status, Trends, and
Initiatives in Water shed Management. EPA-
840-R-00-001.
Vannote, R.L., G.W. Minshall, K.W.
Cummins, J.R. Sedell, and C.E. Gushing.
1980. The river continuum concept.
Canadian Journal of Fisheries and Aquatic
Sciences 37 ': 130-137.
Wang, L., T. D. Simonson, and J. Lyons.
1996. Accuracy and precision of selected
stream habitat estimates. North American
Journal of Fisheries Management 16:340-
347.
28
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3.1 Designating hydrologic condition for
stream reaches
General
This subsection provides guidance for rapidly
designating hydrologic condition in headwater
stream reaches. The categories of hydrologic
condition (discussed in detail below) represent
the degree of departure from a spatially-
continuous flow (or conversely, a completely
dry condition) at a given point in time and
space. These designations describe the level
of connectivity or fragmentation of the aquatic
phase in headwater streams (Boulton 2003).
The degree of hydrologic connectivity is
fundamental in controlling the structure and
function of headwater streams because it
affects physicochemical properties, biotic
dispersal, and refuge availability (e.g.,
Boulton and Lake 1990, Dietrich and
Anderson 1998, Maltchik et al. 1994).
Hydrology of headwater stream reaches may
follow a predictable sequence of hydrologic
conditions related to seasonal (and/or greater
time frames) fluctuations in precipitation and
evapotranspiration. Shannon et al. (2002)
described hydrologic conditions in arid
ephemeral channels that occur at lower
frequencies than would occur in more humid
regions. At a given time, the hydrologic
condition also varies spatially within and
among headwater streams associated with
differences in distance to the groundwater
table, watershed vegetation, groundwater
storage capacity, etc.
The hydrologic designations discussed here
differ from those that represent general flow
regimes over time (e.g., perennial, intermittent
and ephemeral hydrology, Uys and O'Keeffe
1997). However, in the absence of continuous
monitoring of hydrologic condition,
designation of hydrologic conditions at least
once during wet and dry seasons may provide
a simple method for identifying flow regime
types.
Procedure
Hydrologic condition is determined by
visually assessing surface water connectivity
and water velocity within the thalweg of the
study reach. Designation should be based
upon the predominant hydrologic condition
within the study reach. Mark the appropriate
box on the field forms for the hydrologic
condition identified (Figure 3-1).
STUDY REACH HYDROLOGIC CONDITION
D Surface flow continuous (4)
D Flow only interstitial (3)
D Surface water present but no visible flow (2)
D Surface water in pools only (1)
D No surface water (0)
Figure 3-1 Appropriate location for
recording hydrologic condition on page 1 of
field forms.
The text below describes five categories of
hydrologic condition seen in headwater
streams. Each category is represented by
photos and a diagram showing a longitudinal
section along the channel thalweg. Blue
shading indicates surface water, arrows
indicate presence and direction of visible flow,
coarse stone substrate on the streambed is
represented by solid brown, and the hatched
brown areas indicated finer streambed
substrate and underlying geology. The term
"habitat units" refers to riffles and pools, the
dominant habitat types in headwater streams.
The five hydrologic categories and their
numerical descriptors are as follows:
29
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Visible surface flow continuous (4):
Surface water is flowing and
uninterrupted between habitat units
and flowing. Most of the streambed
stones within the thalweg are
submerged.
30
-------
- Visible flow interstitial (3): Surface
water is interrupted between habitat units,
such that the majority of streambed stones in
shallow habitat units (i.e., riffles) are
exposed. However, interstitial flow
connecting habitat units is evident as trickles
or rivulets flowing between stones or visible
at the tail and heads of pools. Soluble
tracers, such as fluorscein dye or NaCl
solution may be added at the upstream end
of a study reach and monitored downstream
to determine if interstitial flow connects
pools within a reach.
-------
- Surface water present but no visible flow
(2): Surface water is uninterrupted between
habitat units, however there is no evidence
that the water is flowing throughout study
reach. Water standing in pools may appear
stagnant. This condition is likely to occur in
low gradient headwater streams rather than in
high gradient streams.
32
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Surface water present in pools only
(1): Surface water is found only in
pools and there is no visible water or
flow connecting pools. Stream bed
sediments between pools may be
moist.
33
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No surface water (0): Surface water is absent from the channel thalweg.
References
Boulton, A. J. 2003. Parallels and contrasts
in the effects of drought on stream
macroinvertebrate assemblages. Freshwater
5/o/ogy 48:1173-1185.
Boulton, AJ. and P.S. Lake. 1990. The
ecology of two intermittent streams in
Victoria, Australia. I. multivariate analyses
of physicochemical features. Freshwater
Biology 24:123-141.
Dietrich, M. andN. H. Anderson. 1998.
Dynamics of abiotic parameters, solute
removal and sediment retention in summer-
dry streams of western Oregon.
Hydrobiologia 379:1-15.
Maltchik, L., S. Molla, C. Casado, and C.
Montes. 1994. Measurement of nutrient
spiraling in a Mediterranean stream:
comparison of two extreme hydrological
periods. Archivfur Hydrobiologie 130:215-
227.
Shannon, J., R. Richardson, and J. Thornes.
2002. Modelling event-based fluxes in
ephemeral streams. Pages 129-172 inL. J.
Bull and M. J. Kirkby (editors). Dryland
rivers: hydrology and geomorphology of
semi-arid channels. John Wiley & Sons,
Chichester, United Kingdom.
Uys, M.C. and J.H. O'Keeffe. 1997. Simple
words and fuzzy zones: early directions for
temporary river research in South Africa.
Environmental Management 21:517-531.
Equipment and supplies
Measuring tape (50 m)
Field forms
34
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3.2 Continuous monitoring ofhydrologic
condition
General
This subsection describes a water sensor for
continuously monitoring hydrologic condition
(i.e., presence or absence of water) that is
economical, light weight, and easy to install.
The sensor described provides information
regarding the timing, duration, and frequency
of channel drying. Other methods such as
float gages and pressure transducers with data
loggers, which are widely used to
continuously measure stream stage and
subsequently, discharge (Rantz et al. 1982),
also provide flow permanence data, but can be
more costly and require more channel
modification and maintenance.
Water sensors may be assembled by
Intermountain Environmental, Inc (IEI)11.
The components of the water sensor include
an Onset Hobo® state data logger, Onset
submersible case, and an encased cable (see
Figure. 3-2, pen shown for scale). The state
data logger was designed to continuously
record binary changes (i.e., open vs. closed;
on vs. off). The modification by IEI has
allowed this data logger to record the timing
and frequency of changes in hydrology (in
terms of presence and absence of water).
When present, water completes the circuit
between the two exposed copper wires on the
contact end of the cable and sends a "closed"
signal to the sensor. When a stream dries and
water no longer is present to complete the
circuit, the data logger records an "open"
signal. The datalogger does not record on
time intervals, and only records the time when
a change of state occurs. The data logger can
1 Intermountain Environmental, Inc.
601 W. 1700 S. Suite B., Logan, UT 84321
(800) 948-62361
Figure 3-2 Primary components of a water
sensor used to continuously monitor
hydrologic condition.
record up to 2000 state changes (checking for
changes of state every 0.5 seconds) and the
battery will last approximately 1 year.
3.2.1 Launching and preparing for
deployment
Procedure
Install the appropriate Onset software onto a
personal computer. (Note that to launch data
loggers via personal laptop computers Onset
Boxcar Pro 4.3® or higher may be required.)
Connect the PC interface cable to an open
Com Port or serial port of the computer and
the 3.5 mm j ack of the data logger. Open the
Onset Boxcar® program and either select
Launch from the Logger menu or select the
icon for launching on the tool bar. A launch
dialog box should appear with setting options
(Figure. 3-3). Note the condition of the
battery; if it does not indicate that the battery
is "good" then close the launch dialog box and
change the battery (CR-2032 lithium). Under
description, type locality information (e.g.,
Hoosier Natl. For.) and change text for "close"
and "open" to "wet" and "dry", respectively.
Do not select "wrap around" (this overwrites
data already stored when >2000 state changes
occur) or "stealth mode" (this turns off
35
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HOBO State iCn93G ONSEt Compute Corp
Dai* I
Current Slate: DRY
CkKeTextfU Sp (WET
OpenTexMQl f| JDRY
Battery: Good
i J~ Wrap around when fiJ (overwrite ddest data)
1 r SreaKhMode
JT Deiai>ed Start:
Figure 3-3 Launch dialog box for Onset Boxcar Pro.
indicator lights while data loggers are
launched). Delayed setting may be selected,
particularly when personal computers are not
accessible near field sites and travel time to
field sites may affect the battery life of data
loggers. Select "start" and allow the launch
progress bar to completely extend before
disconnecting the data logger from the PC
interface cable.
Note that the 6-digit serial number displayed
by the BoxCar software matches the serial
number printed on the data logger. Write this
number on the outside of the submersible case
using a permanent marker.
Note that the red LED (indicating "open" or
"dry" state, see Figure 3-4) should be blinking
if the data logger is properly launched. Place
the data logger in its respective submersible
case. Next connect the 2.5 mm cable to the
2.5 mm jack on side of data logger and place
two desiccant packs inside the submersible
case (Figure 3-4).
Figure 3-4 Desiccant packs and Onset
Hobo® State data logger with jacks and
LEDs shown.
Inspect the rubber O-ring and its seating on
the submersible case (below the threads,
Figure 3-5), making sure that the surfaces are
clean and there are no cracks or damage to the
O-ring. The O-ring should be replaced if there
is cracking or damage. Lubricate the entire
surface of the O-ring using the silicone
compound by applying a thin, even coat.
36
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Place the O-ring in its seating on the
submersible case and screw on the
submersible case cap. Ensure that the O-ring
seats properly and does not extrude when
screwing cap in place.
3.2.2 Deployment
Always make certain that the data logger stays
dry. Record the location of the site using
preferably GPS coordinates (e.g., latitude and
longitude, UTMs) or precise directions. These
directions should include road names,
compass headings, turn directions and
distances.
Figure 3-5 Water sensor with 2.5 mm
cable, O-ring and seat shown.
Select a location within the channel thalweg
that is the approximate average water depth of
the thalweg for the entire 30m reach. Where
available, select a location that also has a steep
bank; this will help to keep the data logger end
of the water sensor dry during high flows.
Use a small sledge hammer to drive a section
of rebar or a stake into the stream bed. Make
certain that the rebar is firmly embedded.
Select a water sensor with the appropriate
encased cable length to extend from the
streambed stake to a safe bank location.
Assemble a stilling well by attaching PVC
pipe to a PVC cap (Figure. 3-6). Three to four
holes should be drilled into the bottom of the
cap to allow the water level within the stilling
well to fluctuate with the stream water level
(Figure. 3-6). This stilling well will prevent
false readings associated with debris
accumulating on the contact wires. The
bottom of the stilling well is positioned so it is
flush with the stream bed. An O-ring (#10,
l/2" inner diameter) may be used to seal out
rain from the stilling well opening around the
flexible cable housing. Place the contact end
of the sensor inside the stilling well so that the
contact wires are a few millimeters above the
PVC cap.
Flexible cable
jf housing
O-ring
3/4" PVC
PVC cap
w/ holes
drilled on
bottom t
Hose clamps
Sensor
- contact inside
PVC
^•r- Rebar
Figure 3-6 Schematic showing assembly of
stilling well and contact end of water
sensor.
37
-------
Attach the contact end of the water sensor to
the stake with 2 hose clamps or cable ties
(above and around stilling well), making
certain that they are tight (Figure 3-7). Extend
the water sensor cable laterally to the bank,
allowing the cable to conform to the contour
of the channel and bank. Insert the second
piece of rebar or stake by pounding it into the
soil adjacent to the stream channel, making
certain that it is firmly embedded.
Figure 3-7 Water sensor securely attached
to rebar above and below stilling well.
Attach the data logger end of the water sensor
to the rebar with 2 cable ties. Gently place
large cobble on top of cable to stabilize and
camouflage the water sensor (Figure 3-8).
Unscrew the cap of the submersible case and
note which light is blinking. Where the
contact end is submersed in water the green
LED ("closed" or "wet" state) should be
blinking. If the contact end is not submersed
in water the red LED ("open" or "dry" state)
Figure 3-8 Water sensor positioned for
continuous monitoring of hydrologic
condition. Meter stick shown for scale.
should be blinking. If this is not occurring
then check the connection of the 2.5 mm cable
and jack. If this does not remedy the situation
then replace the data logger or water sensor
with another.
Record the data logger serial number,
location, date, time, water depth (using a
meter stick) and hydrologic condition for each
sensor deployed. The predominant hydrologic
condition of the study reach is categorized as:
1) visible surface flow continuous, 2) visible
flow interstitial, 3) surface water present but
no visible flow, 4) surface water in pools only
and 5) no surface water.
3.2.3 Retrieval of the water sensor
Using the field sheets or notebook completed
during deployment, return to the study reach
within 1 year after the water sensor was
installed. For each water sensor retrieved
record the data logger serial number, location,
date, time, water depth (using a meter stick)
and hydrologic condition. Remove the rebar
38
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and water sensor from the water. Use clippers
to cut the cable ties to disconnect rebar pieces
from the data logger and contact ends.
3.2.4 Transferring data
Remove data logger from submersible case
and connect PC interface cable to 3.5 mm jack
on data logger. Open the Onset Boxcar®
program and either select "readout" from the
Logger menu or select the icon for readout on
the tool bar. You will then be asked to save
the logger datafile (*.dtf). The serial number
of the data logger and year should be used to
name the file. For example, if the data logger
serial number is 682537 and data were
collected from 15 April to 23 September 2004
then the file is named "682537_04.dtf'. This
will be a unique file name that can then be
linked to field sheets or notebook for further
site description. These files can then be saved
within folders representing each stream.
From the File menu select "export" and the
desired spreadsheet program (e.g., Microsoft
Excel®, Lotus 1-2-3). You will then be asked
to save the text file (*.txt). Use the same
name given to the *.dtf file, but with the *.txt
suffix.
Open the spreadsheet program and open a file
containing water sensor data. The Text
Import Wizard window should open. Select
file type marked "delimited" and select
"next". In the next window select "tab" as the
delimiter and select "finish". This should then
separate date + time, and hydrologic state into
2 separate columns. The number of data rows
minus 1 should indicate the number of
hydrologic state changes occurring over the
period between launching and readout. Some
of the state changes at the beginning and end
of the data set may not represent the
hydrologic changes at the study site. Using
the date, time and hydrologic condition data
from field sheets or notebook, the actual
starting and ending time is entered into the
columns. When entering the starting and
ending date and time, enter each into the
spreadsheet as single cells with a space
between the date and time. For example, if
the water sensor was actually deployed at 1:24
pm on 15 April, enter the date & time as
follows: 4/15/03 1:24 PM. Then highlight the
cell and change its cell format to the "custom"
category and the "mnrss.O" type. Be sure to
also enter the hydrologic state ("wet" or
"dry") for the starting and ending periods in
the appropriate column. Below the cell that
identifies the data logger serial number, enter
the site name including stream name and site
number.
To calculate the number of hours (duration)
that had occurred between each state change
use the following function in the column
adjacent to the column containing hydrologic
state labels. Type "= (A4-A5)*24. This
example subtracts the date+time in cell A5
from a previous date+time in cell A4.
Continue this down the column until the all
durations are calculated. These can then be
easily converted from hours to days by
dividing the number of hours by 24. The total
duration of dry or wetted condition can then
be determined by summing every other cell
within the column.
References
Onset Computer Corporation. 1999. HOBO®
State User's Manual.
Onset Computer Corporation. Directions for
Protective Submersible Case for Onset Data
Loggers
Rantz, S.E. and others. 1982. Measurement
and computation of stream/low. USGS
Water Supply Paper 2175.
Equipment and supplies
39
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Water sensor
Field notebook or field forms
Pencil
Map of area
Metal stakes or rebar (2 per sensor)
Mallet or small sledge hammer
Watch
Hose clamps or cable ties (4 per sensor)
Personal computer (PC) with operating system
that can support data logger software
Onset software Boxcar® 3.0+ or any version
of Boxcar® Pro
PC interface cable (w/ 3.5 mm jack and serial
port)
Submersible case kit (rubber O-ring, 2
dessicant packs, and tube of silicone
compound)
Global Positioning System (GPS) unit
Meter stick
3.3 Identifying the channel head
General
This subsection provides instructions for
identifying and recording the location of the
channel head or channel origin of streams.
Headwater streams link valley hillslopes to
downstream water bodies through the
downstream transfer of sediment and organic
matter (Gomi et al. 2002, Hutchens and
Wallace 2002). The channel head or origin is
the upstream boundary between hillslopes and
channels in the landscape, specifically
between the valley head and channel.
Characteristics of the surrounding valley (e.g.,
slope, geology and land use) determine the
evolution of channels and therefore the
location of channel heads (Dietrich and Dunne
1993, Montgomery 1999). The channel head
rarely extends to the valley divide, so the
valley network envelopes the channel
network. Swales, hollows, and zero-order
basins are other names used to describe
hillslope landforms that drain into channel
heads. These are located upslope from
channel heads (Dietrich and Dunne 1993).
The transition from hillslope to channel may
be abrupt, in the form of headcut or step, or
gradual (Figure 3-9). The channels emerging
from zero-order basins have been called
transitional channels and are often ephemeral
or intermittent (Gomi et al. 2002).
Transitional
channel
Figure 3-9 Drawing showing a valley
hillslope (swale or hollow) relative to
channel. Valley head (A), gradual (B) and
abrupt (C) channel heads are identified.
Gray areas indicate zero-order basins
draining into channel heads. Redrawn
from Dietrich and Dunne (1993).
The following procedure will provide field
characteristics that can be used regardless of
hydrologic status of a site. The observation of
surface flow may not be the best indicator
when defining whether a landform is or is not
a channel. Surface runoff (Horton overland
flow) and throughflow-return flow may be
apparent on hillslopes, and are thus, not
restricted by channel formation (Fetter 1988,
Dietrich and Dunne 1993). Additionally, the
distribution of surface flow in stream
networks expands and contracts with water
table fluctuations (Blythe and Rodda 1973,
Stanley etal. 1997). Hydrologic permanence
at the channel head may be dependent upon
40
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underlying geology and connectivity to
groundwater. Springs or seeps originating
from contact zones, faults, joints and fractures
in the underlying geology can coincide with
and/or control channel head location (Higgins
and Coates 1990). The flow of these springs
may be continuous or discontinuous over time.
The resolution of most topographic maps is
too low to reveal the extent of headwater
channels (e.g., Hansen 2001). Therefore, the
terminations of blue lines (e.g. on USGS 1:24
000 quads) do not accurately represent
channel heads (Mark 1983). Typically
channel heads are located upslope from blue
line terminations, extending into the contour
line crenulations (see Figure 2-2).
The location of the channel head is recorded
once for a given stream during the study
because it is unlikely to change significantly
over the timeframe of most monitoring studies
(1-2 years). However, channel head location
can shift depending upon characteristics of the
surrounding hillslope (e.g., gradient, soil
cohesiveness, land use) and stochastic events
(e.g., mass failures). The channel head is a
particularly sensitive feature in arid and semi-
arid landscapes, where gully erosion caused
by unstable channel heads is a serious socio-
economic and environmental problem (Bull
and Kirkby 2002). Infilling by debris flows
and landslides can move the channel head
downslope, whereas gullying or headcutting
moves the channel head upslope (Benda and
Dunne 1987, Miller et al. 2003). Therefore,
over long time frames (10s to 100s of years),
the position of the channel head may fluctuate
in response to these processes.
Procedure
Hike the channel upstream of the "ephemeral"
or upstream-most study reach (see Section 2.1
for description of study reach selection). You
should focus on characteristics of the
streambed and banks relative to the adjacent
hillslope. The phrase "definable bed and
banks" is often used to determine if a land
form is a stream channel. Problematically,
this phrase is not easily defined in objective
terms although along larger streams and rivers
it is visibly obvious. A channel is a landform
that conveys water and sediment between
banks. Banks are relatively narrow zones that
have steeper gradients than adjacent hillslopes
and the transverse slope of the channel bed
(Dietrich and Dunne 1993).
Characteristics of abrupt channel heads
Abrupt channel heads appear as steep vertical
steps from the valley head down to the
channel (Figure 3-10). These abrupt steps are
also known as "knickpoints" or "headcuts".
No evidence of bank or channel forms is
usually visible above abrupt channel heads.
Figure 3-10 An abrupt channel head in
Wayne National Forest, OH.
41
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Thus, the abrupt channel head represents a
distinct start of continuous streambed and
banks in the downstream direction. These
abrupt changes often correspond to differences
in surface sediment between the valley head
and channel. Surface sediments above the
valley head will be of colluvial origin (e.g.,
transported by gravity from adjacent
hillslopes) and/or have soil nature (e.g., humus
layer). In contrast surface sediment in the
channel will be of a mixture of recently
deposited colluvium and weathered material
exposed from surface flow (e.g., bedrock and
boulders). Vegetation type and density may
also differ up- and downslope of the channel
head. Terrestrial vegetation may be sparse or
absent in the channel below the channel head
compared to the upstream valley head. Be
aware that steep vertical steps and headcuts
are not restricted to channel heads and may
occur within continuous channels. In this
case, definable bed and banks are clearly
evident upstream of the headcut (see Section
3.4). Record coordinates (latitude &
longitude) and description of the hydrologic
condition at the channel head in the Notes
section for the datasheet of the nearest study
reach.
Characteristics of gradual channel heads
Gradual channel heads are less distinct than
abrupt channel heads. These are characterized
by a more gradual or discontinuous transition
in bank and bed features, rather than the
obvious boundary of a step or headcut. As
you approach the channel head, the height and
angle of the banks decline. The defined bed
and banks are often discontinuous and may be
interrupted by debris dams, tree roots, or
bedrock outcropping. For the purposes of this
study, we define the channel head as the point
where the channel no longer has continuous
defined bed and banks. Be aware that steep
channels can have a step-pool or cascade
structure and appear less continuous than
riffle-pool reaches (Church 1992). Banks
typically are less well defined at the "steps"
compared with "pools" in these streams.
However, the channels should be considered
continuous if the steps are composed of
visibly eroded material exposed from surface
flow (e.g., bedrock and boulder) that may or
may not be covered with moss and organic
debris piles (Figure 3-11).
Figure 3-11 Views from gradual channel heads in east-central Kentucky. A) Looking
upslope toward the valley head from the channel head position. B) Looking downslope at
the cascade structure of the transitional channel.
42
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References
Benda, L. and T. Dunne. 1987. Sediment
routing by debris flow. Pages 213-223 in R.
L. Beschta, T. Blinn, G. E. Grant, F. J.
Swanson, and G. G. Ice (editors).
International symposium on erosion and
sedimentation in the Pacific rim.
International Association of Hydrological
Sciences Press, Institute of Hydrology,
Wallingford, United Kingdom.
Blythe, K. and J. C. Rodda. 1973. A stream
length study. Water Resources Research
9:1454-1461.
Bull, L. J. and M. J. Kirkby. 2002. Channel
heads and channel extension. Pages 263-298
in L. J. Bull and M. J. Kirkby (editors).
Dryland rivers: hydrology and
geomorphology of semi-arid channels. John
Wiley & Sons, Chichester, United Kingdom.
Church, M. 1992. Channel morphology and
typology. Pages 126-143 in P. Calow and G.
E. Petts (editors). The rivers handbook:
hydrological and ecological principals
Volume I. Blackwell Scientific, Boston,
Massachusetts.
Dietrich, W. E. and T. Dunne. 1993. The
channel head. Pages 175-219 in K. Beven
and M. J. Kirkby (editors). Channel network
hydrology. John Wiley & Sons, Chichester,
United Kingdom.
Fetter, C. W. 1988. AppliedHydrogeology,
2nd edition. Merrill Publishing Co.,
Columbus, Ohio.
Gomi, T., R. C. Sidle, and J. S. Richardson.
2002. Understanding processes and
downstream linkages of headwater systems.
BioScience 52:905-916.
Higgins, C. G. and D. R. Coates (editors)
1990. Groundwater Geomorphology: The
Role of Subsurface Water in Earth-Surface
Processes andLandforms. Geological
Society of America, Boulder, Colorado.
Hutchens, J. J. and J. B. Wallace. 2002.
Ecosystem linkages between southern
Appalachian headwater streams and their
banks: leaf litter breakdown and invertebrate
assemblages. Ecosystems 5:80-91.
Mark, D. M. 1983. Relations between field-
surveyed channel networks and map-based
geomorphometric measures, Inez, Kentucky.
Annals of the Association of American
Geographers 73:358-372.
Miller, D., C. Luce, and L. Benda. 2003.
Time, space, and episodicity of physical
disturbance in streams. Forest Ecology and
Management 178:121-140.
Montgomery, D. R. 1999. Erosional
processes at an abrupt channel head:
implications for channel entrenchment and
discontinuous gully formation. Pages 247-
276 in S. E. Darby and A. Simon (editors).
Incised river channels. John Wiley & Sons,
Chichester, U.K.
Stanley, E. H., S. G. Fisher, andN. B. Grimm.
1997. Ecosystem expansion and contraction
in streams. BioScience 47:427-435.
Equipment and supplies
GPS unit
Topographic map
Field forms
3.4 Identifying channel headcuts
General
This section provides instructions for the
identification of channel headcuts in
43
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headwater streams. Readouts are abrupt
changes in streambed elevation (i.e.,
knickpoint) that migrate in an upstream
direction (Leopold et al. 1964). This
migration is a natural geomorphic process that
is often accelerated due to human
modification of the channel and/or
surrounding watershed (Patrick et al. 1994,
Montgomery 1999). The upstream migration
of headcuts results in downcutting (i.e.,
degradation) of the streambed and incised
channel morphology (Galay 1983, Simon
1989). Among the ecological effects
downstream of headcuts may be loss of
streamside vegetation, scoured streambeds,
decreased sinuosity, and temporary increase in
downstream sedimentation (Patrick et al.
1994). Headcuts can also influence the
connectivity along headwater streams by steep
changes in streambed elevation and
hydrology. Abrupt changes in summer
baseflow hydrology (and water temperature)
occur at headcuts and are related to
differences in distance from the groundwater
table. As the summer groundwater table
lowers (lower precipitation, higher
evapotranspiration), it falls below the
streambed upstream of the headcut before
dropping below the stream bed downstream of
the headcut. This causes flow to remain for
longer periods downstream (often perennially)
than upstream of headcuts. The presence of
headcuts is determined once for a given reach
during the study because their presence is
unlikely to change significantly over short
time periods (e.g., 1-2 y), however any
upstream advance should be noted.
Procedure
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg. Survey the study reach for abrupt
changes in streambed elevation. If a
knickpoint is located, determine first whether
the formation is simply a natural grade control
point (e.g., large boulders, bedrock outcrops,
or large woody debris). If it is not, then look
for the following: 1) undercutting beneath the
headcut face or headwall (Figure 3-12), 2)
seepage or piping from the headwall, and 3)
alluvial fan or deposits in the channel
downstream of the headcut. Be aware that
headcuts may stall their upstream migration at
grade control features between large floods.
Upstream
streambed
Headwall
A.
Downstream
streambed
Dry
streambed
B.
Seepage from headwall
Wetted streambed
Figure 3-12 Longitudinal view of a headcut, (A.) Blue arrows illustrate flowpaths that lead
to undercutting, failure of the headwall, and eventually upstream migration of the headcut;
(B.) Abrupt change in summer baseflow hydrology at a headcut.
Indicate on the field form (Figure. 3-13) the
presence or absence of a headcut within the
study reach. Note location of headcut on
study reach drawing and make notes
characterizing the formation. Photographs of
headwater streams with headcut formations
are shown in Figures 3-14, 3-15 and 3-16.
44
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PRESENCE OF
HEADCUT IN REACH
Y N
ALGAL COVER INDEX
1 I1'2 2 3 4 5
# CORES FOR SUBSTRATE MOISTURE
(depositional)
Figure 3-13 Portion of page 1 of field forms showing the cell for recording presence of
channel headcuts.
Figure 3-14 Subtle headcut in Falling Rock
Creek in east-central KY (looking
upstream).
Figure 3-15 Huge headcut (~2 m change in
bed elevation) in an unnamed stream in
Athens, GA (looking upstream).
Figure 3-16 Headcut in Taylor Branch in south-central IN (looking downstream), where
streambed elevation at the arrow was ~ 1 m higher than streambed below headcut at the
yellow circle.
45
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References
Galay, V. J. 1983. Causes of riverbed
degradation. Water Resources Research
19:1057-1090.
Leopold, L.B., M.G. Wolman, and J.P. Miller.
1964. Fluvial Processes in Geomorphology.
W.H. Freeman & Co. San Francisco, CA,
USA.
Montgomery, D.R. 1999. Erosional
processes at an abrupt channel head:
implications for channel entrenchment and
discontinuous gully formation. Pages 247-
276 in S.E. Darby and A. Simon (editors).
Incised river channels: processes, forms,
engineering, and management. John Wiley
& Sons, Chichester, U.K.
Patrick, D.M., S.T. Ross, and P.O. Hartfield.
1994. Fluvial geomorphic considerations in
the management and stewardship of fluvial
ecosystems. Pages 90-99 in Riparian
ecosystems in the humid U.S.: Functions,
Values and Management. National
Association of Conservation Districts,
Washington, D.C.
Simon, A. 1989. A model of channel
response in disturbed alluvial channels.
Earth Surface Processes andLandforms
14:11-26.
Equipment and supplies
Measuring tape
Field form
3.5 Measuring channel sinuosity
General
This section provides instructions for rapidly
scoring channel sinuosity of headwater
streams. This procedure is similar to that used
by the Ohio Environmental Protection Agency
(Ohio EPA 2002), where sinuosity is
described as the number of well-defined bends
or meanders over a distance of stream
channel. This differs from the more
quantitative measure, sinuosity index, which is
the ratio of the channel thalweg distance to the
downvalley distance (Gordon et al. 1992,
Platts et al. 1983, Rosgen 1996). In
association with other measures (e.g., channel
slope, substrate particle size), sinuosity
provides useful information regarding the
degree of channel modification to headwater
streams. Retention of nutrients and organic
matter increases with increasing sinuosity,
ensuring transformations that may be
beneficial for downstream waters (e.g.,
Giicker and Boechat 2004, Muotka and
Laasonen 2002). Sinuosity is measured once
for a given reach during the study because it is
unlikely to change significantly over short
time periods (e.g., 1—2 years).
Procedure
Delineate the 30-m study reach so the
Measuring tape is positioned along the
thalweg. Sinuosity is based on the number of
well-defined bends over the 30-m study reach
(approximately 20X the bankfull width of
most headwater streams). Examples showing
various degrees of sinuosity are shown in
Figure 3-17. On the first page of the field
form indicate the sinuosity in the appropriate
cell (Figure 3-18).
46
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0 0.5
1.5
2.5
30m -
Figure 3-17 Examples of stream channels varying in sinuosity (number of bends) along 30-
m study reaches.
MAX. POOL DEPTH (cm)
DEPTH TO BEDROCK /
GROUNDWATER (m)
(3 measures in depositional habitat)
^^
^^
^^
SINUOSITY
(number of bends)
Figure 3-18 Portion of page 1 of field forms showing the cell for recording channel
sinuosity.
References
Gordon, N.D., T.A. McMahon, and B.L.
Finlayson. 1992. Stream Hydrology: An
Introduction for Ecologists. John Wiley &
Sons, Chichester, United Kingdom.
Gucker, B. and I. G. Boechat. 2004. Stream
morphology controls ammonium retention in
tropical headwaters. Ecology 85:2818-2827.
Platts, W.S., W.F. Megahan, and G. W.
Minshall. 1983. Methods for evaluating
stream, riparian, and biotic conditions.
USDA Forest Service General Technical
Report INT-183.
Muotka, T. and P. Laasonen. 2002.
Ecosystem recovery in restored headwater
streams: the role of enhanced leaf retention.
Journal of Applied Ecology 39:145-156.
Ohio EPA. 2002. Field evaluation manual
for Ohio's primary headwater habitat
streams. Final Version 1.0. Ohio
47
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Environmental Protection Agency, Division
of Surface Water, Columbus, Ohio.
http ://www. epa. state, oh.us/dsw/wqs/headwa
ters/PHWHManual 2002 102402.pdf
Rosgen, D.L. 1996. Applied river
morphology. Wildland Hydrology, Pagosa
Springs, CO.
Equipment and supplies
Measuring tape
Field forms
3.6 Designating habitat units
General
This subsection provides instructions for
identifying habitat or channel units within
headwater stream reaches. Habitat units (or
"meso-habitats") are distinct channel units
having characteristic physical properties.
They are smaller than stream reaches and
larger than microhabitats, according to the
hierarchical levels used to describe the
physical template of streams (Frissell et al.
1986). Within headwater streams with
moderate to high gradient (slope > 2 %),
habitat units can range from <1 to 10 m in
linear stream length (K. M. Fritz, personal
observation). Habitat units in sandy, low-
gradient or bedrock-dominated channels may
be > 10 m long. These units are found
longitudinally along the channel and may be
spaced at fairly regular intervals along a
stream reach (Leopold et al. 1964, Beschta
and Platts 1986). Habitat units are delimited
by elevational and lateral changes of the
streambed (Hawkins et al. 1993). This is
particularly evident in streams where the
streambed particles are not primarily sand or
silt (Leopold et al. 1964). Associated with
these distinct channel units are characteristic
water flow and depth regimes. Therefore,
physical variation within a study reach can be
accounted by the proportions of these habitat
types. In many instances these characteristics
lead to differences in the dominant streambed
particle sizes among types of habitat units.
Assessment and restoration of streams are
typically limited to the reach scale. However,
for logistical reasons, biological communities
are often sampled at spatial scales below the
reach level (Cuffney et al. 1993, Lazorchak et
al. 1998, Barbour et al. 1999), often stratified
by habitat type. Inter-habitat variability in
ecological measures can exceed variation seen
among reaches or streams (e.g., Angradi 1996,
Rabeni et al. 2002). Therefore, quantifying
the extent of habitat types within stream
reaches is fundamental to understanding the
ecological status of water resources at larger
spatial scales, not because of the inherent
measurement of habitat units (Poole et al.
1997) but to put other measures in context for
comparison.
The number of the physical parameters needed
for designating habitat types increases as
classification become more complex. The
utility of a complex classification becomes
limited because the variety of habitat types
that can be identified within stream reaches
can vary greatly among regions. To be useful,
the categories of habitat type need to be
applicable for all reaches examined in a study.
In addition, as the specificity of habitat types
increases there is typically a greater level of
subjectivity involved in their designation
(Roper and Scarnecchia 1995). The following
procedure provides guidance to delimit the
most basic categories of habitats within
headwater streams (see Hawkins et al. 1993
and Lazorchak et al. 1998 for descriptions of
finer levels of habitat types). These include
erosional and depositional habitats (Moon
1939). Erosional habitats are identified as
shallow areas with rapid flow and typically
coarse streambed substrate. They include
such habitats as riffles, fast runs, sheets,
48
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cascades and steps (in step-pool reaches). In
contrast, depositional habitats are deeper areas
with little or no visible flow and typically
have fine streambed substrates but may also
be bedrock. They include such habitats as
pools and slow runs. Because water flow and
depth are primary parameters used to
designate habitat type and these can vary
seasonally, this procedure should be carried
out during each sampling period.
Procedure
Delineate the 30-m study reach so that the
measuring tape is marking transects along the
thalweg from downstream to upstream. At
each meter mark along the thalweg of the
study reach (0, 1, 2,.. .30m) assess water flow,
water depth and substrate type to designate
whether the habitat is erosional or
depositional. The dotted line represents the
study reach thalweg and the black arrow is
pointing in the direction of flow in Figure 3-
19.
30m
Figure 3-19 Plan view of study reach (top) and picture showing series of alternating
erosional and depositional habitats along a headwater stream.
On the second page of the field form mark the
habitat type for each transect (Figure 3-20).
There is also a column on the field form for
notes concerning the presence of large woody
debris (LWD, diameter > 10 cm), leaf packs,
bryophytes, herbaceous vegetation, etc. within
the thalweg at that meter mark (Figure 3-20).
49
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The designation of habitat type relies more on
the streambed characteristics where the stream
is dry. Substrate size, streambed elevation and
the distribution of organic matter are useful in
determining habitat type at locations along dry
channels.
* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2BF Page 2 of 4
Meter
#
0
i
2
3
Modal
Sediment
Particle
Size
(mm)*
Water
Depth
(cm)*
Habitat Notes
Type (e.g., LWD,
(E/D) Leafpack)
E Leafpack
E
D LWD
D
Velocity
(m/s)*
Wetted
Width
(m)
BE
Width
(m)
BE
Depth
(m)
ft
EPA
width
(m)§
Figure 3-20 Appropriate location for recording habitat units and notes on Page 2 of the
Field Forms.
References
Angradi, T.R. 1996. Inter-habitat variation in
benthic community structure, function, and
organic matter storage in 3 Appalachian
headwater streams. Journal of the North
American Benthological Society 15:42-63.
Barbour, M.T., J. Gerritsen, B.D. Snyder, and
J.B. Stribling. 1999. RapidBioassessment
Protocols for Use in Streams and Wadeable
Rivers: Periphyton, Benthic
Macroinvertebrates and Fish. Second
edition, EPA/84l/B/98-010. Office of
Water, U.S. Environmental Protection
Agency, Washington, D.C.
Beschta, R. L. and W. S. Platts. 1986.
Morphological features of small streams:
significance and function. Water Resources
Bulletin 22:369-379.
Cuffney, T.F., M.E. Gurtz, and M.R. Meador.
1993. Methods for Collecting Benthic
Invertebrate Samples as Part of the National
Water Quality Assessment Program. U.S.
Geological Survey Open File Report 93-406.
Water Resources Division, United States
Geological Survey, Raleigh, North Carolina,
USA.
Frissell, C.R., W.J. Liss, C.E. Warren, and
M.D. Hurley. A hierarchical framework for
stream habitat classification: viewing
streams in a watershed context.
Environmental Management 10:199-214.
Hawkins, C.P., J.L Kershner, P. A. Bisson,
M.D. Bryant, L.M. Decker, S.V. Gregory,
D.A. McCullough, C.K. Overton, G.H.
Reeves, RJ. Steedman, and M.K. Young.
1993. A hierarchical approach to classifying
stream habitat features. Fisheries 18:3-12.
Lazorchak, J.M., DJ. Klemm, and D.V. Peck
(editors). 1998. Environmental Monitoring
and Assessment Program - Surface Waters:
Field Operations and Methods for
Measuring the Ecological Condition of
Wadeable Streams. EPA/620/R-94/004F.
U.S. Environmental Protection Agency,
Washington, D.C., USA.
50
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Leopold, L.B., M.G. Wolman, and J.P. Miller.
1964. Fluvial Processes in Geomorphology.
W.H. Freeman & Co. San Francisco, CA,
USA.
Moon, H. P. 1939. Aspect of the ecology of
aquatic insects. Transactions of the Society
for British Entomology 6:39-49.
Poole, G.C., C.A. Frissell, and S.C. Ralph.
1997. In-stream habitat unit classification:
inadequacies for monitoring and some
consequences for management. Journal of
the American Water Resources Association
33:879-896.
Rabeni, C.F., K. E. Doisy, and D.L. Galat.
2002. Testing the biological basis of a
stream habitat classification using benthic
invertebrates. Ecological Applications
12:782-796.
Roper, B.B. and D.L. Scarnecchia. 1995.
Observer variability in classifying habitat
types in stream surveys. North American
Journal of Fisheries Management 15:49-53.
Equipment and supplies
Measuring tape
Field forms
3.7 Measuring channel slope
General
The following subsection provides methods
for measuring channel slope or gradient in
headwater streams. Channel slope is the drop
in elevation per unit length of channel ("rise-
over-run", Figure 3-21). Slope is an important
variable because it determines the velocity,
stream power, and tractive forces which shape
channel morphology and control export of
sediment and organic matter. Measurement of
slope can range in spatial scale, generally
losing resolution with increasing spatial
extent. Slope can be determined either at the
streambed or water surface. The following
procedure describes the estimation of slope for
the streambed along the study reach thalweg.
Slope is measured once for a given reach
during the study because it is unlikely to
change significantly over short time periods
(e.g., 1—2 years). This procedure will require
1—2 field crew members to perform depending
on the method chosen.
Unit length of channel ("Run")
Streambed surface
Figure 3-21 Longitudinal section of channel.
Procedure
Delineate the 30-m study reach so that a
measuring tape is marking locations along the
thalweg. Slope is measured at 10-m intervals
(at 0-10, 10-20, and 20-30 m marks) along the
study reach (Figure 3-22).
51
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20m
Figure 3-22 Plan view of study reach showing measurement locations (vertical black tick
marks) for channel slope. Flow is from right to left and the dotted line represents the
thalweg.
3.7.1 Measuring slope with a clinometer
and stadia rod
The procedure requires one person holding
the stadia rod and another person ("viewer")
viewing the stadia rod through the
clinometer. While standing on level ground,
mark the stadia rod at the viewer's eye level
with brightly colored flagging. This will be
the target for the viewer when measuring
slope. Make sure the viewer's posture is the
same (stand-up straight and flat footed)
when marking the stadia rod and when
taking measurements. Alternatively, the
clinometer may be positioned at a set height
(top of meter stick or hiking pole), rather
than held by an observer. The target height
on stadia rod would then be flagged at the
same set height.
The viewer stands at the 0-m mark in the
thalweg, whereas the person holding the
stadia rod stands at the 10-m mark in the
thalweg (Figure 3-23). The stadia rod
should be held perpendicular to the
streambed at the 10-m mark. To standardize
for differences in thalweg depth the viewer
and the stadia rod should be positioned at
the same water depth (e.g., level with
surface of water; see Kaufmann and Robison
1998). However, this difference is often
negligible when all three slope
measurements along the reach are averaged.
The viewer looks through the clinometer
with one eye and at the stadia rod with the
other
Figure 3-23 Crew members measuring
slope of intermittent stream.
eye. Allow the images to appear to be
superimposed on each other and position the
horizontal center line of the clinometer level
with the marking on the stadia rod (Figure 3-
24). Avoid covering side window of
clinometer with your hand while viewing.
This window allows light through, enabling
you to read values. There are two scales
along the measurement wheel: degrees and
percentages. The percentage scale is on the
right side of the measurement wheel of most
52
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clinometers. Tip your head up while
viewing through the clinometer to see unit
markings (e.g., %) and determine which side
is the percentage scale. Slope measurements
are recorded in percentages (to the nearest
0.5%) on the datasheet (Figure 3-25).
Repeat the procedure for 10-20 and 20-30 m
intervals along the reach thalweg.
Stadia rod with
viewer's
eyelevel
Measurement wheel
Figure 3-24 Superimposed views through
clinometer and at stadia rod. Example
shows percent scale on right side and
degrees scale on left side of measurement
wheel.
CHANNEL SLOPE (%)
(for three 10 in sections of study reach)
4.5
10
Figure 3-25 Portion of page 1 of field
forms showing cells for percent slope
values.
Conversion between percent and degrees
can be done using:
degree slope = tan"1 (percent slope /100)
percent slope = (tan (degree slope)) X 100
Modifications to the procedure can
accommodate the use of alternatives to a
clinometer for measuring slope (e.g., Abney
level, theodolite, total station; see Gordon et
al. 1992). This procedure can be modified
to
measure water slope by simply accounting
for differences in water depth (or ensuring
equal water depth) at the stadia rod and
where the viewer is standing.
3.7.2 Hydrostatic (manometer)
measurement of slope
Position stakes at the 0 and 10-m marks
along the thalweg. Fill vinyl tube with
water and ensure no air bubbles are trapped.
Attach the ends of the vinyl tubing to the
stakes and position the tubing along the
thalweg of the streambed (Figure 3-26).
Allow water level within the vinyl tubing to
equilibrate. Using the meter stick, measure
(in meters) the distance between the
streambed and the water level (bottom of
meniscus) within the vinyl tubing at both
ends. Streambed slope (%) is ((h2 - hi) / L)
X 100, where L = 10 m. Slope
measurements are recorded in percentages
on the datasheet (Figure 3-25). Repeat the
procedure for 10-20 and 20-30-m intervals
along the reach thalweg. An alternative to
using rebar and clamps to hold the
manometer in place is to have two people
hold the ends of the manometer against
meter sticks while taking measurements of
hi and h2.
An advantage of this procedure is that it can
be done without a clear line of view along
the reach and it is more accurate than the
clinometer method. A disadvantage is that
water must be available for the manometer.
Water slope can be determined by
measuring the distance between the water
level within the tube and the water surface
(rather than the streambed surface) at both
ends.
53
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Stake
Streambed surface
Figure 3-26 Longitudinal section of channel showing position of manometer and points of
measurement to calculate slope (redrawn fro Gordan et al. 1992). Blue arrow shows
direction of flow. L = horizontal length, hi = height at the upstream end and hi = height at
downstream end.
References
Gordon, N.D., T.A. McMahon, and B.L.
Finlayson. 1992. Stream Hydrology: An
Introduction for Ecologists. John Wiley &
Sons, Chichester, United Kingdom.
Kaufmann, P. R. and E. G. Robison. 1998.
Physical habitat characterization. Pages 77-
118 in J.M. Lazorchak, DJ. Klemm, and
D.V. Peck (editors). Environmental
Monitoring and Assessment Program -
Surface Waters: Field Operations and
Methods for Measuring the Ecological
Condition of Wadeable Streams.
EPA/620/R-94/004F. U.S. Environmental
Protection Agency, Washington, D.C., USA.
Equipment and supplies
Measuring tape (50 m)
Field forms
AorB
A. Stadia rod and clinometer - See Procedure
3.7.1
B. Manometer (clear vinyl tubing, >10 m in
length and -lOmm inner diameter), 2 survey
stakes or pieces of rebar, hose clamps, and
meter stick - See Procedure 3.7.2
3.8 Measuring water depth
General
This subsection provides instructions for
measuring water depth (including maximum)
for reaches of headwater streams. Along with
wetted width (next section), water depth is a
critical measure of the extent of wetted habitat
available and a measure of water persistence
or susceptibility to terrestrial predators. Water
depth is therefore important in governing the
distribution of biota in headwater streams
(e.g., Harvey and Stewart 1991, Taylor 1997).
Because water depth can vary considerably
over time, this procedure should be carried out
during each sampling visit.
Procedure
3.8.1 Longitudinal thalweg measurements
A total of 31 measurements of water depth are
taken along each study reach (Figure 3-27).
Water depth is measured at the center of the
thalweg (illustrated as dotted line in Figure 3-
27) at meter intervals (i.e., 0, 1, 2...30 m).
54
-------
The meter stick is positioned with zero-end
down, side(s) with units facing perpendicular
to the direction of flow and the stick held
perpendicular to the water level (Figure 3-27).
Water depth measurements are recorded to the
nearest 0.5 cm on the field form (Figure 3-28).
30m
Water Depth (c
15 m = 39 cm
Figure 3-27 Overhead view of study reach showing locations for water depth measurement
(vertical black tick marks) along the reach thalweg (dotted line). Water is flowing from
right to left. (A.) overhead view of study reach (B.) channel cross-section, and (C.) lateral
close-up of depth.
55
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* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2BF Page 2 of 4
Meter
#
0
i
2
3
4
5
6
7
8
9
10
11
12
13
14
** •
15
^- ^
Modal
Sediment
Particle
Size
(mm)*
s
Water
Depth
(cm)*
22
18
5.5
1
2.5
3
50
30
35.5
0
0
3
34
12
8
. 39
Habitat
Type
(E/D)
Notes
(e.g., LWD,
Leafpack)
Velocity
(m/s)*
Wetted
Width
(m)
BE Width
(m)
BE
Depth
(m)
ft
EPA
width
(m)§
Figure 3-28 Appropriate location for recording longitudinal water depth measurements on
page 2 of the field forms.
The water level on the meter stick is usually
not perpendicular to the unit markings where
the water velocity is fast (Figure. 3-29).
Measurements should be taken at the middle
of the meter stick, rather than at the upstream
or downstream-facing edges. Where there is
no surface water present, zero water depth is
recorded. Where there is surface water
present, but it is less than 0.5 cm deep, "< 0.5
cm" should be recorded.
56
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Figure 3-29 Schematic showing appropriate reading of water depth where water surface is
turbulent.
3.8.2 Maximum water depth in study reach
A single measurement is recorded for the
greatest water depth within the study reach.
This measurement is not restricted to the 31
(1-m interval) thalweg measurements.
Maximum water depth is recorded to the
nearest 0.5 cm on the field form (Figure 3-30).
54
DEPTH TO BEDROCK /
GROUNDWATER (m)
(3 measures in depositional habitat)
SINUOSITY
(number of bends)
Figure 3-30 Appropriate location for recording maximum pool depth measurement on
page 1 of the field forms.
References
Harvey, B.C. and A.J. Stewart. 1991. Fish
size and habitat depth relationships in
headwater streams. Oecologia 87:336-342.
Taylor, C. M. 1997. Fish species richness
and incidence patterns in isolated and
connected stream pools: effects of pool
volume and spatial position. Oecologia
110:560-566.
Equipment and supplies
Meter stick (with at least 0.5cm increments)
Measuring tape (50m)
Field forms
57
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3.9 Measuring wetted width
General
This subsection provides instructions for
measuring wetted width in headwater streams.
Wetted width (or top width) is the stream
width at the surface water level (Figure 3-31)
and is perpendicular to the channel direction.
This measure (and water depth) describes the
extent of surface water habitat available
within a study reach. Because wetted width
can vary considerably over time, this
procedure should be carried out during each
sampling period.
Wetted Width
Figure 3-31 Channel cross-section illustrating wetted width.
Procedure thalweg. Wetted width is measured at 5-m
Delineate the 30-m study reach so that a
measuring tape is marking locations along the
intervals (at 0, 5, 10, 15, 20, 25 and 30-m
marks) along the study reach (Figure 3-32).
Figure 3-32 Overhead view of study reach showing measurement locations (vertical black
tick marks for wetted width. Flow is from right to left and the dotted line represents the
thalweg.
The meter stick can be used to measure wetted
widths < 1 m, whereas wider channels may
require using a measuring tape (and a survey
stake if done by one individual). At each
location place the zero-end of the meter stick
or tape at the water's edge on one side of the
channel, position the measuring device
perpendicular to channel direction, and
determine the distance to the water's edge on
the other side of the channel. Record the
distance to the nearest 0.01m in the
appropriate cell on the field form (Figure 3-
33).
58
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* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2BF Page 2 of 4
Meter
#
0
Modal
Sediment
Particle
Size
(mm) *
Water
Depth
(cm)*
Habitat
Type
(E/D)
Notes
(e.g., LWD,
Leafpack)
Velocity
(m/s)*
^^"^^^
Wetted
Width
(m)
0.76
BE
Width
(m)
BE
Depth
(m)
ft
EPA
width
(m)§
Figure 3-33 Appropriate location for recording wetted width measurement on page 2 of
the field forms.
If there is no surface water at a measurement
location, indicate on the field form that the
wetted width is 0 m. Where there are
individual boulders or cobbles interrupting the
surface water along the wetted width or there
is visible interstitial flow (see Section 3.1),
include the emergent particles in the
measurement (Figure 3-34 A). If there are
isolated pools along the channel edge (no
surface connection to main channel) or the
channel is braided (where there are vegetated
islands or patches of emergent substrate) do
not include width of isolated side-pools and
islands in the wetted width measurement
(Figure.3-34B, C).
Wetted Width
isolated
side-pool
Figure 3-34 Channel cross-sections showing wetted width measurements where there is
emergent cobble (A.), island (B.), and side-pool (C.).
Equipment and supplies
2 Measuring tapes (50m)
Meter stick
Survey stake (optional)
Field forms
59
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3.10 Measuring basic channel
geomorphology
General
This section provides instructions for rapidly
measuring basic channel form of headwater
streams. Specifically, this section provides
directions for measuring three channel
parameters: bankfull width, bankfull depth,
and flood-prone area width. The stream
channel is composed of the banks and the
streambed. The banks often have steeper
gradient (in cross-section) and are often
composed of finer sediments than the
streambed (Figure 3-35). Bankfull discharge
occurs when there is sufficient flow to fill the
entire channel. This level is called bankfull
stage and typically occurs once every 1-2
years. Bankfull width is the horizontal
distance between the banks (perpendicular to
flow) at bankfull stage. Bankfull depth is the
vertical distance between the streambed and
the bankfull stage height at the thalweg.
Flood-prone area width is the distance across
the channel at a vertical level equaling 2X the
bankfull depth. Entrenchment ratio is the ratio
of the flood-prone area width to the bankfull
width and is used to describe the degree of
channel incision or "down-cutting" (Rosgen
1994, 1996). Channel dimensions vary with
flow, the sediment being transported, and the
material composition of the bed and banks.
Channel geomorphology influences many
structural and functional aspects in streams,
including streambed substrates, organic matter
retention, and biotic response to floods. The
scouring forces of floods are dissipated on the
banks to greater extent in wide, shallow
channels, whereas these forces are focused on
the streambed in constrained or incised
Figure 3-35 Headwater stream channel showing the location of the streambed and the
banks (white arrows).
60
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channels (Carling 1983). Geomorphology
also governs the distribution of water as
streams dry. Wetted widths will contract
faster in wide, shallow channels than in
incised channels. Wide, shallow channels
may be more prone to surface water drying
than incised channels because the summer
groundwater table is more likely to be above
the streambed (Stanley et al. 1997).
However, where drying is severe, incised
channels offer less interstitial refugia
because the substrate layer above underlying
bedrock may be thin. Habitat simplification
reduces the biotic diversity directly, but also
affects diversity indirectly through loss of
refugia (Lake 2003).
Channel geomorphology is measured once
for a given reach during the study because
they are unlikely to change significantly
over short time periods (e.g., 1-2 years).
However, floods can significantly reshape
channel geometry over short periods of time
and should be taken into account when
investigators need fine temporal resolution
data. The following procedure will require
2-3 field crew members, depending upon the
channel width.
3.10.1 Bankfull width (BF width)
Field determination of bankfull stage is
particularly difficult for small channels
where the floodplain may not be well-
developed or may be absent. Useful
indicators of bankfull stage include breaks in
sediment particle size and bank vegetation.
Swift and Ledford (1994) identifies the
following characteristics for estimating
bankfull stage in small southern
Appalachian streams:
1. Topographic break from vertical bank to
floodplain
2. Topographic break from steep to gentle
slope
3. Top of point bar
4. Change in vegetation from temporary to
permanent
5. Upper elevation of fine debris deposition
6. Rocks and/or roots exposed in banks
7. Change in size distribution of deposits
8. Change in texture of fines lodged
between rocks
20m
15m
Figure 3-36 Plan view of study reach showing 5-m intervals. Direction of arrows shows
direction of flow, and the dotted line represents the thalweg.
Procedure
Delineate the 30-m study reach so that a
measuring tape is marking locations along the
thalweg. Bankfull width and depth are
measured at 5-m intervals (at 0, 5, 10, 15, 20,
25 and 30-m marks) along the study reach,
whereas flood-prone area width is measured at
15-m intervals (at 0, 15, and 30-m marks;
Figure 3-36). Measurements should be taken
at the next meter mark (upstream or
downstream) along the study reach where
obstacles (e.g., large woody debris) or certain
channel features (e.g., meanders, knickpoints)
are present at original measurement locations
61
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(0, 5, 10, 15, 20, 25, and 30 m). Note on the
field form where measurements were taken.
It is useful to look upstream and downstream
along both banks of measurement location to
identify appropriate bankfull stage. When a
consensus among crew members is made
about the appropriate bankfull stage, the end
of a measuring tape is staked at bankfull stage.
The tape is pulled across the channel
(perpendicular to direction of flow) to the
other bank to determine bankfull width
(Figure 3-37). A second crew member,
standing downstream, provides instruction for
adjusting the tape position so that it is
horizontally level at the bankfull stage. This
can be done more accurately if a laser level is
used to adjust the tape position. Ensure that
the tape is taut and record the distance (to the
nearest 0.01 m) in the appropriate cell on the
second page of the field form (Figure 3-38).
3.10.2 Bankfull depth (BF depth)
While the tape is still positioned for measuring
bankfull width, a crew member uses the meter
stick to measure bankfull depth (Figure 3-37).
The meter stick (zero-end down) is positioned
perpendicular to the tape measuring bankfull
width at the center of the thalweg. Record the
distance (to the nearest 0.01 m) between the
.
Figure 3-37 Photograph shows measurement of bankfull (BF) width and bankfull depth.
62
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* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2BF Page 2 of 4
Meter
#
0
Modal
Sediment
Particle
Size
(mm) *
Water
Depth
(cm)*
Habita
t Type
(E/D)
Notes
(e.g., LWD,
Leafpack)
Velocity
(m/s)*
Wetted
Width
(m)
BE BF EPA
Width Depth widtn
(m) (m) (m) §
ft
1.43 0.2 3.02
Figure 3-38 Appropriate location for recording bankfull (BF) width (red), bankfull depth
(blue), and flood prone area (FPA) width (black) measurements on page 2 of the field
forms.
streambed and the tape in the appropriate cell
on the second page of the field form (Figure 3-
38).
3.10.3 Flood-prone area (FPA) width and
entrenchment ratio
At the 0, 15 and 30-m locations the crew
members then locate 2X the bankfull depth
and raise the tape to that level for measuring
the width of the flood-prone area (FPA width,
Figure 3-39). The crew member with the tape
adjusts ends of the tape so that it is
horizontally level and extended tautly across
the channel to touch soil at both ends. Where
the distance of the flood-prone area width is
>2.2X the bankfull width, record ">2.2X
BFW", otherwise record to the nearest 0.01 m
Figure 3-39 Photograph illustrating flood-prone area (FPA) width.
in the appropriate cell (Figure 3-38). The
significance of the 2.2X bankfull width is
based on Rosgen (1994, 1996) channel
classification, where entrenchment ratios >2.2
63
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(FPA > 2.2X BFW) are classified as slightly
entrenched (stream types C, D, or E). As was
done when measuring the bankfull width, a
crewmember provides instruction for
adjusting the tape position so that it is
horizontally level at the 2X bankfull depth.
This can be done more accurately if a laser
level is used to adjust the tape position.
References
Carling, P. A. 1983. Threshold of coarse
sediment transport in broad and narrow
natural streams. Earth Surface Processes
and Landforms 8:1-18.
Lake, P. S. 2003. Ecological effects of
perturbation by drought in flowing waters.
Freshwater Biology 48:1161-1172.
Rosgen, D. 1994. A classification of natural
rivers. Catena 22:169-199.
Rosgen, D. 1996. Applied River Morphology,
2nd edition. Wildland Hydrology, Pagosa
Springs, CO.
Stanley, E. H., S. G. Fisher, and N. B. Grimm.
1997. Ecosystem expansion and contraction
in streams BioScience 47:427-435.
Swift, L. W. Jr. and E. S. Ledford. 1994.
Where is bankfull in small southern
Appalachian stream channels? Pages 306-
311 in Riparian ecosystems in the humid
U.S.: Functions, Values and Management.
National Association of Conservation
Districts, Washington, D.C.
Equipment and supplies
2 Measuring tapes (50m)
Meter stick
Field forms
Survey stakes
Laser level (optional)
3.11 Measuring water velocity
General
The following subsection provides methods
for measuring water velocity in headwater
streams. Water velocity is the rate of water
moving through a point and represents one
aspect of stream flow. Hydraulics is among
the more complex and dynamic characteristics
of the stream environment (Statzner et al.
1988, Vogel 1994). For example, the
relevancy of a velocity is dependent on
organism size. Under the same velocity,
smaller organisms may experience the near-
bed velocity as laminar syrup, whereas larger
organisms would experience a turbulent
maelstrom. Although water velocity is just
one aspect of stream hydraulics, it provides
ecologically-relevant information. The
following methods will offer coarse estimates
that are useful in for making relative
comparisons. For fine-scale and less-invasive
measurements, alternative methods such as
acoustic Doppler velocimeter (ADV,
Bouckaert and Davis 1998, Finelli et al. 1999)
and thermistor probes (LaBarbera and Vogel
1976, Dodds and Biggs 2002) are more
suitable. As already discussed in Subsection
3.6, water velocity is useful for designating
habitat units and can directly (e.g., food
availability, dispersal) and indirectly (e.g.,
refuge from predators) affect the distribution
of organisms (Hart and Finelli 1999). Mean
water velocity for a stream reach may not
necessarily decline as streams first begin to
dry, but it will drop dramatically when
streambed materials such as cobbles and
boulders become emergent and flow becomes
mostly interstitial. Because water velocity can
vary considerably over time, measurements
should be taken during each sampling visit.
Below we detail four simple procedures for
measuring water velocity along a stream
reach; additional procedures are discussed by
John (1978), Newbury (1984), and
Ciborowski (1991).
64
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Procedure
Delineate the 30-m study reach so that the
measuring tape is marking locations along the
thalweg. Point measurements of water
velocity (Procedures 3.11.1,3.11.2 and
3.11.3) at the streambed are taken at 5-m
intervals (at 0, 5, 10, 15, 20, 25 and 30-m
marks) along the study reach thalweg (Figure
3-40). Below are four procedures that can be
used. In most cases (and when available) the
velocity meter procedure is preferred;
30m
20m
15m
Figure 3-40 Plan view of study reach showing measurement locations (vertical black tick
marks) for current velocity measurements. Flow is from right to left and the dotted line
represents the thalweg.
however under some circumstances the other
three procedures may be more suitable.
3.11.1 Velocity meter procedure
Before arriving at the field site read the
instruction manual for the velocity meter (e.g.,
electromagnetic, propeller). Attach the
wading rod to the velocity meter probe.
Check to see that the meter is functioning
properly and is calibrated. Set the selector
switch to m/sec and the time constant switch
to the lowest setting that gives stable readings.
Stand downstream and to the side of each of
the measurement locations when taking
velocity readings. Hold the rod perpendicular
to the water surface with the front of the
velocity probe facing upstream, perpendicular
to the channel cross-section (Figure 3-41). Set
the bottom of probe -0.5 cm off the
streambed and take flow reading. Write the
water velocity in the appropriate cell on the
second page of the field forms (Figure 3-42).
If no surface water is found at a measurement
location, indicate on the field form that the
water velocity is 0. If there is flowing surface
water at a location but it is too shallow to
measure with a velocity meter, then indicate
that the water velocity is ">0".
65
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Figure 3-41 Longitudinal section across the channel thalweg showing orientation of the
velocity probe for measurements.
* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2BF Page 2 of 4
Meter
#
0
Modal
Sediment
Particle
Size
(mm)*
Water
Depth
(cm)*
Habitat
Type
(E/D)
Notes
(e.g., LWD,
Leafpack)
Velocity
(m/s)*
0.02
Wetted
Width
(m)
BE
Width
(m)
BE
Depth
(m)
EPA
width
(m)§
Figure 3-42 Appropriate location for recording water velocity on page 2 of the field forms.
3.11.2 Velocity-area procedure using a bag
meter (Gessner meter)
A simple alternative to electromagnetic or
propeller meters is the bag meter or Gessner
meter (Gessner 1950). To assemble the bag
meter: tape a plastic bag (e.g., small plastic
grocery or bread bag) over the larger opening
of a small plastic funnel with duct tape. Make
sure that it is completely sealed and there are
no holes in the plastic bag. Then tape a
cylinder (e.g., plastic cup with bottom cut out,
PVC pipe) that has a diameter slightly larger
than the large opening of the funnel), to the
outside of the large funnel opening and over
the plastic bag (Figure 3-43). Calculate the
area of the small funnel opening (i.e., A = 71
66
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Funnel
Bag
Cylinder
Figure 3-43 Bag meter used to measure water velocity.
Stand downstream and to the side of each of
the measurement locations when measuring
velocity. Before taking a measurement, empty
and deflate the bag as much as possible.
While holding the bag meter by the cylinder in
one hand, use your other to cover the small
funnel opening. Submerge and hold the bag
meter near (will depend on diameter of large
funnel opening) and parallel to the stream bed,
so that the small opening is facing into the
current. Simultaneously note the second hand
position on your wristwatch (alternatively
signal "start" to another crew member with a
stopwatch) and uncover the small funnel
opening. Let the bag fill with water for 10
seconds (or shorter time in very fast current)
and recover the funnel opening. Carefully
pour the water from the bag into the calibrated
container. Determine the volume to the
nearest 0.005 liter. In the cells for water
velocity on the field form (Figure 3-42) write
the volume and fill time (e.g., 0.25 L /10 sec).
Indicate on page 3 of the field form that the
bag meter was used to measure discharge and
the area of the small funnel opening.
After returning from the field, the water
velocities can be calculated by first converting
the volumes from liters to m3 (i.e., divide by
1000). The volume is divided by the filling
time (e.g., 10 sec) and then the resulting value
is divided by the area of the small funnel
opening (in meters). Repeat this for all
measurement locations. If there is no surface
water at a measurement location, indicate on
the field form that the water velocity is 0. If
there is flowing surface water at a location but
it is too shallow for this method indicate that
the water velocity is >0.
3.11.3 Neutrally-buoyant obj ect procedure
This procedure can be used when a velocity
meter is not available or if flow is too shallow
for accurate meter readings. Indicate on page
3 of the field form that this procedure was
used. Designate the upstream and
downstream boundaries of 2-m segments that
are centered on each of the measurement
locations (1 m upstream and downstream of
the 0, 5, 10,...30-m locations, Figure 3-44).
While standing downstream of the release
point and outside the thalweg, hold the
neutrally-buoyant object (see Equipment and
supplies for examples; consistently use the
same object across all measurements) in the
thalweg (at 0.4X the water depth). In unison,
gently release the neutrally-buoyant object and
start the stop watch. Note the time required
for the object to travel the 2-m segment. If the
object becomes stuck or drags along the
bottom repeat the release and/or slide the
segment position upstream or downstream to
avoid areas where the object sticks or drags.
In fast segments 2 people may be required to
67
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accurately measure segment travel time. One
person at the upstream boundary
simultaneously releases the object and signals
"start". This indicates to the second person
who is standing at the downstream boundary
to start the stopwatch. The second person then
stops the watch when the object crosses the
downstream boundary. Divide the segment
length by the travel time and write this in the
appropriate cell on the field form. If there is
no surface water at a measurement location,
indicate on the field form that the water
velocity is 0. If there is flowing surface water
at a location but it is too shallow for this
method indicate that the water velocity is >0.
30m
15m
Figure 3-44 Overview of study reach showing measurement locations (black tick marks
crossing the thalweg, shown as dotted line), upstream (dashed blue lines) and downstream
segment boundaries (solid red lines) for the neutrally-buoyant procedure to measure water
velocity.
3.11.4 Fluorescent dye procedure
This procedure can be used when a velocity
meter is not available or if flow is too shallow
for accurate meter readings. Indicate on page
3 of the field form that this procedure was
used. This procedure provides only a general
measure of water velocity for the entire reach,
in contrast to the methods described above
which provide estimates for average and
variation of water velocity. Pour ~ 1 ml
fluoroscene dye (or rhodamine WT) into a 1 L
plastic bottle and add 500 ml of stream water.
Cap and shake bottle until dye is thoroughly
dissolved. In fast-flowing reaches 2 people
may be required for this method, one person
with the dye at the 30-m location (upstream
boundary of study reach) and the other person
with a stopwatch at the 0-m location
(downstream boundary of study reach).
Before starting, make sure that other field
personnel are outside of the study reach. The
person at the upstream boundary will
simultaneously release the dye (gently pouring
bottle contents from ~ 5 cm above the water
level) into the thalweg at the 30-m location
and signal "Start". This indicates to the
person at the downstream boundary to start the
stopwatch. The downstream person records
the time when the "leading" and "trailing"
edges of the dye plume cross the downstream
boundary (Figure 3-45). The trailing edge is
identified as the last visible portion of the
plume in the thalweg. Ignore any dye that
may have gotten caught in backwater pockets.
On the field form write the distance of the dye
release (should be 30 m if entire study reach is
flowing) and travel times for leading and
trailing edges in seconds.
68
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Figure 3-45 Overhead view of study reach showing leading and trailing edges of
fluoroscene plume.
References
Bouckaert, F. W. and J. Davis. 1998.
Microflow regimes and the distribution of
macroinvertebrates around stream
boulders. Freshwater Biology 40:77-86.
Ciborowski, J. J. H. 1991. Head tube: a
simple device for estimating velocity in
runnint water. Hydrobiologia 222:109-
114.
Dodds, W. K. and B. J. F. Biggs. 2002.
Water velocity attenuation by stream
periophyton and macrophytes in relation to
growth form and architecture. Journal of
the North American Benthological Society
21:2-15.
Finelli, C. M., D. D. Hart, and D. M.
Fonesca. 1999. Evaluating the spatial
resolution of an acoustic Doppler
velocimeter and the consequences for
measuring near-bed flows. Limnology and
Oceanography 44:1793-1801.
Gessner, F. 1950. Die okologische
Bedeutung der Stromungsgeschwindigkeit
flieBender Gewasser und ihre Messung auf
kleinstem Raum. Archivfur
Hydrobiologie 43:159-165.
Hart, D. D. and C. M. Finelli. 1999.
Physical-biological coupling in streams:
the pervasive effects of flow on benthic
organisms. Annual Review of Ecology and
Systematics 30:363-395.
John, P. H. 1978. Discharge measurement
in lower order streams. International
Revue der Gestamten Hydrobiologie
63:731-755.
LaBarbera, M. and S. Vogel. 1976. An
inexpensive thermistor flow meter for
aquatic biology. Limnology and
Oceanography 21:750-756.
Newbury, R. 1984. Hydrologic
determinants of aquatic insect habitats.
Pages 323-357 in V. H. Resh and D. M.
Rosenberg (editors). The ecology of
aquatic insects. Praeger, New York.
Statzner, B., J.A. Gore, and V.H. Resh.
1988. Hydraulic stream ecology: observed
patterns and potential applications.
Journal of the North American
Benthological Society 7:307-360.
69
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Vogel, S. 1994. Life in Moving Fluids, 2nd
edition. Princeton University Press,
Princeton, New Jersey.
Equipment and supplies
Measuring tape (50 m)
Field forms
A,B, CorD
A. Velocity meter (electromagnetic,
propeller, or cup) and spare batteries -
see Procedure 3.11.1
B. Stopwatch and bag meter - see
Procedure 3.11.2
C. Stopwatch and neutrally buoyant object
(e.g., piece of orange peel, film canister
partially filled with stream water, small
stick) - see Procedure 3.11.3
D. Stopwatch, 1L plastic bottle, fluoroscene
dye (1 ml per 500 ml streamwater) - see
Procedure 3.11.4
3.12 Measuring discharge
General
This subsection provides methods for
measuring discharge (Q) or flow rate of
water in headwater streams. Discharge (in
conjunction with stream size or drainage
area) is a quantitative measure for describing
the hydrologic condition. This measure of
flow is useful in following and describing
temporal patterns in water chemistry. When
conditions are allowable, discharge should
be measured during each sampling visit.
The methods described in this subsection are
modified from those described by John
(1978), Platts et al. (1983), Kilpatrick and
Cobb (1985), Gordon et al. (1992), Gore
(1996), and Kaufmann (1998). For long-
term studies continuous discharge
monitoring may be considered. The
simplest method is a staff gauge, where
discharge can be determined by monitoring
the stage (or water depth) at a permanent
location. Stage-discharge relationships
(rating curves) are plotted by measurements
of stage against discharge over a range of
flows (Gordon et al. 1992). Peak flow
between field visits can be determined from
crest gauges (Gordon et al. 1992, Harrelson
et al. 1994). A simple crest gauge consists
of stilling well, a meter stick, and ground
cork. The stilling well can be a length of
plastic pipe (3 to 4 cm diameter) with caps
on both ends. Holes are drilled in the
bottom cap so the water level within the
stilling well represents the stage. The top
cap of the well should be loose fitting or
vented. Finely ground cork and the meter
stick are placed in the well. After a peak
flow the cork will adhere to the meter stick
at the crest or peak stage. The gauge is then
easily reset by washing the cork off the
meter stick and back into the well. The
design and equipment for gauging stations
can vary from a simple staff gauges to more
permanent flumes and weirs. Gauging
station design and data storage are discussed
in John (1978), Herschy (1995), Clemmons
et al. (2001), and Bureau of Reclamation
(2001).
Procedure
Delineate the 30-m study reach so that a
measuring tape is positioned along the
thalweg.
3.12.1 Velocity-area procedure using a
velocity meter
Before arriving at the field site read the
instruction manual for the velocity meter.
Attach the wading rod to the velocity meter
probe. Check to see that the meter is
functioning properly and is calibrated. Set
the selector switch to m/sec and the time
constant switch to the lowest setting that
gives stable readings (unit setting may be
switched to ft/s under extremely low flow
conditions). The location for discharge
70
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20m
15m
Wetted Width = 1.01 m
A
Cell midpoints at 0.4X water
depth from the streambed
Figure 3-46 Plan view of study reach (top) showing discharge measurement cross-section
(red dashed line). Cross-section for discharge measurement (bottom) showing
measurement cells.
measurement is not restricted to the 30-m
study reach; however, the discharge at the
measurement location should be
representative of the discharge seen in the
study reach. Locate a channel cross-section
that has the following characteristics (or can
be modified to have these characteristics*):
1) channel immediately upstream and
downstream is straight (~ 3 m in both
directions of discharge transect), 2) free of
obstructions (e.g., woody debris,
macrophytes, emergent stones, braided
channel), 3) "U" shaped so that > 90% of the
cross-section has water depths sufficiently
deep for accurately measuring water
velocity with the velocity meter, and 4)
water velocity across the channel is
relatively uniform and > 90% of the cross-
section has water velocities >0.01 ms" .
Runs and glides are typically good habitat
units for measuring discharge.
At the measurement cross-section, stretch
the second measuring tape taut across the
channel so that it is perpendicular to flow
and > 5 cm above the stream surface (Figure
3-46). Determine the wetted width of the
channel to the nearest 0.01 m. Divide the
wetted width into 6 to 12 equally sized
intervals or cells. Cells should be > 5 cm
wide.
Write the wetted and cell widths in the
appropriate blanks on the field form (Figure
3-47). Water depth and water velocity are
measured midway across each cell or cell
midpoint (Figure 3-46).
* The channel can be modified (e.g., remove rocks,
obstructions) prior to taking any discharge
measurements. Once measurements have begun
however, do not modify the channel.
71
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Start measurements from one bank and move
across. Stand downstream and to the side of
each depth and velocity measurement. Use
the meter stick to measure water depth (to the
nearest 0.5 cm). Water velocity is then
measured at ~ 0.4X water depth from the
streambed for each cell. If this depth is too
shallow to submerge the velocity meter probe
or propeller, measure velocity closer to the
streambed. Write the water depth and its
associated water velocity measurement in the
cells on the field form (Figure 3-47).
Discharge is calculated by multiplying the cell
width * water depth * water velocity of each
cell then summing across all cells.
STREAM DISCHARGE Page 3 of 4
Wetted Width (
Depth(cm)
Velocity(m/s)
m): 1.01
4
0
Ofm3s ')= 0.02908
7
0.05
CELL WIDTH (
11
0.09
16.5
0.12
m) : 0.1
25
0.22
28
0.25
27
0.18
10.5
0.08
5.5
0
Discharge procedure: Velocity-area
Velocity procedure/meter model: Marsh-McBirnev Flowmate
Figure 3-47 Appropriate location for recording discharge and procedures used on page 3
of field forms. Example values shown in red.
3.12.2 Velocity-area procedure using a bag
meter
To assemble the bag meter: tape a plastic bag
(e.g., small plastic grocery or bread bag) over
the larger opening of a small plastic funnel
with duct tape. Make sure that it is
completely sealed and there are no holes in the
plastic bag. Then tape a cylinder (e.g., plastic
cup with bottom cut out, PVC pipe that has a
diameter slightly larger than the large opening
of the funnel), to the outside of the large
funnel opening and over the plastic bag
(Figure 3-48). Calculate the area of the small
funnel opening (i.e., A = 71 r2).
Funnel
Bag
Cylinder
Figure 3-48 Bag meter used to measure discharge.
72
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Select and delimit a measurement cross-
section as described in 3.12.1. Follow the
same procedures except use the bag meter to
measure water velocity at 0.4X the water
depth from the stream bed. Before taking a
measurement completely empty the bag of
water and deflate the bag of air as much as
possible. While holding the bag meter by the
cylinder in one hand use your other to cover
the small funnel opening. Submerge and hold
the bag meter at the appropriate measuring
depth, so that the small opening is facing into
the current and the bag meter is perpendicular
to the measurement cross-section.
Simultaneously note the second hand position
on your wristwatch (alternatively shout "start"
to another crew member with a stopwatch)
and uncover the small funnel opening. Let the
bag fill with water for 10 seconds (or shorter
time in very fast current) and recover the
funnel opening. Carefully pour the water
from the bag into the calibrated container.
Determine the volume to the nearest 0.005
liter. In the cells for water velocity on the
field form write the volume and fill time (e.g.,
0.25 L /10 sec). Indicate on the field form
that the bag meter was used to measure
discharge and the area of the small funnel
opening.
After returning from the field, the cell water
velocities can be calculated by first converting
the volumes from liters to m3 (i.e., divide by
1000). The volume is divided by the filling
time (e.g., 10 s) and then the resulting value is
divided by the area of the small funnel
opening (in meters). Repeat this for all cells
of the measurement cross-section and
determine discharge as instructed in 3.12.1.
3.12.3 Timed filling procedure
This method can be used where the channel is
small and there are one or more natural
spillways or plunges along the reach where the
entire stream flow can be captured (the
channel can be modified to ensure that all the
flow is funneled). Simultaneously start the
stopwatch and position the wide-mouth
container (i.e., bucket or basin) under the
spillway to collect the entire flow. Collect
water for 10-30 seconds, depending upon the
level of discharge. Transfer the water from
the wide-mouth container to a calibrated one
and determine the volume (to the nearest
0.005 liter). Alternatively, one may simply
record the time required to fill a bucket or
basin to a known volume (e.g., 2 L). Repeat
this procedure 3 times at given spillway.
Indicate that the timed filling procedure was
used to measure discharge and write the
volume and respective filling time for each
trial on the field form.
3.12.4 Dilution gauging procedures
These methods use dilution over time of
biologically inert substances introduced into a
stream reach. Commonly used substances
(tracers) included salt solutions (NaCl, KBr)
and dyes (e.g. fluorescene, rhodamine WT).
Tracers should be readily detectable at low
concentrations (low or no background
concentrations), and soluble in water at stream
conditions (Gordon et al. 1992). Depending
upon the tracer used, general (electrical
conductivity meter, fluorometer) or tracer-
specific probes can be used for in situ
measurements. Alternatively, samples can be
collected in bottles and returned to the
laboratory for analysis. An estimate of
discharge is needed to determine the initial
tracer concentration so that the measured
concentration is easily detectable (5 to 10
times background). The two general methods
for dilution gauging are the slug injection and
constant injection. The slug injection method
involves releasing a known volume and
concentration of a tracer as a single pulse.
Background measurement for the tracer
should be measured before beginning the
injection. The point of injection should be a
73
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zone with turbulent mixing. Tracer
concentration is measured at regular intervals
at a downstream station from the start of the
injection until concentrations reach
background levels. The measurement interval
will depend upon the level of discharge and
the size of the study reach. Discharge (Q) is
determined from using the area under the
concentration curve (Figure 3-49). The
following equation from Gordon et al.
(1992)is used:
Where c\ is the stabilized concentration, Qt is
the tracer injection rate (1 s"1), and the other
variables are the same as shown in the
previous equation.
Figure 3-50 Example of a concentration
curve from a continuous injection.
Discharge (m3s *) is the hatched area under
c
c\ -
Concentration
Where Fis the slug volume (in liters), ct is
the initial tracer concentration, c0 is the
background concentration in the stream
water, c is the concentration at time t.
Concentration
ti
Time
Figure 3-49 Example of a concentration
curve from a slug injection. Discharge
(mV1) is the hatched area under the curve.
The constant injection method also uses a
known concentration of the tracer, but the rate
of injection is constant over the duration of the
measurement rather than as a slug. Tracer
concentration will increase and then stabilize
at the downstream station (Figure 3-50).
Constant injection can be done using a
peristaltic pump or a Mariotte bottle (see
Webster and Ehman 1996). Discharge using
this method is calculated using the equation
from Gordon et al. (1992):
(c -c
l
Injection
stopped
K
Time
the curve.
= 1000-
Qt
Although these methods may be more
accurate and feasible during low flows than
previously described methods, insufficient
mixing and anastomosing flow through
reaches may also limit discharge measurement
using dilution gauging methods. Some
disadvantages of dilution methods compared
to other methods include need for prior
knowledge of approximate discharge level,
additional equipment bulk, and drift response
by biota (Wood and Dykes 2002).
References
Bureau of Reclamation. 2001. Water
Methods Manual, 3rd edition. U. S.
Department of Interior, Washington D.C.
Clemmens, A. 1, T. L. Wahl, M. G. Bos, and
J. A. Replogle. 2001. Water Measurement
with Flumes and Weirs. Water Resources
Publications, Highlands Ranch, Colorado.
Gordon, N. D., T. A. McMahon, and B. L.
Finlayson. 1992. Stream Hydrology: An
74
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Introduction for Ecologists. John Wiley &
Sons, Chichester, United Kingdom.
Gore, J. A. 1996. Discharge measurement
and streamflow analysis. Pages 53-74 in:
F.R. Hauer and G. A. Lamberti (editors).
Methods in stream ecology. Academic Press,
San Diego, California.
Harrelson, C. C., C. L. Rawlins, J. P.
Potyondy. 1994. Stream channel reference
sites: an illustrated guide to field technique.
General Technical Report RM-245. Fort
Collins, CO: U.S. Department of
Agriculture, Forest Service, Rocky
Mountain Forest and Range Experiment
Station.
Herschy, R. W. 1998. Streamflow
Measurement, 2nd edition. Kluwer
Academic, New York.
John, P. H. 1978. Discharge measurement in
lower order streams. International Revue der
Gestamten Hydrobiologie 63:731-755.
Kaufmann, P. R. 1998. Stream discharge.
Pages 67-76 in: J.M. Lazorchak, D.J.
Klemm, and D.V. Peck (editors).
Environmental Monitoring and Assessment
Program - Surface Waters: Field
Operations and Methods for Measuring the
Ecological Condition of Wadeable Streams.
EPA/620/R-94/004F. U.S. Environmental
Protection Agency, Washington, D.C.
Kilpatrick, F. A. and E. D. Cobb. 1985.
Measurement of Discharge Using Tracers.
Techniques of Water-Resources
Investigations 3-A16, U.S. Geological
Survey, Washington, D.C.
Platts, W. S., W. F. Megahan, and G. W.
Minshall. 1983. Methods for Evaluating
Stream, Riparian, andBiotic Conditions.
USDA Forest Service General Technical
Report INT-183.
Webster, J.R. and T. P. Ehrman. 1996. Solute
dynamics. Pages 145-160 in: F.R. Hauer and
G. A. Lamberti (editors). Methods in Stream
Ecology. Academic Press, San Diego,
California.
Wood, P. J. and A. P. Dykes. 2002. The use
of salt dilution gauging techniques:
ecological considerations and insights.
Water Research 36:3054-3062.
Equipment and supplies
Measuring tape (50 m)
Field forms
A, B, C, or D
A. Second measuring tape, survey stakes,
meter stick, velocity meter
(electromagnetic, propeller, or cup) and
spare batteries - see Procedure 3.12.1
B. Second measuring tape, meter stick,
wristwatch with second hand, bagmeter
(small funnel taped to plastic bag enclosed
in plastic pipe), and calibrated container
(e.g., volumetric cylinder) - see
Procedure 3.12.2
C. Stopwatch, wide-mouth container (e.g.,
bucket, wash basin), and calibrated
container (e.g., volumetric cylinder) - see
Procedure 3.12.3
D. Stopwatch, tracer substance (stock
solution), calibrated pipette and tips,
volumetric cylinder, mixing container,
tracer probe or fluorometer, peristaltic
pump or Mariotte bottle - see Procedure
3.12.4
3.13 Measuring depth to bedrock and
groundwater table
General
75
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This section provides instructions for
measuring depth to underlying bedrock and
groundwater table in headwater streams. The
hyporheic zone is the interface between the
surface stream and the underlying
groundwater (Boulton et al. 1998, Jones and
Mulholland 2000). The importance of the
hyporheic zone to the structure and function of
streams depends upon the permeability and
discharge through the hyporheic zone to the
overlying surface water (Brunke and Gonser
1997). Because the subsurface environment
(e.g., temperature, flow) is relatively more
stable than the overlying streambed surface,
the hyporheic zone may serve as a refuge for
stream organisms from disturbances such as
floods and drying (e.g., Clinton et al. 1996,
Dole-Olivier et al. 1997). This rapid method
provides an estimate of the extent and
hydrologic status of the hyporheic zone, and
therefore the potential for it to serve as refuge.
Depth to groundwater table can vary with
intra- and interannual differences in catchment
precipitation and evapotranspiration, and it is
important to measure whenever surface water
is absent. Depth to bedrock is unlikely to
change significantly over short time periods
(e.g., 1-2 years), and therefore only needs to
be measured once during the study period.
Because these measurements use the same
procedure (e.g., sounding rod, Valett 1993),
we recommend taking both measurements
during drier periods (when more than one
sampling visit is planned).
The development of ground-penetrating radar
(GPR) offers an alternative, non-invasive
method to describe subsurface features of
streambeds, including the depth to bedrock
and groundwater (Naegeli et al. 1996,
Huggenberger et al. 1998). However, the
utility of GPR can be limited where interfaces
are not clearly defined (e.g., saturated fine
sediments) or below dense layers, such as
clays (Poole et al. 1997). The cost and bulk of
equipment are other considerations that may
limit the application of GPR in large scale
assessments of headwater streams.
Procedure
Delineate the 30-m study reach so that a
measuring tape is marking locations along the
thalweg. Locate 3 depositional habitat units
near the 0, 15, and 30-m marks of the study
reach. Depth measures are taken in the
thalweg at these 3 locations.
3.13.1 Depth to bedrock
Hammer the sounding rod or "T"-bar
vertically into the stream bed at intervals of 5-
10 cm with the hand sledge (Figure 3-51).
Wiggle the upper end of the sounding rod in
circular motion by hand (Figure 3-52). This
will prevent the rod from becoming stuck
within the stream bed. Continue tapping the
rod until it strikes bedrock (or large boulder).
This will be evident from the "pinging" sound
the rod makes when hammered (and resistance
to further downward movement). Some
stream beds have cobble deposition that may
impede the rod's downward progress. You
can penetrate through cobble layers by
rotating the rod tip in a circular motion while
continuing to hammer (Figure 3-52). This
process will often allow the rod to pass
through interstitial spaces between the
cobbles. If not, simply shift the rod location
slightly and repeat the process. When you
have struck bedrock, use your forefinger and
thumb (or cable tie) to mark the point on the
rod where it is even with the stream bed
surface. Pull the rod out of the stream bed and
measure the distance with the meter stick (to
the nearest 1 cm) between the lower end of the
rod and your finger. Write this measurement
in the appropriate cell on the field form
(Figure 3-53). If the depth to bedrock appears
to exceed the length of the sounding rod (> 85
cm for 91 cm sounding rod) then indicate
76
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">85 cm" on the field form. Where the stream
bed surface is bedrock then indicate "0 cm" on
the field form.
Figure 3-51 Using sounding rod and hand sledge hammer to estimate depth to bedrock
and the groundwater table.
3.13.2 Depth to groundwater table
Where the stream contains surface water the
depth to the groundwater table will equal the
water depth at the measurement location.
Indicate this on the form by writing "+" and
the water depth. Where the stream bed is dry
begin by following the same procedure used to
measure depth to bedrock. After the
groundwater table is reached, water seeping
into the hole will create resistance on the rod.
Moving the top of the rod in a circular motion
or gently lifting the rod a few centimeters will
help you determine if you have entered the
groundwater table. If the rod has entered the
water table, you may either hear a "slurping"
sound or feel suction resistance when the rod
is lifted. Before fully removing the sounding
rod from the streambed, mark the point (with a
finger or cable tie) on the sounding rod where
it is even with the stream bed surface.
77
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Figure 3-52 Cross-section of a dry channel illustrating depth to underlying bedrock (A)
and depth to the groundwater table (B).
Immediately after removing the sounding rod
from the stream bed identify the highest point
along the rod where there is water (wet
enough to drip). Measure the distance
between stream bed level and the highest
wetted point on the rod with the meter stick
(to the nearest 1 cm). Write this measurement
in the appropriate cell on the field form
(Figure 3-53). Indicate that this represents a
measurement below the stream bed surface by
writing a "-" before the distance. If the depth
to the groundwater table appears to exceed the
length of sounding rod (>85 cm for 91 cm
sounding rod) then indicate ">-85 cm" on the
field form.
MAX. POOL DEPTH (cm)
DEPTH TO BEDROCK /
GROUNDWATER (cm)
(3 measures in depositional habitat)
SINUOSITY
(number of bends)
Figure 3-53 Appropriate location for recording depth to bedrock (example values in blue)
and depth to groundwater (example values in red) on page 1 of field forms.
References
Brunke, M. and T. Gonser. 1997. The
ecological significance of exchange
processes between rivers and groundwater.
Freshwater Biology 37:1-33.
Boulton, A.J., S. Findlay, P. Marmonier,
E.H. Stanley, and H.M. Valett. 1998. The
functional significance of the hyporheic
zone in streams and rivers. Annual Review
of Ecology and Systematics 29:59-81.
Clinton, S.M., N. B. Grimm, and S.G.
Fisher. 1996. Response of a hyporheic
invertebrate assemblage to drying
disturbance in a desert stream. Journal of
the North American Benthological Society
15:700-712.
Dole-Olivier, M.-J., P. Marmonier, and J.-L.
Beffy. 1997. Responses of invertebrates
to lotic disturbance: is the hyporheic zone
a patchy refugium? Freshwater Biology
37:257-276.
78
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Huggenberger, P., E. Hoehn, R. Beschta,
and W. Woessner. 1998. Abiotic aspects
of channels and floodplains in riparian
ecology. Freshwater Biology 40:407-425.
Jones, J.B. andP.J. Mulholland. 2000.
Streams and Ground Waters. Academic
Press, San Diego, California.
Naegeli, M. W., P. Huggenberger, U.
Uehlinger. 1996. Ground penetrating
radar for assessing sediment structures in
the hyporheic zone of a prealpine river.
Journal of the North American
Benthological Society 15:353-366.
Poole, G. C., R. J. Naiman, J. Pastor, and J.
A.Stanford. 1997. Uses and limitations
of ground penetrating RADAR in two
riparian systems. Pages 140-148 in: J.
Gibert, J. Mathieu, and F. Fournier
(editors). Groundwater/Surface Water
Ecotones: Biological and Hydrological
Interactions and Management Options.
Cambridge University Press, Cambridge,
United Kingdom.
Valett, H. M. 1993. Surface-hyporheic
interactions in a Sonoran Desert stream:
hydrologic exchange and diel periodicity.
Hydrobiologia 259:133-144.
Equipment and supplies
Measuring tape (50 m)
Sounding rod (steel rod, > 3 ft or 91 cm) or
"T"-bar
Hand sledge hammer
Meter stick
Field forms
3.14 Gravimetrically measuring streambed
sediment moisture
General
This subsection provides instructions for
measuring the relative moisture of
streambed sediments in dry headwater
channels. In other words, this procedure
quantifies the degree of "dryness" or
desiccation of the benthic habitat in streams
when visible surface water is absent. This is
especially relevant to organisms that inhabit
intermittent or ephemeral streams and
possess life histories or physiological traits
(i.e., diapause, quiescence, and aestivation)
for surviving the dry periods (Davis 1972,
McKee and Mackie 1983, Danks 1987,
Williams 1998, Dunphy et al. 2001).
Mortality during such periods can depend on
the desiccation level of the surrounding
sediments, and therefore can influence the
spatial distribution of organisms (Suemoto
et al. 2005). Soil moisture is measured
during the summer sampling period (when
sediment moisture is expected to be lowest)
within study reaches that have no visible
surface water.
Procedure
Delineate the 30-m study reach so that a
measuring tape is positioned along the
thalweg. Mark three 15-cm pieces of 3/4"
PVC with a line 10 cm from one end using a
permanent marker (volume -28.5 cm3).
3.14.1 Sediment collection
Three individual sediment cores are taken
along study reaches lacking visible surface
water. Cores are extracted from the
streambed in depositional habitat units with
fine sediments (e.g., sand, silt, fine gravel).
In addition to being more feasible to collect,
moisture content is expected to be relatively
high in thick patches of fine sediment (i.e.,
capillary fringe) because of greater capillary
tension compared to levels associated with
coarser particles (Dunne and Leopold 1978).
Where possible, cores from each study reach
79
-------
should be taken from separate depositional
units within the thalweg.
Find a suitable location and brush aside
detritus (i.e., leaf litter) from the streambed
surface. Position the core vertically so that
the 10-cm mark is away from the streambed
(Figure 3-54). Tap the core vertically into
the streambed with the hand sledge until the
Upper core
Streambed surface
Figure 3-54 Sampling sediment moisture.
10-cm mark is flush with the streambed
(Figure. 3-55). Place a rubber stopper into
the upper core opening. Carefully pull the
core out of the streambed and place a second
rubber stopper into the lower core opening.
Place core in a resealable plastic bag. Label
the bag and/or core using a permanent
marker with relevant information (e.g.,
locality, date, collector's initials). Remove
excess air within the bag when sealing. Log
the number of sediment core samples taken
at each site on the field forms (Figure 3-56).
Store samples in a cooler with ice or in a
refrigerator until the samples can be
measured in the laboratory. Measure
moisture of the sample within 4 days of
collection. A soil borer or auger can be used
collect samples rather than PVC cores. Care
must be taken to keep sediment samples
airtight (e.g., Shelby tube) to maintain soil
moisture levels.
Figure 3-55 Tapping core vertically into
streambed.
PRESENCE OF
HEADCUT IN REACH
Y
N
ALGAL COVER INDEX
1 I1'2
2
3
4 5
# CORES FOR SUBSTRATE MOISTURE
(depositional)
3
Figure 3-56 Appropriate location for recording the number of sediment moisture cores
collected on page 1 of field forms.
3.14.2 Laboratory measurement
In the laboratory, use a weighing spatula or
thin metal rod to transfer sediment from cores
into separate evaporating dishes or crucibles.
Be sure that the dishes are uniquely identified
(e.g., dish #s), so that results can be associated
with specific samples. Measure the wet
weight of the sediment samples with an
analytical balance to the nearest O.Olg.
Record dish identification, sample
abbreviation, and wet weight on the data
sheets (Figure 3-57).
80
-------
HISS Sediment Moisture Data Sheet DATE: 9/23/03 Page 1 of 1
Dish#
123
144
135
142
Study Site Abbrev.
Four-FC-3
Four FC-3
FourFC-3
Four-FC-4
Sediment Core
A
B
C
A
Wet Weight (g)
39.36
38.47
38.95
29.29
Dry Weight (g)
Figure 3-57 Example of the sediment moisture data sheet.
Place samples into the drying oven for 24 h
with temperature set at 90° C. Remove
samples from the oven using tongs and allow
them to cool to room temperature. If a
desiccator is available, the samples can be
directly placed into the desiccator to cool.
Measure the dry weight of the sediment
samples with the balance to the nearest O.Olg
and record on the data sheet. Percent moisture
is calculated using the following equation:
Percent Moisture =
Wet Weight - Dry Weight
Cores can be collected from locations
where/when surface water is present to
provide a relative comparison of sediment
moisture. Alternatively, water can be added to
previously dried samples until visibly
saturated. The cores are then weighed to
determine percent moisture at saturation. The
amount of organic matter within the sediments
can be determined by ashing the core contents
in a muffle furnace at 550° C for two hours
and then reweighing to determine ash-free dry
mass (AFDM).
Alternative means for measuring soil moisture
include the use of soil moisture probes (e.g.,
tensiometers, capacitance sensors; see Miller
et al. 1997); but these are not commonly used
in the relatively coarse sediments of
intermittent streambeds. A procedure
described by Greacen et al. (1989) indirectly
measures sediment moisture by way of water
absorption onto filter paper and then
gravimetric determination of water content.
Techniques that have been used to extract
water from soil cores include centrifugation,
squeezing, and vacuum extraction (e.g.,
Adams etal. 1980).
Wet Weight
x 100
References
Adams, F. C., C. H. Burmester, N. H. Hue,
and F.L. Long. 1980. A comparison of
column displacement and centrifuge
methods for obtaining soil solutions. Soil
Science Society of America Journal. 44:733-
735.
Banks, H. V. 1987. Insect Dormancy: An
Ecological Perspective. Biological Survey
of Canada Monograph series No. 1., Ottawa,
Canada.
Davis, J. S. 1972. Survival records in the
algae, and the survival role of certain algal
pigments, fat, and mucilaginous substances.
The Biologist 54:52-93.
Dunne, T. and L. B. Leopold. 1978. Water in
Environmental Planning. W. H. Freemann
and Company, New York.
Dunphy, M. E., D. C. McDevit, C. E. Lane,
and C. W. Schneider. The survival of
Vaucheria (Vaucheriaceae) propagules in
81
-------
desiccated New England riparian sediment.
Rhodora 103:416-426.
Greacen, E. L., G. R. Walker, and P. G. Cook.
1989. Procedure for the Filter Paper
Method of Measuring Soil Water Suction.
Division of Soils Division Report 108,
CSIRO, Melbourne.
McKee P. M. and G. L. Mackie. 1983.
Respiratory adaptations of the fingernail
clams Sphaerium occidentale and
Musculium secures to ephemeral habitats.
Canadian Journal of Zoology 40:783 -791.
Miller, J. D., G. J. Gaskin, and H. A.
Anderson. 1997. From drought to flood:
catchment responses revealed using novel
soil water probes. Hydrological Processes
11:533-541.
Suemoto, T., K. Kawai, and H. Imabayashi.
2005. Dried-up zone as a temporal stock of
chironomid larvae: survival periods and
density in a reservoir bank. Hydrobiologia
54:145-152.
Williams, D. D. 1998. The role of dormancy
in the evolution and structure of temporary
water invertebrate communities. Archivfur
Hydrobiologie Special Issue, Advances in
Limnology 52:109-124.
Equipment and supplies
Measuring tape (50 m)
Cores (15 cm long, % inch inner diameter
PVC pipe)
Hand sledge hammer
Rubber stoppers (2 per core, No. 1 or 2)
Resealable plastic bags (1 per core)
Cooler
Ice or ice packs
Permanent marker
Field forms
Weighing spatula or metal rod (laboratory)
Evaporating dishes or crucibles (laboratory)
Drying oven (laboratory)
Analytical balance (laboratory)
Desiccator (laboratory)
Lab notebook or bench sheets (laboratory)
3.15 Characterizing the size distribution of
streambed sediments
General
This subsection provides a simple method for
characterizing the size structure of streambed
sediments within headwater stream reaches.
Sediment characteristics influence many other
physical properties, including habitat stability,
interstitial habitat volume, nearbed velocities,
organic matter retention, and re-aeration.
Consequently, streambed sediments directly
and indirectly influence community structure
and stream processes. The characteristics of
the streambed are expected to influence
stream processes to a greater degree in
headwater streams than in larger rivers
because headwaters have a higher ratio of
streambed surface area to instantaneous flow
volume (m2/m3) than larger streams and rivers.
Geology, climate, topography, and drainage
area are factors that naturally govern the
natural composition of stream sediments.
Land-use changes can cause deleterious
alteration to streambed properties (e.g.,
siltation) and subsequent shifts in biological
integrity.
Particle size is the most common measure
used to characterize streambed sediments,
mainly because of the ease to which it can be
objectively quantified compared to other
characteristics (e.g., sphericity, specific
density). A frequently used method to
characterize sediments on streambed surfaces
is the Wolman pebble count procedure
(Wolman 1954, Leopold 1970, Kondolf & Li
1992), where the sizes of individual stones are
randomly selected and measured along a
82
-------
reach. Vertical characterization can be done
by coring (Cummins 1962, Everest et al. 1980,
Wesche et al. 1989) and ground-penetrating
radar (Naegeli et al. 1996, Huggenberger et al.
1998). Other aspects of the streambed
sediments that have been measured include
texture (Downes et al. 1998, Bergey 1999),
porosity (Maridet et al. 1992), bed roughness
(Statzner 1981, Ziser 1985), topographic
complexity or fractal geometry (Schmid 2000,
Robson et al. 2002, Stewart and Garcia 2002)
and stability (Biggs et al. 1997, Duncan et al.
1999). The composition of streambed
sediments influences aspects related to the rate
of stream drying (i.e., permeability), wetted
surface area as stream levels decline (boulder-
dominated reaches will have more emergent
sediments at low flows than gravel reaches),
and the availability of interstitial refugia when
streams are dry.
The protocol below is based on methods
described in Walters et al. (2003) for particle
size characterization by patches rather than
individual grains or stones. Streambed
sediment characterization is measured once
for a given reach during the study because
reach-level particle size measures are unlikely
to change significantly over the timeframe of
most ecological studies (1-2 years).
Procedure
Streambed surface sediments are measured at
31 locations, longitudinally at every meter
mark along the thalweg of each 30-m study
reach (Figure 3-58). Each particle size
measurement is based upon 0.25 m2 patches of
particles, rather than a single particle
measurement. The patches are centered
around each meter mark (0, 1, 2,... 30 m)
along the study reach thalweg. The modal
particle size class or the size class with the
greatest patch coverage is estimated for each
patch location. Once the patch is located,
visually assess the size classes within each
patch, determine which size class has the
greatest coverage, and select a representative
particle of that size class. The dimension used
to determine particle size is the intermediate
axis (i.e., p-axis) or the median value among
the length, width, and height of the particle.
Exact measurement of the intermediate axis is
not needed because size classes are used.
Particle size classes are based upon the
Wentworth size classification or phi (O) scale
(Cummins 1962, Table 3-1). The value for
sediment particle size to be entered in the field
form (Figure 3-59) is the upper bound value of
the size class (bold-faced values in Table 3-1).
The particle size classes are also listed on the
bottom of page 3 of the field forms.
83
-------
30m
Modal particle size
class = 128 mm
Figure 3-58 Schematic of study reach illustrating thalweg (dotted line) and patch locations for determining modal sediment
particle size class. Inset provides a close-up of a patch (overlaid) with measuring tape used in designating patch locations
longitudinally along the study reach).
84
-------
Table 3-1 Modified Wentworth scale for sediment particle size classes. Bold-faced
numbers indicate values to be entered on field forms
_Class
Sand, silt, and clay
Fine gravel
Medium gravel
Coarse gravel
Small pebble
Large pebble
Small cobble
Large cobble
Boulder
Bedrock and hardpan
Size_range (mm) Phi ()
<2 >0
>2to4 -lto-2
>4 to 8 -2 to -3
>8 to 16 -3 to -4
>16to32 -4 to-5
>32 to 64 -5 to -6
>64 to 128 -6 to -7
>128 to 256 -7 to -8
>256tp512 -8 toi-9
~>512 <-9
References
Bergey, E. A. 1999. Crevices as refugia for
stream diatoms: effects of crevice size on
abraded substrates. Limnology and
Oceanography 44:1522-1529.
Biggs, B. J. F., M. J. Duncan, S. N. Francoeur,
and W. D. Meyer. 1997. Physical
characterisation of microform bed cluster
refugia in 12 headwater streams, New
Zealand. New Zealand Journal of Marine
and Freshwater Research 31:413-422.
Cummins, K. W. 1962. An evaluation of
some techniques for the collection and
analysis of benthic samples with special
emphasis on lotic waters. American Midland
Naturalist 67:477-504.
Downes, B. J., P. S. Lake, E. S. G. Schreiber,
and A. Glaister. 1998. Habitat structure and
regulation of local species diversity in a
stony, upland stream. Ecological
Monographs 68:237-257.
Duncan, M. J., A. M. Suren, and S. L. R.
Brown. 1996. Assessment of streambed
stability in steep, bouldery streams:
development of a new analytical technique.
Journal of the North American
BenthologicalSociety 15: 445-456.
Everest, F. H., C. E. McLemore, and J. F.
Ward. 1980. An Improved Tri-tube
Cryogenic Gravel Sampler. Research Note
PNW-350, U.S. Forest Service, Northwest
Forest and Range Experiment Station,
Portland, OR.
Kondolf, G. M. and S. Li. 1992. The pebble
count technique for quantifying surface bed
material size in intstream flow studies.
Rivers 3:80-87.
Leopold, L. B. 1970. An improved method
for size distribution of stream-bed gravel.
Water Resources Research 6:1357-1366.
Maridet, L., J.-G. Wasson, and M. Philippe.
1992. Vertical distribution of fauna in the
bed sediment of three running water sites:
influence of physical and trophic factors.
Regulated Rivers: Research & Management
7:45-55.
Naegeli, M. W., P. Huggenberger, U.
Uehlinger. 1996. Ground penetrating radar
for assessing sediment structures in the
hyporheic zone of a prealpine river. Journal
85
-------
of the North American Benthological Society
15:353-366.
American Journal of Fisheries Management
9:234-238.
Ohio EPA. 2002. Field Evaluation Manual
for Ohio's Primary Headwater Habitat
Streams. Final Version 1.0. Ohio
Environmental Protection Agency, Division
of Surface Water, Columbus, Ohio.
http ://www. epa. state, oh.us/dsw/wqs/headwa
ters/PHWHManual 2002 102402.pdf
Robson, B. J., E. T. Chester, and L. A.
Barmuta. 2002. Using fractal geometry to
make rapid field measurements of riverbed
topography at ecologically useful spatial
scales. Marine and Freshwater Research
53:999-1003.
Schmid, P. E. 2000. Fractal properties of
habitat and patch structure in benthic
ecosystems. Advances in Ecological
Research 30:339-401.
Statzner, B. 1981. A method to estimate the
population size of benthic
macroinvertebrates in streams. Oecologia
51:157-161.
Stewart, T. W. and J. E. Garcia. 2002.
Environmental factors causing local
variation in density and biomass of the snail
Leptoxis carinata in Fishpond Creek,
Virginia. American Midland Naturalist
148:172-180.
Walters, D.M., D.S. Leigh, M.C. Freeman,
BJ. Freeman, and C.M. Pringle. 2003.
Geomorphology and fish assemblages in a
Piedmont river basin, U.S.A. Freshwater
Biology 48:1950-1970.
Wesche, T. A., D. W. Reiser, V. R.
Hasfurther, W. A. Hubert, and Q. D.
Skinner. 1989. New technique for
measuring fine sediment in streams. North
Wolman, M. G. 1954. A method of sampling
coarse river-bed material. Transactions of
the American Geophysical Union 35:951-
956.
Ziser, S. W. 1985. The effects of a small
reservoir on the seasonality and stability of
physiochemical parameters and
macrobenthic community structure in a
Rocky Mountain stream. Freshwater
Invertebrate Biology 4:160-177.
Equipment and supplies
Meter stick or ruler
Measuring tape (50m)
Field forms
86
-------
* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2BF Page 2 of 4
Meter #
0
i
2
3
4
5
6
7
8
9
10
11
12
13
14
Modal
Sediment
Particle
Size
(mm)*
<2
<2
4
16
>512
>512
16
64
64
<2
<2
32
8
>512
256
(^•^128
Water
Depth
(cm)*
Habitat
Type
(E/D)
Notes
(e.g., LWD,
Leafpack)
Velocity
(m/s)*
Wetted
Width
(m)
BE
Width
(m)
BE
Depth
(m)
ft
EPA
width
(m)§
Figure 3-59 Appropriate location for recording modal particle size data on page 2 of field
forms (example from Figure 3-58 highlighted).
3.16 In situ water chemistry measurements
General
This subsection provides procedures for
measuring in situ water chemistry of
headwater streams. The basic water chemistry
measurements discussed in this section are: 1)
temperature, 2) conductivity, 3) pH, and 4)
dissolved oxygen. Instructions for collecting
water samples and measuring additional
chemical parameters (i.e., nutrients, cations,
anions) can be found in Wetzel and Likens
(1991), Herlihy (1998), and APHA (2005).
Because characteristics of water change with
residence time, these measurements may be
useful in distinguishing between groundwater
and throughflow (i.e., water in unsaturated soil
zones during and immediately after
precipitation). Physicochemical amplitudes
(seasonal and diel) are typically greater in
temporary waterbodies than in perennial
counterparts (Zale et al. 1989, Boulton et al.
2000). As flow begins to decline, deeper
groundwater inputs may represent the
dominant source of surface flows, resulting in
relatively subtle physicochemical shifts
(Dahm et al. 2003). More dramatic changes in
water physiochemistry can occur as
waterbodies dry, and such changes can have
equally dramatic effects on the inhabiting
biota (Moore and Burn 1968, Magoulick and
87
-------
Kobza 2003). Maximum diel variation and
absolute extremes are commonly measured
when surface water becomes limited to
disconnected pools (Stehr and Branson 1938,
Boulton and Lake 1990) and depending upon
the pool depth, vertical stratification can occur
(e.g., Neel 1951, Wood et al. 1992).
Conductivity and water temperature typically
increases as streams dry (e.g., Baron et al.
1998), whereas dissolved oxygen tends to
decrease (e.g., Slack and Feltz 1968, Chapman
and Kramer 1991). Declines in water volume
from evaporation and evapotranspiration lead
to greater water surface area to volume ratios
that subsequently cause water temperatures to
rise from rapid solar heating. Warmer water
and contraction of surface water intensifies
community respiration that can lead to
declines in dissolved oxygen. Evaporation,
increased soil residence time, and organic
matter breakdown elevates stream water
concentrations of dissolved ions and alters pH
(Williams and Melack 1997, Hamilton et al.
2005). The buffering capacity (or acid-
neutralizing capacity, ANC) of stream water
will determine the direction of pH change
during drying. In some streams, high leachate
concentrations from organic matter may
decrease pH (Slack and Feltz 1968). Increases
in pH during dry seasons can occur where
ANC is strongly influenced by acid rain or
snowmelt during wet seasons (Wigington et
al. 1996) or where stream water is naturally
low in base cations (e.g., Ca++, K+, Mg++) and
drying concentrates strong acid anions (e.g.,
SO4", Cr, NCV, Bayley et al. 1992).
Although water quality can decline with
drying, these changes may be mitigated where
there is intact forest to buffer the stream
environment (e.g., Feminella 1996).
Conversely, reduced flows and drying
exacerbates water quality problems in areas
with nutrient input and removal of riparian
canopy (Casey and Ladle 1976, Chessman and
Robinson 1987), particularly if remaining
flow is effluent-dominated (e.g., Lewis and
Burraychak 1979, Jennings and Gasith 1993,
Suren et al. 2003, Brooks et al. 2006).
Because in situ water chemistry can vary
considerably over time, measurements should
be taken during each sampling visit. Note that
the following procedure is for taking point
measurements rather than measuring diel
variation or extremes.
Procedure
Before arriving at the field sites read the
instruction manual for portable meters and
check batteries. Check to see that the meters
are functioning properly and are calibrated.
Use standards to calibrate meters at least
daily. Record pre- and post-calibration values
on the instrument log sheet (Figure 3-60).
Calibrate the dissolved oxygen meter for the
appropriate elevation for each study site
(elevation can be read from the 7.5 min.
topographic maps, or GPS units). Suggested
data quality objectives (DQO) for in situ water
chemistry are shown in Table 3.16.1.
88
-------
Date
9/22/03
9/22/03
Instrument
Hydrolab
Quanta (#2)
Hydrolab
Quanta (#2)
Inspected
by
KMF
KMF
Pass
inspection
(Y or N)
Y
Y
Degree deviated
from calibration
standard
(+ or - include
units)
pH
@3:+0.1
@ 7: +0.2
Cond
@45:+7|iS
@ 147: -12|iS
Recalibrated
(YorN)
Y
Y
Figure 3-60 An example of an instrument inspection and calibration log sheet.
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg. In situ water chemistry
measurements should be taken before all other
measurements. Note the location and time of
measurements on the field form (Figure 3-61).
If the pH, conductivity, and dissolved oxygen
meters also measure temperature, consistently
use one of these to measure temperature.
When available, submerge probes in the area
of flowing water (note that some probes
cannot be completely submerged) and monitor
the readout until values stabilize. Where
hydrologic condition is "surface water in
pools only" (see Section 3.1 for designation of
hydrologic condition), in situ water chemistry
should be measured in all pools where
biological samples are taken. Write values for
measurements in the appropriate cells on the
field form (Figure 3.16.2). Record time of day
when measurements were taken in
"comments" section. If additional space is
needed use space on page 3 of the field form.
Turn off meters and then repeat measurements
to meet DQO in Table 3.16.1. If repeat
measurements do not meet DQO standards
then flag those values on the field forms to
indicate that they are suspect.
Table 3-2 Data Quality Objectives (DQO) for in situ water chemistry measurements
Measurement
Data Quality Required
Temperature
PH
Dissolved Oxygen (DO)
Conductivity
two measurements taken with less than 5% deviation.
two measurements with less than 10% deviation
two measurements with less than 10% deviation
two measurements with less than 10% deviation
89
-------
IN SITU WATER QUALITY MEASUREMENTS
Location of
Measurements
10m
Cond
OS/cm)
24
Temp
(°C)
10
DO
(mg/1)
1.23
PH
6.3
Comments
@ 9:30 am
Figure 3-61 Appropriate locations for recording in situ water quality measurements on
page 1 of field forms, example values shown in red.
References
American Public Health Association,
American Water Works Association, and
Water Environment Federation. 2005.
Standard Methods for the Examination of
Water and Wastewater, 21st edition.
American Public Health Association,
Washington, D.C.
Baron, J. S., T. LaFrancois, and B. C.
Kondratieff. 1998. Chemical and biological
characteristics of desert rock pools in
intermittent streams of Capitol Reef
National Park, Utah. Great Basin Naturalist
58:250-264.
Bayley, S. E., D. W. Schindler, B. R. Parker,
M. P. Stainton, and K. G. Beaty. 1992.
Effects of forest fire and drought on acidity
of a base-poor boreal forest stream:
similarities between climatic warming and
acidic precipitation. Biogeochemistry
17:191-204.
Boulton, A. J. and P. S. Lake. 1990. The
ecology of two intermittent streams in
Victoria, Australia. I. Multivariate analyses
of physiochemical features. Freshwater
Biology 24:123-141.
Boulton, A. J., F. Sheldon, M. C. Thorns, and
E. H. Stanley. 2000. Problems and
constraints in managing rivers with variable
flow regimes. Pages 415-430 in P. J. Boon,
B. R. Davies, and G. E. Petts (editors).
Global Perspectives on River Conservation:
Science, Policy and Practice. John Wiley &
Sons, Chichester, United Kingdom.
Brooks, B. W., T. M. Riley, and R. D. Taylor.
2006. Water quality of effluent-dominated
ecosystems: ecotoxicological, hydrological,
and management considerations.
Hydrobiologia 556:365-379.
Casey, H. and M. Ladle. 1976. Chemistry
and biology of the South Winterbourne,
Dorset, England. Freshwater Biology 6:1-
12.
Chapman L. J. and D. L. Kramer.
Limnological observations of an intermittent
tropical dry forest stream. Hydrobiologia
226:153-166.
Chessman, B. C. and D. P. Robinson. 1987.
Some effects of the 1982-83 drought on
water quality and macroinvertebrate fauna in
the Lower LaTrobe River, Victoria.
Australian Journal of Marine and
Freshwater Research 38:289-299.
Dahm, C. N., M. A. Baker, D. I. Moore, and J.
R. Thibault. 2003. Coupled
biogeochemical and hydrological responses
of streams and rivers to drought. Freshwater
Biology 48:1219-1231.
Feminella, J. W. 1996. Comparison of
benthic macroinvertebrate assemblages in
small streams along a gradient of flow
90
-------
permanence. Journal of the North American
Benthological Society 15:651-669.
Hamilton, S. K., S. E. Bunn, M. C. Thorns, J.
C.Marshall. 2005. Persistence of aquatic
refugia between flow pulses in a dryland
river system (Cooper Creek, Australia).
Limnology and Oceanography 50:743-754.
Herlihy, A. T. 1998. Water chemistry. Pages
57-65 in: J.M. Lazorchak, DJ. Klemm, and
D.V. Peck (editors). Environmental
Monitoring and Assessment Program -
Surface Waters: Field Operations and
Methods for Measuring the Ecological
Condition of Wadeable Streams.
EPA/620/R-94/004F. U.S. Environmental
Protection Agency, Washington, D.C.
Jennings, J.-J. and A. Gasith. 1993. Spatial
and temporal changes in habitat condition in
the Na'aman stream ecosystem, Israel.
Water Science and Technology 27:387-395.
Lewis, M. A. and R. Burraychak. 1979.
Impact of copper mining on a desert
intermittent stream in central Arizona.
Journal of the Arizona-Nevada Academy of
Science 14:22-29.
Moore, W. G. and A. Burn. 1968. Lethal
oxygen thresholds for certain temporary
pond invertebrates and their applicability to
field situations. Ecology 49:349-351.
Neel, J. K. 1951. Interrelations of certain
physical and chemical features in a
headwater limestone stream. Ecology
32:368-391.
Slack, K. V. and H. R. Feltz. 1968. Tree leaf
control on low flow water quality in a small
Virginia stream. Environmental Science and
Technology 2:126-131.
Stehr, W. C. and J. W. Branson 1938. An
ecological study of an intermittent stream.
Ecology 19:294-310.
Suren, A. M., B. J. Biggs, C. Kilroy, and L.
Bergey. 2003. Benthic community
dynamics during summer low-flows in two
rivers of contrasting enrichment 1.
Periphyton. New Zealand Journal of Marine
and Freshwater Research 37:53-70.
Wetzel, R. G. and G. E. Likens. 1991.
Limnological Analyses, 2nd edition.
Springer-Verlag, New York.
Wigington, P. J., Jr., D. R. DeWalle, P. S.
Murdock, W. A. Krester, H. A. Simonin, J.
Van Sickle, and J. P. Baker. 1996.
Acidification of small streams in the
northeastern United States: ionic controls of
episodes. Ecological Applications 6:389-
407.
Williams, M. R. and J. M. Melack. 1997.
Effects of prescribed burning and drought on
the solute chemistry of mixed-conifer forest
streams of the Sierra Nevada, California.
Biogeochemistry 39:225-253.
Wood, D. J., S. G. Fisher, and N. B. Grimm.
1992. Pools in desert streams: limnology
and response to disturbance. Journal of the
Arizona-Nevada Academy of Science
26:171-179.
Zale, A. V., D. M. Leslie, W. L. Fisher, and S.
G. Merrifield. 1989. The Physicochemistry,
Flora, and Fauna of Intermittent Prairie
Streams: A Review of the Literature.
Biological Report 89(5), US Department of
Interior, Fish and Wildlife Service.
Equipment and supplies
Measuring tape (50 m)
pH meter
91
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Conductivity meter
Dissolved oxygen meter
Thermometer
Associated calibration standards for meters
Field form
Spare batteries
3.17 Measuring riparian canopy cover
General
This subsection provides instructions for
measuring riparian canopy cover for
headwater streams. Canopy cover is a useful
measure of riparian condition that can strongly
influence the structure (e.g, organic substrate,
algal biomass) and function (e.g., primary
production) of streams (Gregory et al. 1991,
Naiman and Decamps 1997). This procedure
is a modification of the original method
described by Lemmon (1957) for use with a
convex spherical densiometer. Measurements
of irradiance with pyrheliometers or
photosynthetically active radiation with
quanta sensors provide quantitative measures
of incoming solar energy (Moulton et al 2002,
also see reviews by Hauer and Hill 1996,
Jennings et al. 1999). A disadvantage of these
measures is their sensitivity to cloud cover and
angle of the sun. Another method for
estimating canopy cover is the use of fisheye
or hemispheric photography (Davies-Colley
and Payne 1998, Ringold et al. 2003, Kelly
and Krueger 2005). Especially with the
advent of digital photography and analytical
software this method offers short processing
times, consistency, and precision. One
limitation of photographic methods is ensuring
proper lighting conditions. Direct overhead
sunlight, reflection on vegetation, and dark
clouds can lead to data misinterpretation
(Kelly and Krueger 2005). Measurements of
canopy cover are taken during each season
(spring and summer) because this will likely
change through time.
Procedure
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg. Canopy cover is measured while
facing upstream, downstream, left bank, and
right bank at the 15-m mark of the study
reach.
Canopy measurements are taken by holding
the densitometer about 0.3 m above the stream
surface at the thalweg. Level the densitometer
using the bubble level and position it so that
your reflection is just below the mirror grid
(Figure 3-62). Calculate the percent cover by
first identifying the grid intersections (of 37
total intersections) that are covered by
vegetation (e.g., leaves, branches, trunks) or
stream banks. Percent cover values for
intersections are equivalent to the number of
squares meeting at an intersection (Figure 3-
62), ranging from 1% to 4%. For example, an
intersection where 4 squares meet and is
covered by vegetation is equivalent to 4%
cover (Note that based on this value system,
the total percentage is 98% and therefore
approximates 100%.). Sum percent cover and
record in the appropriate cell on the field form
(Figure 3-63).
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Bubble
level
Figure 3-62 Plan view of a convex spherical densitometer, showing percent cover values
associated with intersections. Values are equivalent to the number of squares meeting at
each intersection.
Where canopy over the stream channel is
heavy, it is more efficient to measure the
percentage of open by identifying and
summing grid intersections values that are not
covered by vegetation, etc. Percent canopy
cover is then simply calculated by subtracting
total percent open canopy from 100%.
DISTANCE TO NEAREST
SURFACE WATER (m)
0 <100 100-500 >500
CHANNEL SLOPE (%)
(for three 10 in sections of study reach)
% CANOPY COVER
( facing upstream, downstream, right & left
banks
Figure 3-63 Appropriate location for recording percent canopy cover on page 1 of field
forms.
The methods used by USEPA's EMAP and
USGS's National Water-Quality Monitoring
Program (NAWQA) differ slightly from the
method discussed above. Rather than using
all 37 intersections on the convex mirror for
measurements, only 17 intersections are
evaluated (Figure. 3-64, Fitzpatrick et al.
1998, Kaufmann and Robison 1998). A "V"
is taped on the mirror surface to delimit the 17
intersections. This modification is intended to
minimize repeated observations of cover
structures during multiple readings from the
same position (e.g., facing upstream,
downstream, left and right bank) and reduces
measurement time (Strichler 1959). Each
intersection is weighted equally, rather than by
the number of squares meeting at the
intersections. The number of covered
intersections is recorded for measurements
facing upstream, downstream, left and right
banks (standing at mid-channel) for the 11
transects (per study reach) in the EMAP
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protocol (Kaufmann and Robison 1998),
whereas two measurements, facing the each
bank at the water's edge, are taken at the 11
transects (per study reach) in the National
Water Quality Assessment (NAQWA)
protocol (Fitzpatrick et al. 1998). The
NAQWA protocol measures canopy closure,
rather than canopy cover (also called canopy
density). Canopy closure includes the
overhead area bracketed by vegetation,
whereas canopy density includes only area of
sky completely blocked by vegetation.
Canopy closure is intended to be less
influenced by season (i.e., leaf abscission)
than canopy density (Strichler 1959). For
both protocols, percent cover or closure is
calculated as the ratio of covered to total
intersections.
Bubble
level
Figure 3-64 Plan view of a convex spherical densitometer, modified for measuring over 17
intersections (open circles) that are delimited by a "V" taped to the convex mirror.
References
Davies-Colley, R. J. and G. W. Payne. 1998.
Measuring stream shade. Journal of the
North American Benthological Society
17:250-260.
Fitzpatrick, F. A., I. R. Waite, P. J. D'Arconte,
M. R. Meador, M. A. Maupin, and M. E.
Gurtz. 1998. Revised Methods for
Characterizing Stream Habitat in the
National Water-Quality Assessment
Program. U.S. Geological Survey Water-
Resources Investigations Report 98-4052,
Raleigh, North Carolina.
Gregory, S.V., FJ. Swanson, W.A. McKee,
and K.W. Cummins. 1991. An ecosystem
perspective of riparian zones. BioScience
41:540-551.
Hauer, F. R. and W. R. Hill. 1996.
Temperature, light, and oxygen. Pages 93-
106 in: F.R. Hauer and G. A. Lamberti
(editors). Methods in stream ecology.
Academic Press, San Diego, California.
Jennings, S. B., N. D. Brown, and D. Sheil.
1999. Assessing forest canopies and
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understorey illumination: canopy closure,
canopy cover and other measures. Forestry
72: 59-73.
Kaufmann, P. R. and E. G. Robison. 1998.
Physical habitat characterization. Pages 77-
118 in J.M. Lazorchak, DJ. Klemm, and
D.V. Peck (editors). Environmental
Monitoring and Assessment Program -
Surface Waters: Field Operations and
Methods for Measuring the Ecological
Condition of Wadeable Streams.
EPA/620/R-94/004F. U.S. Environmental
Protection Agency, Washington, D.C.
Kelley, C. E. and W. C. Krueger. 2005.
Canopy cover and shade determinations in
riparian zones. Journal of the American
Water Resources Association 41:37-46.
Lemmon, P.E. 1957. A new instrument for
measuring forestry overstory density.
Journal of Forestry 55:667-669.
Moulton, S. R. II, J. G. Kennen, R. M.
Goldstein, and J. A. Hambrook. 2002.
Revised Protocols for Sampling Algal,
Invertebrate, and Fish Communities As Part
of the National Water-Quality Assessment
Program. Department of Interior, U.S.
Geological Survey Open-File Report 02-
150, 75 p.
Naiman, RJ. and H. Decamps. 1997. The
ecology of interfaces: riparian zones. Annual
Review of Ecology and Systematics 28:621-
658.
Ringold, P. L., J. Van Sickle, K. Rasar, J.
Schacher. 2003. Use of hemispheric
imagery for estimating stream solar
exposure. Journal of the American Water
Resources Assoication 39:1373-1384.
Strichler, G. S. 1959. Use of the
Densitometer to Estimate Density of Forest
Canopy on Permanent Sample Plots. USDA
Forest Service, Pacific Northwest Forest and
Range Experimental Station Research Note
180, Portland, Oregon.
Equipment and supplies
Measuring tape (50 m)
Convex spherical densiometer
Field forms
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4 BIOLOGICAL SAMPLING
This section provides background material
related to biological sampling in headwater
streams. The incorporation of biomonitoring
(i.e., the use of organisms to evaluate changes
in an environment) into assessment programs
is advocated because biota are ubiquitous
across the landscape, represent a diversity of
responses, integrate stressor effects over time,
and are relevant to societal needs (Rosenberg
and Resh 1993, Whitton and Kelly 1995).
The primary biological levels used in
biomonitoring are community and population
levels, although biological measurements can
range from molecules to ecosystems. The
response measures in bioassessment are
typically abundance, biomass, and diversity;
however, there is a trend toward quantifying
characteristics (i.e., species traits) and
functional roles of biota for predicting
biological responses to specific disturbances
or stressors. Some of these traits include life
span, maximum size, phenology, and
physiology (Muotka and Virtanen 1995, Biggs
et al. 1998, Charvet et al. 1998, Usseglio-
Polatera et al. 2000). Species traits may
provide important insight in understanding
stressor-specific responses and have a place in
bioassessment, as do tolerance values (e.g.,
Hilsenhoff 1987, Van Dam et al. 1994) and
functional feeding groups.
The choice of biological level and group
should match the study objectives. Be aware
of attributes and limitations of particular
taxonomic groups. For example, primary
producers will respond immediately to
changes in light and nutrients, whereas a lag-
response is expected for consumers. Long-
term monitoring of individuals is possible for
most bryophytes, but not so for invertebrates
with relatively short life spans. In addition,
ethical and legal considerations (e.g.,
sampling permits) are more prevalent for
some biota than others. Particular sampling
regimes may also be more conducive to some
groups than others. For instance, organisms
with a patchy distribution may require larger
sample areas (or more samples) than those
with a uniform distribution.
As discussed for physical habitat assessments,
methods of biological sampling can range
from qualitative to quantitative. Sampling
methods should match the investigators' study
objectives. Objectivity, comparability and
precision of the methods increase as the level
of quantification increases. For example,
quantitative methods measure biota over a
specified area or volume (e.g., Hess sampler)
with greater precision and repeatability than
semi-quantitative (e.g., kick nets) or
qualitative methods (e.g., dip net jabs).
However, the level of effort (especially time)
and training may increase with more
quantitative methods. Therefore, when
deciding on a sampling method, one should
consider the purpose of the resulting data
(e.g., species list for the area, statistical
comparison among treatments) and the
resources available to accomplish the study
objectives. Although not discussed in detail in
this manual, post-sampling procedures are
equally important to consider and should
match the study objectives (see Klemm et al.
1990 and Charles et al. 2002).
Their small size and likelihood of drying make
headwater intermittent streams unique habitats
for sampling biota. Many methods developed
for perennial streams may not be as effective
or consistent in headwater streams. Water
depth and flow may not always be sufficient
for some sampling methods. In addition, the
sampling area may need to be reduced to
96
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minimize damage to streams and populations
(or even to logistically collect a sample). The
fluctuation of flow affects the wetted surface
area to a greater extent in headwater streams
than in larger, downstream water bodies.
Therefore, it is critical to monitor wetted
surface area when sampling biota. As streams
dry, surface water contracts and organisms
may track surface water and concentrate (e.g.,
Stanley et al. 1994). A density increase may
be misinterpreted as increased abundance in
response to drying if the context of wetted
surface area is ignored. Studies that compare
biological responses across time periods
and/or among habitat types should use
sampling methods that are equally efficient
across the range of associated hydrologic
conditions (Resh 1979, Boulton 1985)
The study objectives and the spatial
distribution of the fauna should determine the
number of samples or total sample area.
Where diversity (or richness) is of interest,
species-area relationships should be assessed
to determine the appropriate sample area.
Under ideal circumstances the number of
species collected over area sampled should
level-off. Therefore, the appropriate sampling
area should coincide with the asymptote
(where slope ~ 0) of the species-area curve.
However, because of the diverse and patchy,
but numerically skewed nature of aquatic
assemblages, the effort needed to reach the
asymptote is typically enormous and
logistically unattainable (Figure 4-1A). In
addition to the preponderance of rare taxa, the
limitations associated with fine-scale sample
stratification among habitat patches contribute
to the inability to attain the species-area
asymptote. An alternative goal that is more
feasible to achieve (in time and effort) is the
asymptote of the relationship between species
gained and sample area (Figure 4-IB). This
relationship measures the amount of
information gained per unit effort. Similar
considerations should be applied where
laboratory subsampling is done prior to
enumerating and identifying organisms
(Vinson and Hawkins 1996, Larsen and
Herlihy 1998). Because taxa richness
increases with the number of organisms
sampled, another consideration when
comparing among sites or treatments is to
standardize for the number of individuals or
rarefy the data (e.g., Downes et al. 1998,
McCabe and Gotelli 2000).
The following subsections are organized
according to biological group. Included are
more traditionally used communities of algae
and invertebrates as well as less commonly
used bryophytes and amphibians. Sampling
methods used in Headwater Intermittent
Streams Study are detailed. The identification
of indicators of flow permanence was the
objective of this study, sampling commonly
was restricted to the thalweg. This was done
because it was a consistent and conservative
target when comparing across sites with
varying hydrologic permanence and
ecological condition. Other spatial
configurations of field samples may be more
suitable depending upon the study objectives.
Alternative sampling methods are briefly
discussed at the end of each subsection.
97
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180 -|
140-
o>
.Q
E
1100-
o>
'
E
O 60
20
60 n
A.
0 2 4 6 8 10 ^2
Number of Samples
15-
0 2 4 6 8 10 12
Number of Samples
Figure 4-1 Examples of a species-area curve (A) and a species gained-area curve (B) for
benthic invertebrates samples (sample area = 0.053 m2) collected from a perennial site on
Falling Rock Branch, Robinson Forest, KY. Each point represents the mean (± 1 SE) of
100 permutations.
References
Biggs, B. J. F., R. J. Stevenson, and R. L.
Lowe. 1998. A habitat matrix conceptual
model for stream periphyton. Archivfur
Hydrobiologie 143:21-56.
Boulton, A. J. 1985. A sampling device that
quantitatively collects benthos in flowing or
standing waters. Hydrobiologia 127:31-39.
Charles, D. F., C. Knowles, and R. S. Davis
(editors) 2002. Protocols for the Analysis
of Algal Samples Collected as Part of the
United States Geological Survey National
Water Quality Assessment Program. Report
No. 02-06. The Academy of Natural
Sciences, Philadelphia, Pennsylvannia.
Charvet, S., A. Kosmala, and B. Statzner.
1998. Biomonitoring through biological
traits of benthic macroinvertebrates:
perspectives for a general tool in stream
management. Archivfur Hydrobiologie
142:415-432.
Downes, B. J., P. S. Lake, E. S. G. Schreiber,
and A. Glaister. 1998. Habitat structure and
regulation of local species diversity in a
stony, upland stream. Ecological
Monographs 68:237-257.
Hilsenhoff, W. L. 1987. An improved biotic
index of organic stream pollution. Great
Lakes Entomologist 20:31 -3 9.
98
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Klemm, D. I, P. A. Lewis, F. Fulk, and J. M.
Lazorchak. 1990. Macroinvertebrate Field
and Laboratory Methods for Evaluating the
Biological Integrity of Surface Waters.
EPA/600/4-90/030. U.S. Environmental
Protection Agency, Washington, D.C., USA.
Larsen, D. P. and A. T. Herlihy. 1998. The
dilemma of sampling streams for
macroinvertebrate richness. Journal of the
North American Benthological Society
17:359-366.
McCabe, D. J. and N. J. Gotelli. 2000.
Effects of disturbance frequency, intensity,
and area on assemblages of stream
macroinvertebrates. Oecologia 124:270-279.
Muotka, T. and R. Virtanen. 1995. The
stream as a habitat templet for bryophytes:
species' distributions along gradients in
disturbance and substratum heterogeneity.
Freshwater Biology 33:141-160.
Resh, V. H. 1979. Sampling variability and
life history features: basic considerations in
the design of aquatic insect studies. Journal
of the Fisheries Research Board Canada.
36:290-311.
Rosenberg, D. M. and V. H. Resh. 1993.
Freshwater Biomonitoring andBenthic
Macroinvertebrates. Chapman & Hall, NY.
Usseglio-Polatera, P., M. Bournaud, P.
Richoux, and H. Tachet. 2000. Biological
and ecological traits of benthic freshwater
macroinvertebrates: relationships and
definition of groups with similar traits.
Freshwater Biology 43:175-205.
Van Dam, H., A. Mertens, and J. Sinkeldam.
1994. A coded checklist and ecological
indicator values of freshwater diatoms from
the Netherlands. Netherlands Journal of
Aquatic Ecology 28:117-133.
Vinson, M. R. and C. P. Hawkins. 1996.
Effects of sampling area and subsampling
procedure on comparisons of taxa richness
among streams. Journal of the North
American Benthological Society 15:392-399.
Whitton, B. A. and M. G. Kelly. 1995. Use
of algae and other plants for monitoring
rivers. Australian Journal of Ecology 20:45-
56.
4.1 Sampling the bryophyte assemblage
General
This section describes sampling methods for
bryophyte assemblages (mosses and
liverworts) in headwater streams. Bryophytes
include the nonvascular, seedless plants
belonging to the classes Musci (mosses,
Figure 4-2) and Hepaticae (liverworts, Figure
4-3). Both groups share a life cycle composed
of two generations, the sporophyte (spore-
producing) and gametophyte (gamete-
producing). The sporophyte is directly
attached to and nutritionally dependent upon
the larger and longer-lived gametophyte.
Figure 4-2 Sporophyte and gametophyte
generations of a moss. (Photo by Michael
Luth)
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Figure 4-3 An epilithic moss (Musci) growing in a headwater stream.
Figure 4-4 An epilithic liverwort (Hepaticae) growing in a headwater stream.
100
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A fundamental gradient that governs the
spatial distribution of bryophytes is moisture
(Craw 1976, Glime and Vitt 1979).
Bryophytes range from being xerophytes
(adapted to dry habitats) to obligate
hydrophytes (requiring water). This range
enables mosses and liverworts to be
potentially useful indicators of headwater
stream hydrology. Additionally, because
bryophytes are sessile and relatively long-
lived compared to other stream-dwelling
organisms, their distributions may be useful
descriptors of hydrologic and ecological
conditions over several years. Bryophytes
have been used to monitor heavy metals and
other pollutants through accumulation in
tissue (e.g., Glime 1992, Engleman and
McDiffett 1996), biochemical change (Lopez
and Carbeilleira 1989), and species
composition (Vrhovsek et al. 1984,
Stephenson et al. 1995). Shifts in biomass,
species dominance and composition of
bryophyte assemblages have been linked to
changes in water chemistry (Omerod et al.
1987, Bowden et al. 1994), sediment particle
size (Vuori and Joensuu 1996), and hydrology
(Englund et al. 1997, Downes et al. 2003).
Texts for taxonomic identification of
bryophytes are referenced at the end of this
subsection.
Procedure
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg.
4.1.1 Qualitative sampling
Bryophyte sampling is confined to the thalweg
(deepest flow path) of the 30-m study reach.
Avoid unattached specimens or specimens
growing on loose woody debris because these
may have recently been deposited from
adjacent forest or upstream. Most specimens
suitable for sampling will be growing on stone
substrate and submerged in water (see Figures
4-2 and 4-3). In dry channels be careful to
select only specimens growing within the
thalweg. If no specimens are found note this
on the field form (Figure 4-4). Scrape small
samples (-10 cm2) of all representative
species seen in the thalweg using a scoopula
or similar tool. Collect specimens with the
sporophyte generation whenever possible,
because the sporophyte characteristics are
often critical for species-level identification.
Place all collected specimens from a study site
into a single 24-oz Whirl-Pak® bag with a
sample label that includes relevant
information (e.g., locality, date, collector's
initials). Keep samples cool (cooler with ice
bags or ice packs) while transporting them to
the laboratory and until the sample can be air-
dried in the laboratory. Protect samples from
ice meltwater. In the laboratory remove
samples and associated labels from Whirl-
Pak® bags and place them into paper bags or
envelopes for air drying. Write the label
information on the outside of the envelopes
with a permanent marker.
BRYOPHYTES SAMPLED: Y N
SAMPLE ID
COMMENTS
Figure 4-5 Appropriate location for recording bryophyte sample information on page 1 of
field forms.
4.1.2 Quantitative sampling
As opposed to the qualitative sampling
described above, quantitative sampling of
bryophyte assemblages requires field
identification (or at least recognition of
distinct taxa) and therefore some expertise on
101
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the flora (Slack 1984, Bowden et al. 2006).
There are three primary measures used to
quantify bryophytes by taxa: frequency,
percent cover, and standing crop. Frequency
measures the proportion of the samples
collected that contains a taxon, whereas
percent cover measures the proportion of the
sample area that is covered by a taxon.
Standing crop biomass is a measure of the
biomass of a taxon within a sample and is
usually reported as g m"2 dry weight or ash-
free dry mass. The advantage of the percent
cover and frequency over standing crop
biomass is that they are non-destructive,
enabling subsequent measurements. Percent
cover is usually considered an estimate and
therefore more subjective than frequency or
standing crop biomass. Some investigators
have subdivided the sample area using
plexiglass grids or other viewing devices to
improve repeatability. In addition, rather than
using absolute percentages, percentage
categories are often used when estimating
cover (e.g., Braun-Blanquet cover scale, see
Mueller-Dombois and Ellenberg 1974,
Bowden et al. 2006). Voucher specimens for
each taxon are collected for later identification
or confirmation in the laboratory.
Three sampling methods commonly used and
are listed by increasing level of effort: point-
intercept, transect, and quadrat (i.e., plot).
Points, transects and quadrats should be
randomly or haphazardly selected across the
study reach to avoid samples biasing their
reach representation. Depending upon the
study objectives, it may be useful to stratify
sampling within a study reach (e.g., habitat
type, stream margin versus mid-channel,
height relative to water surface). Stratified
sampling (or floristic habitat sampling) has
been advocated when compiling
comprehensive surveys of bryophyte diversity
(Newmaster et al. 2005). The point-intercept
method uses a grid or coordinate system.
Each randomly or haphazardly selected
coordinate (point) is sampled by simply
recording the species present at the point, on
the nearest substrate (e.g., cobble), or a
surrounding area (making it similar to quadrat
sampling). The transect method uses
randomly or haphazardly placed transects
(measuring tape or string) typically positioned
perpendicular to the direction of flow.
Sampling along transects may span only the
wetted width, entire active, or into the
adjacent riparian zone. Percent cover for each
species is determined by the percent of the
transect length that is intercepted by each
species. Frequency may be assessed among
transects or within transects by recording
individual species-patches along each
transects. Some investigators treat transects as
belts, where bryophytes are sampled within a
set distance (e.g., 0.1 m) upstream and
downstream of each transect (e.g., Steinman
and Boston 1993, Suren and Duncan 1999).
The quadrat method uses circular, square, or
rectangular plots of known area that are
randomly or haphazardly positioned in the
study area. The percent cover of each species
within the quadrat is recorded and frequency
is typically assessed across replicate quadrats.
Some investigators have sampled quadrats
along transects to quantify assemblage shifts
across geomorphic units (e.g., Jonsson 1996).
Potential edge effects (perimeter:area) are
lower for circular plots than for square or
rectangular plots (Krebs 1999). The number
and size of replicate sampling units depends
on the patchiness of bryophytes with the study
reaches and the resources available for the
study. Studies comparing transect, point
intercept, and quadrat methods have generally
found similar estimates of bryophyte and
macroalgae abundance (Rout and Gaur 1990,
Neechi et al. 1995). However, because the
quadrat method usually covers a larger
sampling area, this method will include more
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rare taxa and tend to have higher estimates of
taxa richness.
References
Bowden, W.B., J. C. Finlay, and P. E.
Maloney. 1994. Long-term effects of PC>4
fertilization on the distribution of
bryophytes in an arctic river. Freshwater
Biology 32:445-454.
Bowden, W.B., J. M. Glime, and T. Riis.
2006. Macrophytes and bryophytes. Pages
381-414 in: F. R. Hauer and and G. A.
Lamberti (editors), Methods in Stream
Ecology, 2nd edition. Academic Press, San
Diego, California.
Craw, R. C. 1976. Streamside bryophyte
zonations. New Zealand Journal of Botany
14:19-28.
Downes, B. J., T. J. Entwisle, and P. Reich.
2003. Effects of flow regulation on
disturbance frequencies and in-channel
bryophytes and macroalgae in some upland
streams. River Research Applications 19:27-
42.
Engleman, C. J. and W. F. McDiffett. 1996.
Accumulation of aluminum and iron by
bryophytes in streams affected by acid-mine
drainage. Environmental Pollution 94:67-74.
Englund, G., E.G. Jonsson, and B. Malmqvist.
1997. Effects of flow regulation on
bryophytes in north Swedish rivers.
Biological Conservation 79:79-86.
Glime, J. M. 1992. Effects of pollutants on
aquatic species. Pages 333-361 in: J. W.
Bates and A. M. Farmer (editors),
Bryophytes and Lichens in a Changing
Environment. Clarendon Press, Oxford,
United Kingdom.
Glime, J.M. and D.H.Vitt. 1979. The
physiological adaptations of aquatic Musci.
Lindbergia 10:41-52.
Jonsson, B. G. 1996. Riparian bryophyte
vegetation in the Cascade mountain range,
northwest U.S.A.: patterns at different
spatial scales. Canadian Journal of Botany
75:744-761.
Krebs, C. J. 1999. Ecological Methodology,
2nd edition, Benjamin/Cummings, Menlo
Park, California.
Lopez, J. and A. Carballeria. 1989. A
comparative study of pigment contents and
responses to stress in five species of aquatic
bryophytes. Lindbergia 15:188-194.
Mueller-Dombois, D. and H. Ellenberg. 1974.
Aims and Methods of Vegetative Ecology.
Wiley-Interscience, New York.
Necchi, O., L. H. Z. Branco, C. C. Z. Branco.
1995. Comparison of three techniques for
estimating periphyton abundance in bedrock
streams. Archivfur Hydrobiologie 134:393-
402.
Newmaster, S. G., R. J. Belland, A. Arsenault,
D. H. Vitt, and T. R. Stephens. The ones we
left behind: comparing plot sampling and
floristic habitat sampling for estimating
bryophyte diversity. Diversity and
Distributions 11:57-72.
Ormerod, S. J., K. R. Wade, and A. S. Gee.
1987. Macro-floral assemblages in upland
Welsh streams in relation to acidity, and
their importance to invertebrates.
Freshwater Biology 18:545-557.
Rout, J. and J. P. Gaur. 1990. Comparative
assessment of line transect and point
intercept methods for stream periphyton.
Archivfur Hydrobiologie 119:293-298.
103
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Slack, N. G. 1984. A new look at bryophyte
community analysis: field and statistical
methods. Journal of the Hattori Botanical
Laboratory 55:113-132.
Steinman, A. D. and H. L. Boston. 1993. The
ecological role of aquatic bryophytes in a
woodland stream. Journal of the North
American Benthological Society 12:17-26.
Stephenson, S. L., S. Moyle Studlar, C. J.
McQuattie, and P. J. Edwards. 1995.
Effects of acidification on bryophyte
communities in West Virginia mountain
streams. Journal of Environmental Quality
24:116-125.
Suren, A. S. andM. J. Duncan. 1999. Rolling
stones and mosses: effect of substrate
stability on bryophyte communities in
streams. Journal of the North American
Benthological Society 18:457-467.
Vrhovsek, D., A. Martincic, and M. Kralj.
1984. The applicability of some numerical
methods and the evaluation of Bryophyta
indicators species for the comparison of the
degree of pollution between two rivers.
Archivfur Hydrobiologie 100:431 -444.
Vuori, K.M. and I. Joensuu. 1996. Impact of
forest drainage on the macroinvertebrates of
a small boreal headwater stream: do buffer
zones protect lotic biodiversity? Biological
Conservation 77:87-95.
Taxonomic Texts
Breen, R. S. 1963. Mosses of Florida, an
Illustrated Manual. University of Florida
Press, Gainesville, Florida.
Churchill, S. P. 1985. A synopsis of the
Kansas mosses with keys and distribution
maps. University of Kansas Science Bulletin
53:1-64.
Conard, H.S. and P. L. Redfearn, Jr. 1979.
How to Know the Mosses and Liverworts,
2nd edition. WCB McGraw-Hill, Boston,
Massachusetts.
Crum, H. 1983. Mosses of the Great Lakes
Forest, 3rd edition. University of Michigan
Press, Ann Arbor, Michigan.
Crum, H. 1991. Liverworts andHornworts of
South Michigan. University of Michigan
Press, Ann Arbor, Michigan.
Crum, H. and L. E. Anderson. 1981. Mosses
of Eastern North America. Columbia
University Press, New York.
Flowers, S. and A. Holmgren. 1973. Mosses:
Utah and the West. Brigham Young
University Press, Provo, Utah.
Hicks, M. L. 1992. Guide to the Liverworts
of North Carolina. Duke University Press,
Durham, North Carolina.
Ireland, R. R. and G. Bellolio-Trucco. 1987.
Illustrated Guide to Some Hornw or ts,
Liverworts and Mosses of Eastern Canada.
Syllogeus 62, National Museum of Canada,
Ottawa.
Jennings, O. E. 1951. A Manual of the
Mosses of Western Pennsylvania and
Adjacent Regions, 2nd edition. Notre Dame
University Press, South Bend, Indiana.
Lawton, E. 1965. Keys for the Identification
of Mosses of Washington and Oregon.
American Bryological and Lichenological
Society.
104
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Lawton, E. 1971. Moss Flora of the Pacific
Northwest. Journal of the Hattori Botanical
Laboratory, Supplement No. 1, Hattori
Botanical Laboratory, Nichinan, Japan.
Lawton, E. 1971. Keys for the Identification
of the Mosses of the Pacific Northwest.
Journal of the Hattori Botanical Laboratory,
Supplement No. 2, Hattori Botanical
Laboratory, Nichinan, Japan.
Malcolm, B. andN. Malcolm. 2000. Mosses
and Other Bryophytes : An Illustrated
Glossary. Micro-Optic Press, Nelson, New
Zealand.
Redfearn, P. L. Jr. 1983. Mosses of the
Interior Highlands of North America.
Missouri Botanical Garden Press, St. Louis,
Missouri.
Reese, W.D. 1984. Mosses of the Gulf South:
From the Rio Grande to the Apalachicola.
Louisiana State University Press, Baton
Rouge, Louisiana.
Schofield, W.B. 2002. Field guide to
Liverwort Genera of Pacific North America.
Global Forest Society and University of
Washington Press, San Francisco,
California.
Schuster, R. M. 1966-1992. The Hepaticae
and Anthocerotae of North America East of
the Hundredth Meridian, 6 volumes. New
York, London and Chicago.
Smith, H. L. 1966. Mosses of the Great
Plains and Arkansas River lowlands of
Kansas. University of Kansas Science
Bulletin 46: 433-474.
Vitt, D. H., J. E. Marsh, and R. B. Bovey.
1993. Mosses, Lichens, and Ferns of
Northwest North America. Lone Pine
Publishing, Auburn, WA.
Weber, W.A. 1973. Guide to the Mosses of
Colorado. Occasional Paper No. 6, Institute
of Arctic and Alpine Research, University of
Colorado, Boulder, Colorado.
Welch, W.H. 1957. Mosses of Indiana.
Purdue University Press (Reissue edition),
Lafayette, Indiana.
Equipment and supplies
Measuring tape (50 m)
Metal scoopula or spatula
24 oz Whirl-Pak® bags
Pencils
Permanent marker
Label paper
Field forms
Cooler
Ice or ice packs
Paper bags or envelopes
4.2 Sampling the epilithic algal assemblage
General
This subsection describes methods for
sampling the algal assemblage in headwater
streams. The particular algae sampled in these
procedures are epilithic algae (or algae
associated with stone surfaces) and includes
diatoms (Bacillariophyta) and "soft" algae
(i.e., Chlorophyta, Cyanophyta, Rhodophyta,
and Chrysophyta). Epilithic algae are
associated with fungi, bacteria, heterotrophic
protists, and organic matter and together they
form a matrix called periphyton, biofilm, or
aufwuchs. The target organisms for
laboratory identification are the algae within
the periphyton, but because algae are difficult
to exclusively collect, the periphyton is
sampled. Algal assemblages have been shown
to be useful indicators of ecological condition
in wadable streams (e.g., Pan et al. 1996, Hill
105
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et al. 2000). The ubiquity, diversity, sampling
efficiency, and responsiveness to physical and
chemical stressors are all attributes for the use
of algae in bioassessment (Patrick 1973,
Stevenson and Lowe 1986). Despite being
ubiquitous, algae have received less attention
than invertebrates in temporary streams
research (see review by Stanley et al. 2004).
Algae are potentially useful indicators of
hydrologic permanence because algae inhabit
a wide range of habitats (terrestrial to aquatic)
and varying in desiccation tolerance and
presence of resistant structures (e.g., akinetes,
cysts, zygotes, mucilage) among taxonomic
groups (Davis 1972).
4.2.1.Quantitative sampling of epilithic algae
This first procedure is modified from the
procedure described by Hill (1998) and
focuses only on the collection of periphyton
on natural substrates to determine the
taxonomic composition of the algal
assemblage (by abundance and biovolume).
The algae assemblage is sampled during each
season (spring and summer) because it is
likely to vary with season.
Procedure
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg. Identify erosional and depositional
habitats with the study reach. Two separate
algae samples are taken from each study
reach, one from each habitat type (erosional
and depositional). Sampling is confined to the
thalweg of the study reach and is done
regardless of hydrologic condition. Each
sample is a composite of 12 cm2 areas from
upper surface of 6 individual stones.
4.2.1.1. Substrate collection
Begin by haphazardly collecting 6 stones (>12
cm2 upper surface area) from the thalweg of a
habitat type and placing them in the large
basin with the upper surface facing upward.
Avoid disturbing the streambed as much as
possible when collecting stones and make sure
that the stones have not been disturbed by
other sampling activities (communicate with
fellow crewmembers). Spread the sampling
across multiple units of each habitat type
along the study reach. However, where
hydrologic conditions vary among units of (or
stones from) a habitat type in a study reach
(i.e., there are pools with and without surface
water), restrict sampling to the dominant
hydrologic condition represented by the
habitat units within the study reach. For
example, if a study reach has 5 depositional
habitat units and 4 had surface water and one
was dry, collect the 6 stones from the 4 wet
units. Indicate on the field form the number
of stones collected and whether the stones
collected were wet or dry (Figure 4-5). Stones
can be randomly selected within available
habitat units in the reach. Numbers ranging
from 1 to 100 can be drawn and the nearest
stone along the thalweg coinciding
proportionally with the unit length is selected.
# STONES SAMPLED: Depositional I
SAMPLE ID
Four - JC - 1 - D
Four - JC - 1 - E
ALGAE
labitats 6 Erosional Habitats 6 TOTAL AREA SAMPLED: 144 cm2 (each 12 cm2)
HABITAT TYPE
Depositional
Erosional
NUMBER OF
BAGS
lof 1
lof 1
COLLECTED
BY:
KMF
KMF
COMMENTS
Wet
Wet
Figure 4-6 Appropriate location for recording algal sample information on page 1 of field
forms. Example information is shown in red.
106
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4.2.1.2. Compositing and preserving sample
Fill the wash bottle to the 50-ml mark with
stream water. Place a 1.5" diameter PVC
circle (i.e., delimiter) on the upper surface of a
stone to define a 12 cm2 area. Use the metal
spatula and a firm-bristle toothbrush (trim
bristles to half original length) to dislodge
algae from the stone surface within the
delimiter (Figure 4-6). Rigorously scrape and
brush the surface for 30 seconds. Be aware
that because clay particles have similar
density (mass per unit volume), excess clay
particles in the sample may hamper
identification and enumeration of algae.
Using the wash bottle, sparingly wash the
dislodged algae from the delimited area into
the small plastic container. Repeat this step
with the 5 remaining stones to make a
composite sample. Use the remaining water
in the wash bottle to rinse off tooth brush and
spatula into the small basin. Pour the
composite sample from the small container
through the small funnel and into a 50-ml
centrifuge tube. Use a syringe or bulb pipette
to add 2 ml of 10% formalin to preserve the
sample*.
If formalin is not taken to the field, keep
sample in the dark and on ice until it is
preserved. Tightly cap the centrifuge tube and
seal with electrical tape. Gently shake the
tube to distribute the formalin throughout the
sample. Make a label (waterproof paper and
pencil) that includes relevant information
(e.g., locality, habitat type, date, collector's
initials). Attach the sample label securely to
the outside of the centrifuge tube using
packing tape or clear tape strips. Also write
* Wear gloves and safety eyeglasses when using
formalin and work in a well-ventilated area. Formalin
is extremely caustic and potentially carcinogenic. It
may cause severe 'irritation on contact with skin and
eyes. Rinse immediately with water in case of contact
with skin or eyes.
label information on the field form (Figure 4-
5). Thoroughly rinse all sampling equipment
with clean stream water to prevent cross-
contamination between samples. If additional
measures of biomass and/or pigment are
needed, the sample can be split volumetrically
(see Hill 1998).
Repeat procedures outlined in 4.2.1.1 and
4.2.1.2 for the remaining habitat type.
Figure 4-7 Collecting epilithic algae from a
stone within the sample delimiter.
4.2.1.3. Sample transport and shipping
Before leaving a site, check that all samples
are labeled properly and safely stowed. At the
vehicle, consolidate all algae samples in one
location. All the samples can then be placed
in a crush-resistant and leak-proof container
and transported to the laboratory for
processing. If samples need to be shipped,
include any special shipping forms that may
be required for the formalin-preserved
samples.
4.2.2 Alternative methods
Here we briefly describe various techniques
used to sample epilithic algae in streams.
More details are available in reviews by
Stevenson and Lowe (1986) and Aloi (1990).
Algal sampling methods can be separated into
107
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two broad categories: natural substrates and
artificial substrates. As previously described,
natural substrate sampling involves
quantitatively collecting epilithon found
growing naturally on substrates in streams. In
contrast, artificial substrate sampling involves
placing substrates (e.g., glass slides, ceramic
tiles, bricks) into streams for periphyton
colonization. Because the exposure time is
known, artificial substrates provide the
investigator with more control and perhaps
less variability among sampling units than
natural substrates. However, assemblages
colonizing artificial substrates may not
provide a realistic characterization of the algal
assemblage in streams. Taxonomic
composition and measures of biomass
(chlorophyll a and AFDM) on artificial
substrates can differ from these algal measures
on natural substrates (Lay and Ward 1987,
Cattaneo and Amireault 1992). Homogeneous
substrate texture and short incubation times of
artificial substrate procedures have been
identified as the likely causes for lower algal
biomass and lower representation by green
and blue-green algae than seen in adjacent
natural substrates. Both methods have
benefits and drawbacks for monitoring algal
assemblages and these should be weighed
carefully when designing monitoring studies.
Algae in headwater streams are often
logistically easier to collect than from deeper
rivers and lakes. Many substrate types are
easily removed from the stream for
subsequent collection of algae. Rather than
sub sampling periphyton on a substrate
particle, several investigators have used the
entire substrate as a sampling unit (e.g., Biggs
and Close 1989, Dodds et al. 1999, Mosisch
2001). Surface area of substrates can be
estimated using substrate dimensions and
geometric equations (e.g., Graham et al.
1988). Others have determined stone surface
area by covering stones with aluminum foil,
plastic wrap, or ink stamps (Doeg and Lake
1981, Lay and Ward 1987). Surface area -
weight relationships are then used to
determine surface area of substrates. Large
boulders and bedrock common to steep
headwater streams can pose a problem in
retrieving samples and quantifying surface
area. Syringe type samplers (e.g., Loeb 1981,
Flower 1981, Peters et al. 2005) offer a
solution, where a sample can be collected in
situ. Syringe samplers use brushes to remove
attached periphyton within an enclosed area;
then the sample is suctioned and transferred
onto a filter or into a sample container. One
drawback noted about syringe samplers is an
underestimation of chlorophyll a
concentrations from stream samples but not
from lake samples (Cattaneo and Roberge
1991). Sampler brushes are likely ineffective
at removing tightly attached members of the
periphyton assemblage in streams. Davies and
Gee (1993) developed a scouring disc for
periphyton removal and reported higher
concentrations of chlorophyll a were obtained
from the scouring disc than from either
brushing or scraping. Algae associated with
fine particles (epipsammon and epipelon) can
be sampled simply by using an area delimiter
(e.g., inverted petri dish) and spatula or
trowel. The delimiter is positioned into the
upper sediment layers and the spatula is
positioned beneath. The spatula is carefully
lifted and the sample is transferred to a sample
container using a funnel. Because algae may
be firmly attached to sand grains, additional
laboratory steps (i.e., sonication) are needed
prior to microscopic or fluorometric
measurement (Miller et al. 1987, Romani and
Sabater 2001). Further details on sampling
algae from various substrates are discussed in
Moulton et al. 2002.
References
108
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Aloi, J. E. 1990. A critical review of recent
freshwater periphyton field methods.
Canadian Journal of Fisheries and Aquatic
Sciences 47:656-670.
Biggs, B. J. F. and M. E. Close. 1989.
Periphyton biomass dynamics in gravel bed
rivers: the relative effects of flows and
nutrients. Freshwater Biology 22:209-231.
Cattaneo, A. and M. C. Amireault. 1992.
How artificial are artificial substrata for
periphyton. Journal of the North American
Benthological Society 11:244-256.
Cattaneo, A. and G. Roberge. 1991.
Efficiency of a brush sampler to measure
periphyton in streams and lakes. Canadian
Journal of Fisheries and Aquatic Sciences
48:1877-1881.
Davies, A. L. and J. H. R. Gee. 1993. A
simple periphyton sampler for algal biomass
estimates in streams. Freshwater Biology
30:47-51.
Davis, J. S. 1972. Survival records in the
algae, and the survival role of certain algal
pigments, fat, and mucilaginous substances.
The Biologist 54:52-93.
Dodds, W. K., R. E. Hutson, A. C. Eichem,
M. A. Evans, D. A. Gudder, K. M. Fritz, and
L. Gray. 1996. The relationship of floods,
drying, flow and light to primary production
and producer biomass in a prairie stream.
Hydrobiologia 333:151-159.
Doeg, T. and P. S. Lake. 1981. A technique
for assessing the composition and density of
the macroinvertebrate fauna of large stones
in streams. Hydrobiologia 80:3-6.
Flower, R. J. 1985. An improved epilithon
sampler and its evaluation in two acid lakes.
British Phycological Journal 20:109-115.
Graham, A. A., D. J. McCaughan, F. S.
McKee. 1988. Measurement of surface
area of stones. Hydrobiologia 157:85-87.
Hill, B.H. 1998. Periphyton. Pages 119-132
in: J.M. Lazorchak, DJ. Klemm, and D.V.
Peck (editors), Environmental Monitoring
and Assessment Program - Surface Waters:
Field Operations and Methods for
Measuring the Ecological Condition of
Wadeable Streams. EPA/620/R-94/004F.
U.S. Environmental Protection Agency,
Washington, D.C.
Hill, B. H., A. T. Herlihy, P. R. Kaufmann, R.
J. Stevenson, F. H. McCormick, and C. B.
Johnson. 2000. Use of periphyton
assemblage data as an index of biotic
integrity. Journal of the North American
Benthological Society 19:50-67.
Lay, J. A. and A. K. Ward. 1987. Algal
community dynamics in two streams
associated with different geological regions
in the southeastern United States. Archivfur
Hydrobiologie 108:305-324.
Loeb, S. L. 1981. An in situ method for
measuring the primary productivity and
standing crop of the epilithic periphyton
community in lentic systems. Limnology and
Oceanography 26:394-399.
Miller, A. R., R. L. Lowe, and J. T.
Rotenberry. 1987. Succession of diatom
communities on sand grains. Journal of
Ecology 75:693-709.
Mosisch, T. 2001. Effects of desiccation on
stream epilithic algae. New Zealand Journal
109
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of Marine and Freshwater Research 35:173-
179.
Moulton, S. R. II, J. G. Kennen, R. M.
Goldstein, and J. A. Hambrook. 2002.
Revised Protocols for Sampling Algal,
Invertebrate, and Fish Communities As Part
of the National Water-Quality Assessment
Program. Department of Interior, U.S.
Geological Survey Open-File Report 02-
150, 75 p.
Pan, Y., R. J. Stevenson, B. H. Hill, A. T.
Herlihy, and G. B. Collins. 1996. Using
diatoms as indicators of ecological condition
in lotic systems: a regional assessment.
Journal of the North American
Benthological Society 15:481 -495.
Patrick, R. 1973. Use of algae, especially
diatoms, in assessment of water quality.
Pages 76-95 in: J. Cairns and K. L. Dickson
(editors). Biological Methods for the
Assessment of Water Quality. ASTM
Special Technical Publication 528,
American Society for Testing and Materials,
Philadephia, PA.
Peters, L., N. Scheifhacken, M. Kahlert, and
K. O. Rothhaupt. 2005. An in efficient in
situ method for sampling periphyton in lakes
and streams. Archivfur Hydrobiologie
163:133-141.
Romani, A. M. and S. Sabater. 2001.
Structure and activity of rock and sand
biofilms in a Mediterranean stream. Ecology
82:3232-3245.
Stanley, E. H., S. G. Fisher, and J. B. Jones.
2004. Effects of water loss on primary
production: a landscape-scale model.
Aquatic Sciences 66:130-138.
Stevenson, R. J. and R. L. Lowe. 1986.
Sampling and interpretation of algal patterns
for water quality assessments. Pages 118-
149 in: E. G. Isom (editor), Rationale for
Sampling and Interpretation of Ecological
Data in the Assessment of Freshwater
Ecosystems. ASTM Special Technical
Publication 894, American Society for
Testing and Materials, Philadelphia,
Pennsylvania.
110
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Figure 4-8 Equipment used to collect and preserve algal assemblage samples. Numbers
correspond to the Equipment and Supplies list.
Equipment and supplies (Numbers correspond
to items in Figure 4-7)
Measuring tape (50 m)
1. Plastic wash basin (approximately 35 cm x
29 cm x 14 cm)
2. Small plastic container or basin (e.g.,
Tupperware® or Rubbermaid® container)
(approximately 25 cm x 16 cm x 6 cm,
large enough to contain a cobble and large
gravel particle and can be used to store
items listed below)
3. PVC ring delimiter (1.5 in or 3.8 cm
diameter pipe cut 2 to 3 cm in length)
4. Firm-bristle toothbrush (2) - trim bristles to
half their original length
5. Spatula or scoopula
6. Water squirt bottle (with 50 ml volume
marked)
7. Buffered formalin (10%)
8. Small syringe or bulb pipette
9. 50 ml centrifuge tubes
10. Small funnel
11. Electric tape
12. Label paper
13. Pencils
14. Packing tape or clear tape strips
4.3 Visual and tactile assessment of algal
cover
General
This subsection provides instructions for
rapidly assessing algal cover in headwater
streams. The method uses a categorical index,
Algal Cover Index (ACT) that is based on
visual and tactile characteristics of periphyton
(and associated algae). The ACT scores and
associated characteristics are shown in Table
111
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1. The ACT has been field tested and ACT
scores have explained 68-85% of the variation
in measured levels of algal (chlorophyll a) and
periphyton (AFDM) biomass in streams
(Feminella and Hawkins 2000). The protocol
described here is modified from Hawkins et
al. (2001). Another field-based rapid
periphyton method that separately
characterizes macroalgal and microalgal cover
is described in Stevenson and Bahls (1999).
EMAP protocols include percent classes for
filamentous algae (Kaufmann and Robison
1998). The NAQWA qualitative algae
sampling protocol includes designating the
abundance classes (dense to none) for
periphyton at a site (Moulton et al. 2002). The
ACT is measured during each season (spring
and summer) because it is likely to vary with
season.
Table 4-1 Algal Cover Index (ACI) scores and their associated characteristics
ACI Score Visual and Tactile Characteristics
1 substrate is rough with no apparent growth
1.5 substrate is slimy, but biofilm not visible (i.e., tracks from scratching rock
with back of fingernail is not visible)
2 thin layer visible (0.5-1 mm thick, i.e., tracks from scratching rock with
back of fingernail is visible)
3 algal mat thickness ranges from 1-5 mm thick and filamentous algae is rare
4 algal mat thickness ranges from 5-20 mm thick and filamentous algae
common
5 al§al mai thickness >2 cm and/or filamentous algae dominates
Procedure
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg. Algal cover assessment is based on
> 25 substrate particles on the streambed
surface. Particles assessed should be spread
along the thalweg of the entire study reach.
Final ACI score for the study reach is based
on the dominant score of the assessed
particles. Where there is a clear discrepancy
between habitat types (i.e., deposit!onal and
erosional), note ACI scores for both habitat
types. Algal cover can be assessed while
sampling benthic invertebrates (assessing
surface cobble and gravel while scrubbing
attached invertebrates). Photographic
examples of ACI scores are shown in Figure
4-8. The ACI score is circled on the field
form (Figure 4-9). The ACI scores and
associated characteristics are also listed on the
bottom of page 3 of the field forms.
112
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Figure 4-9 Categorical examples of algal cover based on visual and tactile characteristics.
Numbers represent Algal Cover Index (ACI) scores associated with periphyton on stones in
the photographs.
PRESENCE OF
HEADCUT IN REACH
Y N
ALGAL COVER INDEX
1 I1'2
2
345
# CORES FOR SUBSTRATE MOISTURE
(depositional)
Figure 4-10 Appropriate location for recording the dominant reach score for the Algal
Cover Index on page 1 of field forms.
113
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References
Feminella, J.W. and C.P. Hawkins. 2000.
Eyeing the aufwuchs and groping the
growth: a rapid method for estimating
epilithic periphyton abundance in streams.
Bulletin of the North American
Benthological Society 17: 180.
Hawkins, C., J. Ostermiller, M. Vinson, and
R.J.Stevenson. 2001. Stream algae,
invertebrate, and environmental sampling
associated with biological water quality
assessment field protocols. Unpublished
document, Utah State University.
Kaufmann, P. R. and E. G. Robison. 1998.
Physical habitat characterization. Pages 77-
118 in J.M. Lazorchak, DJ. Klemm, and
D.V. Peck (editors). Environmental
Monitoring and Assessment Program -
Surface Waters: Field Operations and
Methods for Measuring the Ecological
Condition of Wadeable Streams.
EPA/620/R-94/004F. U.S. Environmental
Protection Agency, Washington, D.C.
Moulton, S. R. II, J. G. Kennen, R. M.
Goldstein, and J. A. Hambrook. 2002.
Revised Protocols for Sampling Algal,
Invertebrate, and Fish Communities As Part
of the National Water-Quality Assessment
Program. Department of Interior, U.S.
Geological Survey Open-File Report 02-
150, 75 p.
Stevenson, R. J. and L. L. Bahls. 1999.
Periphyton protocols. Pages 6.1 -6.22 in. M.
T. Barbour, J. Gerrisen, B. D. Snyder, and J.
B. Stribling (editors). Rapid bioassessment
protocols for use in streams andwadeable
rivers: periphyton, benthic
macroinvertebrtes and fish. 2nd edition,
EPA/841/B/98-010. Office of Water, U.S.
Environmental Protection Agency,
Washington, D.C.
Equipment and supplies
Measuring tape (50 m)
Field forms
4.4 Sampling the benthic invertebrate
assemblage
General
This subsection provides methods for
quantitatively sampling the benthic
invertebrate assemblage in headwater streams.
Benthic invertebrate surveys are widely used
to evaluate the condition or health of water
bodies (Hellawell 1986, Rosenberg and Resh
1993, Rader et al. 2001). Invertebrate
assemblages are composed of a wide range of
taxonomic and functional groups, many of
which can be found in headwater streams.
Furthermore, a diversity of life histories (e.g.,
voltinism, cohort production interval,
dormancy stages) and physiological tolerances
are found among aquatic invertebrates
(Williams 1996, Frouz et al. 2003). Habitat
characteristics (e.g., predictability, disturbance
intensity, productivity) set the template
governing the evolution of life histories and
therefore the composition of assemblages
(Southwood 1977, Townsend and Hildrew
1994). Flow is considered one of the ultimate
drivers of lotic systems (Lytle and Poff 2004),
and may be even more critical to temporary
water bodies (Walker et al. 1995, Schwartz
and Jenkins 2000). Thus, the composition of
invertebrate assemblages should reflect the
flow permanence in headwater streams.
However, among past investigations there is
no consensus regarding the distinctiveness of
invertebrate communities among stream
reaches of different flow permanence (Deluchi
1988, Feminella 1996, Dietrich and Anderson
2000, Fritz and Dodds 2002, Price et al.
2004). As is often the case in ecological
systems, this disparity suggests that the
relationship between flow permanence and
assemblage organization may be complex.
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Plasticity of life histories, subtle variation of
drying intensity, degree of connectivity to
refugia, physiographic variation,
anthropogenic impacts, and other factors may
influence assemblage structure.
4.4.1 Quantitative bucket sampling for
invertebrate assemblage
The surface water conditions of many
headwater streams fluctuate from continuous
flow to only standing water in pools to
complete absence. Therefore, the method
used to collect benthic invertebrates needs to
be effective and consistent across the range of
hydrological conditions seen in headwater
streams. Many existing sampling methods
take advantage of flowing conditions to trap
invertebrates in nets positioned downstream of
the sampling area (e.g., Surber sampler, kick
net); however, flow in headwater streams is
often too low (sometimes absent) to
effectively use these methods. The
quantitative bucket sampling method
described here is: 1) not dependent upon flow
conditions, 2) performed by a single operator,
3) light weight, and 4) inexpensive. The
bucket sampling method is modified from
methods described by Wilding (1940) and
Statzner (1981). The sample area of a 5
gallon bucket sampler is 0.053 m2 (26-cm
diameter). A smaller sample area (e.g., coffee
can) may be required to collect benthos from
step-pool streams dominated by boulders and
large woody debris. The benthic invertebrate
assemblage is sampled during each season
(spring and summer) because it is likely to
vary with season.
Procedure
Before hiking to the study reach(es) make sure
all the equipment is stowed in the backpacks
and there are ample Whirl-pak bags and
ethanol. Transfer the 95% ethanol (usually
from 5 gal. container) into transport jug(s)
using a funnel. Typically 1 to 1.5 liters is
sufficient to preserve all the samples taken
from a study reach.
Delineate the 30-m study reach so that the
measuring tape is positioned along the
thalweg. Identify erosional and depositional
habitats within the study reach (see Subsection
3.4 for designating habitat units). Preliminary
data from perennial and intermittent
headwater streams in Kentucky, Indiana, and
Ohio indicate that the number of additional
taxa collected within a 30-m reaches an
asymptote after 8 samples (0.42 m2, see Figure
4. IB). We recommend that investigators
independently assess species-area curves for
study reaches, particularly if estimation of
invertebrate diversity (a common metric used
in ecological condition assessments) is an
objective.
Eight separate invertebrate samples (each
0.053 m2) are haphazardly taken from each
study reach, 4 from each habitat type
(erosional and depositional). Samples are kept
separate to 1) determine the sampling effort
needed to sufficiently represent the
invertebrate assemblage and 2) provide
within-reach measures of variance. Where
these objectives are not a concern, samples
may be composited. Sampling is confined to
the thalweg of the study reach and only where
surface water is present (see Subsection 3.1
for designating hydrologic condition). For
instance, if there is continuous surface flow
throughout the study reach, 4 samples are
taken from each habitat type. If there is
surface water only in depositional units, only
4 depositional samples are taken. Samples
should be spread across multiple units of each
habitat type (i.e., erosional and depositional)
along the study reach. Where the study reach
has < 1 habitat unit (e.g., a pool) with surface
water and the other units of that type are dry,
do not sample that habitat type. Do not
sample areas where the water depth exceeds
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the height of the bucket sampler. Samples can
be randomly positioned within available
habitat units in the reach. For example,
numbers ranging from 1 to 100 can be drawn
and the center of the bucket sampler is
positioned along the thalweg coinciding
proportionally along the unit length.
4.4.1.1. Sample collection and preservation
Attach the canvas skirt to the bucket sampler
by sliding the elastic band of the canvas skirt
over and around the bottom edge of the bucket
sampler (Figure 4-11). A foam ring, rather
than the canvas skirt, should be fitted to the
bucket where the streambed is bedrock.
Sample collection should proceed in an
Figure 4-11 Photographs of bucket
sampler and canvas skirt A) unassembled
and B) assembled for sampling.
upstream direction. Avoid disturbing the
streambed outside of the sampling area as
much as possible by walking along the banks
rather than in the thalweg. Make sure that the
areas sampled have not been disturbed by
other sampling activities. Begin by
identifying a sampling location within the
channel thalweg and place the wash basin,
sieve, hand net, and trowel on a bank or gravel
bar near the sampling location. With both
hands lift the weighted edge of the canvas
skirt above the bottom edge of the sampling
bucket (to prevent the skirt from being inside
the sampling area once the bucket encloses the
sampling area). Push the bottom edge of the
bucket 3 to 5 cm vertically into the streambed.
Adjust the weighted edge of the skirt to seal
the sampling area. By hand remove the coarse
surface substrates (i.e., large gravel and
cobble) from the enclosed sampling area and
place them into the wash basin or sieve to be
scrubbed (Figure 4-12). Stir by hand or trowel
the remaining substrate within the bucket for
10 seconds to a depth of 10 cm (or bedrock,
whichever is shallower). This step helps to
suspend invertebrates from streambed
interstices into the water column.
Figure 4-12 Coarse surface substrate set
aside in basin for scrubbing.
Immediately sweep the hand net through the
water column for 10 seconds to capture the
suspended invertebrates (Figures 4-12 and 4-
13). Repeat the substrate stirring and net
sweeping steps 2 more times. After the
sweeping is completed, empty the net contents
into the wash basin. Look for any
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Figure 4-13 Sweep the hand net through
the water column to collect suspended
invertebrates within the bucket area.
invertebrates that may be attached to the net
and put these in the basin or sieve. (If the net
becomes full before completing the three sets
of sweepings, empty its contents into the basin
and then continue sampling.) Where the water
depth within the bucket area is too shallow to
effectively sweep the water column, additional
substrate will need to be excavated and placed
into the wash basin.
Remove all invertebrates attached to coarse
surface substrate by scrubbing them by hand
or scrub brush into the sieve or basin (Figure
4-14). Carefully add stream water to the basin
(Figure 4-15). Rinse invertebrates from large
detritus (e.g., leaves and sticks) that may be in
Figure 4-15 Carefully adding water to the
wash basin before sample elutriation.
the basin and discard the detritus. Elutriate
the remaining contents by swirling the basin
by hand and pouring the water and low-
density contents (e.g., invertebrates and fine
organic matter) into the sieve (Figure 4-16).
This step will separate most of the fine
sediment particles from the invertebrates.
Repeat the elutriation until no organic matter
is remaining in the basin. Carefully search the
remaining basin contents for heavy-bodied
invertebrates (i.e., mollusks, mineral-cased
caddisflies, Figure 4-17). Place any heavy-
bodied invertebrates into the sieve. Empty
Figure 4-14 Scrubbing attached
invertebrates off the coarse surface
substrate in the wash basin (or sieve).
Figure 4-16 Sample elutriation in the wash
basin and pouring invertebrates and fine
detritus into the sieve.
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Figure 4-17 Carefully search the basin for
heavy-bodied invertebrates that were not
transferred to the sieve.
and rinse the wash basin. At the stream edge
carefully wash the sieve contents to one side
by gently agitating the sieve while the sieve
mesh is partially submerged in water (Figure
4-18). With the wash basin positioned
underneath, transfer the majority of the sieve
contents by hand or a minimal amount of
water using a wash bottle into a 24-oz Whirl-
Pak ® bag or other container. Agitate the
sieve again if necessary to combine the
remaining contents against one side of the
sieve (Figure 4-19). Wash the remaining
sieve contents into the bag with 95% ethanol
using the wash bottle (Figure 4-20). Ensure
Figure 4-18 Washing sieve contents to one
side by gentle agitation while sieve is
partially submerged.
Figure 4-19 Sieve contents condensed for
transfer to sample bag.
there is enough space in the bag to sufficiently
preserve the sample with additional ethanol.
Use more than one bag if necessary to contain
a sample. Pour more ethanol into the bag until
the sample is completely submerged and the
final preservative concentration is > 70%
ethanol. Note that the amount of ethanol
needed for sufficient preservation increases
with the amount of organic matter within a
sample.
Figure 4-20 Sieve contents rinsed into
sample bag (over basin) using ethanol
squirt bottle.
4.4.1.2. Sample labeling
Make a label (waterproof paper and pencil)
that includes relevant information (e.g.,
locality, habitat type, date, collector's initials).
Where more than one bag is needed to contain
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an entire sample, indicate this on the label by
writing "1 of 2", "2 of 2", etc. Place label
inside the Whirl-Pak ® with the sample and
seal the bag. When sealing the bag, remove as
much air space as possible from the bag (this
will make samples more compact for
transport). Seal the bag by folding the tab
over a couple of times then while holding the
wire ends, whirl the bag 3-4 times, and lastly
twist wire ends together. Write label
information on the field form (Figure 4-21)
and on the outside of the bag with a permanent
marker.
INVERTEBRATES
# BUCKET SAMPLES: Depositional Habitats_4_ Erosional Habitats _4_ TOTAL AREA SAMPLED: O4 m2 (each -0.05 m2)
SAMPLE ID
Four -JC- 1-0(1-4)
Four -JC-l-E (1-4)
HABITAT TYPE
Depositional
Erosional
NUMBER OF
BAGS
5
6
COLLECTED
BY:
KMF
KMF
COMMENTS
Rep #1 in 2 bags
Reps. 3 & 4 in 2 bags
Figure 4-21 Appropriate location for recording invertebrate sampling information on page
1 of field forms. Example sample information shown in red.
Repeat procedures outlined in 4.4.1.1 and
4.4.1.2 for the remaining sample replicates in
each habitat type. Thoroughly rinse sampling
equipment with stream water between sample
replicates to prevent transporting any attached
invertebrates.
4.4.1.3. Sample transport and shipping
Before leaving a site, check that all samples
are labeled properly and safely stowed in a
backpack or container (e.g., 5-gallon bucket)
for transport to the vehicle. At the vehicle
consolidate all invertebrate samples in a
crush-resistant and leak-proof container (e.g.,
cooler) for transport to the laboratory for
processing. If samples need to be shipped,
include any special shipping forms that may
be required for the ethanol-preserved samples.
4.4.2 Alternative methods
Here we will briefly identify other flow-
independent methods for quantitatively
sampling macroinvertebrates. More detailed
reviews of sampling methods for stream
invertebrates can be found in Peckarsky
(1984), Klemm et al. 1990, and Merritt et al.
(1993). Among the simplest methods is stone
sampling, where individual stones are used as
the sampling units (Ball 1979, Doeg and Lake
1981, Wrona et al. 1986, Scrimgeor et al.
1993). Stone surface area is estimated as
described in Subsection 4.2.2. Some
advantages of this method include: 1)
simplification of streambed heterogeneity, 2)
represent natural sampling units, and 3)
efficient, cost-effective method (samples
contain little detritus from which to sort
invertebrates). For small headwater streams
this method also causes minimal degradation
to the habitat (if stone area or dimensions are
measured immediately after collection) and
logistically feasible where channel width
limits use of larger samplers. Some
disadvantages of stone sampling are the
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exclusion of some habitats (e.g., interstitial
spaces, bedrock), overestimation of
extrapolations to 1 m2, and depending upon
the typical stone surface area, this method
may require large sample sizes to reduce
sample variability (Morin 1985).
Like the bucket sampler, vacuum samplers
(Boulton 1985, Brown et al. 1987, Brooks
1994) are flow-independent and can be used
with equal efficiency across habitat types and
flow conditions. Vacuum samplers are
devices that enclose a sampling area and
transfer sample material by bilge or peristaltic
pump to a sieve or net. The major drawbacks
to vacuum samplers are related to their overall
size. Most of these samplers require more
than one person for sample collection and are
likely too heavy or unwieldy for long hikes
often necessary to reach headwater study sites.
Artificial substrates, such as rock baskets and
multi-plate samplers, are commonly used for
comparing invertebrate assemblages among
sites (e.g., Poulton et al. 2003, Rinella and
Feminella 2005). Artificial substrate methods
provide the investigator with control over the
colonization or exposure time and the
standardized size may reduce sample
variability. Artificial substrates minimize
streambed disturbance to small headwater
streams. Some limitations of artificial
substrates are differential colonization among
taxa, the requirement of multiple visits for
deployment and retrieval, and susceptibility to
vandalism or natural disturbance.
Modifications to standardized substrates (e.g.,
Hester-Dendy multiplate samples) may be
required to ensure complete submergence in
shallow headwater streams (e.g., Winterbourn
1982). Variable submergence among units
counters one of the advantages of artificial
samplers.
Aquatic insects often represent a significant
proportion of the invertebrate assemblage in
headwater streams. A wide variety of traps
have been used to capture the adult life stage
of aquatic insects as they emerge from streams
(Davies 1984). Emergence traps can be
designed to collect and preserve emerging
adults daily or for up to several weeks
(LeSage and Harrison 1979, Whiles and
Goldowitz 2001). This method can be used to
easily sample across a wide range of
hydrologic permanence because it is not
dependent upon the presence of water (Progar
and Moldenke 2002, Price et al. 2003).
However, this method requires multiple visits,
is susceptible to vandalism or natural
disturbance, excludes invertebrate taxa that do
not have a winged-adult stage and, depending
upon trap design and position, may
differentially capture taxa.
References
Boulton, A. J. 1985. A sampling device that
quantitatively collects benthos in flowing or
standing waters. Hydrobiologia 127:31-39.
Brooks, S. 1994. An efficient and
quantitative aquatic benthos sampler for use
in diverse habitats with variable flow
regimes. Hydrobiologia 281:123-128.
Brown, A. V., M. D. Schram, and P. P.
Brussock. 1987. A vacuum benthos
sampler suitable for diverse habitats.
Hydrobiologia 153:241-247.
Ball, P. C. 1979. A sampling technique for
littoral stone dwelling organisms. Oikos
33:106-112.
Davies, I. J. 1984. Sampling aquatic insect
emergence. Pages 161-227 in: J. A.
Downing and F. H. Rigler (editors). A
Manual on Methods for the Assessment of
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Secondary Productivity. Blackwell
Scientific, Oxford, United Kingdom.
Delucchi, C. M. 1988. Comparison of
community structure among streams with
different temporal flow regimes. Canadian
Journal of Zoology 66:579-586.
Dietrich, M. and N. H. Anderson. 2000. The
invertebrate fauna of summer-dry streams in
western Oregon. Archivfur Hydrobiologie
147:273-295.
Feminella, J. W. 1996. Comparison of
benthic macroinvertebrate assemblages in
small streams along a gradient of flow
permanence. Journal of the North American
Benthological Society 15:651-669.
Fritz, K. M. and W. K. Dodds. 2002.
Macroinvertebrate assemblage structure
across a tallgrass prairie stream landscape.
Archivfur Hydrobiologie 154:79-102.
Frouz, J., J. Matena, and A. Ali. 2003.
Survival strategies of chironomids (Diptera:
Chironomidae) living in temporary habitats:
a review. European Journal of Entomology
100:459-465.
Hellawell, J. M. 1986. Biological Indicators
of Freshwater Pollution and Environmental
Management. Elsevier Applied Science
Publishers, London, United Kingdom.
Klemm, D. J., P. A. Lewis, F. Fulk, and J. M.
Lazorchak. 1990. Macroinvertebrate Field
and Laboratory Methods for Evaluating the
Biological Integrity of Surface Waters.
EPA/600/4-90/030. U.S. Environmental
Protection Agency, Washington, D.C.
LeSage, L. and A. D. Harrison. 1979.
Improved traps and techniques for the study
of emerging aquatic insects. Entomological
News 90:65-78.
Lytle, D. A. and N. L. Poff 2004. Adaptation
to natural flow regimes. Trends in Ecology
and Evolution 19:94-100.
Morin, A. 1985. Variability of density
estimates and the organization of sampling
programs for stream benthos. Canadian
Journal of Fisheries and Aquatic Sciences
42:1530-1534.
Peckarsky, B. L. 1984. Sampling the stream
benthos. Pages 131-160 in: J. A. Downing
and F. H. Rigler (editors). A Manual on
Methods for the Assessment of Secondary
Productivity. Blackwell Scientific, Oxford,
UK.
Poulton, B. C., M. L. Wildhaber, C. S.
Charbonneau, J. F. Fairchild, B. G. Mueller,
and C. J. Schmitt. 2003. A longitudinal
assessment of the aquatic macroinvertebrate
community in the channelized lower
Missour River. Environmental Monitoring
and Assessment 85:23-53.
Price, K., A. Suski, J. McGarvie, B. Beasley,
and J. S. Richardson. 2003. Communities
of aquatic insects of old-growth and clearcut
coastal headwater streams of varying flow
permanence. Canadian Journal of Forest
Research 33:1416-1432.
Progar, R. A. and A. R. Moldenke. 2002.
Insect production from temporary and
perennially flowing headwater streams in
western Oregon. Journal of Freshwater
Ecology 17:391-407.
Rader, R. B., D. P. Batzer, and S. A.
Wissinger (editors) 2001. Bioassessment
and Management of North American
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Freshwater Wetlands. John Wiley & Sons,
New York.
Rinella, D. J. and J W. Feminella. 2005.
Comparison of benthic macroinvertebrates
colonizing sand, wood, and artificial
substrates in a low-gradient stream. Journal
of Freshwater Ecology 20:209-220.
Rosenberg, D. M. and V. H. Resh (editors)
1993. Freshwater Biomonitoring and
Benthic Macroinvertebrates. Chapman &
Hall, New York.
Schwartz, S. S. andD. G. Jenkins. 2000.
Temporary aquatic habitats: constraints and
opportunities. Aquatic Ecology 34:3-8.
Scrimgeour, G. J., J. M. Gulp, and N. E.
Glozier. 1993. An improved technique for
sampling lotic invertebrates. Hydrobiologia
254:65-71.
Southwood, T. R. E. 1977. Habitat, the
templet for ecological strategies? Journal of
Animal Ecology 46:33 7-365.
Statzner, B. 1981. A method to estimate the
population size of benthic
macroinvertebrates in streams. Oecologia
51:157-161.
Townsend, C. R. and A. G. Hildrew. 1994.
Species traits in relation to a habitat templet
for river systems. Freshwater Biology
31:265-275.
Walker, K. F., F. Sheldon, and J. T.
Puckridge. 1995. A perspective on dryland
river ecosystems. Regulated Rivers:
Research & Management 11:85-104.
Wilding, J.L. 1940. A new square-foot
aquatic sampler. Limnological Society of
America Special Publication 4:1-4.
Williams, D. D. 1996. Environmental
constraints in temporary freshwaters and
their consequences for the insect fauna.
Journal of the North American
Benthological Society 15:634-650.
Winterbourn, M. J. 1982. The invertebrate
fauna of a forest stream and its association
with fine particulate matter. New Zealand
Journal of Marine and Freshwater Research
16:271-281.
Whiles, M. R. andB. S. Goldowitz. 2001.
Hydrologic influences on insect production
from central Platte River wetlands.
Ecological Applications 11:1829-1842.
Wrona, F. J., P. Calow, I. Ford, D. J. Baird,
and L. Maltby. 1986. Estimating the
abundance of stone-dwelling organisms: a
new method. Canadian Journal of Fisheries
and Aquatic Sciences 43:2025-2035.
Equipment and supplies (Numbers correspond
to items in Figure 4-22)
Measuring tape (50 m)
1. Bucket sampler = 5-gallon bucket with
bottom cut out w/ bottom area ~ 0.053 m2
2. Canvas skirt
3. Hand-net (243 |j,m mesh)
4. Plastic wash basins
5. Hand trowel
6. 250 |j,m sieves
7. Funnel
8. Ethanol transport jug(s)
9. Squirt bottle (250 ml)
10. 24 oz Whirl-pak bags or other sample
containers
11. Label paper
Ethanol (95%)
Sharpies & Pencils
Field forms
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Figure 4-22 Equipment used to collect and preserve benthic invertebrate samples.
Numbers correspond to the equipment and supplies list above.
4.5 Surveying the amphibian assemblage
General
This subsection provides instructions for
characterizing amphibian assemblages in
headwater streams using a visual encounter
survey. All amphibians are highly dependent
on water and the amount of moisture in the
environment influences their geographic
range, life history characteristics, and
behavior. With the exception of the tailed
frog in the western United States, most
headwater stream-dwelling amphibians are
urodels (salamanders) rather than anurans
(frogs and toads). This discussion therefore
focuses on the use of salamanders as
indicators of hydroperiod, but the methods are
similarly effective for anurans populations.
Following hatching, all stream salamanders go
through a gilled larval stage during which they
are obligate to the aquatic environment. The
larval stage may last from months to several
years depending on species and locality. At
the end of the larval stage, most species
metamorphose into juveniles and leave the
stream to become semi-aquatic or terrestrial as
adults. Adults subsequently return to streams
for courtship and egg-laying. Some
salamanders are permanently aquatic as adults
(eg., Cryptobranchus alleganiensis, Necturus
spp., etc.) and retain their gills. All stream
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salamanders are predatory and they are often
the top predators in high-gradient, fishless
headwaters (Davic and Welch 2004, Johnson
and Wallace 2005).
The fact that stream-dwelling salamander
larvae are obligate to the aquatic environment
and have larval periods that can vary greatly
in length means they are potentially ideal
indicators of stream hydroperiod. This
protocol focuses on the larval stage because
adult salamanders are less dependent on water
and may move far from the stream channel.
Unfortunately, identification of larval
salamanders can be difficult and few good
comprehensive larval keys are available.
Field crews should therefore attempt to
become familiar with the salamander species
in their area prior to sampling. Larvae of the
two-lined salamanders (Eurycea bislineata, E.
cirrigera, andE. wilderae) are among the
most commonly encountered salamanders in
streams of the eastern United States. Larvae
are dusky colored dorsally, have branched
external gills, and have 6-9 pairs of light
dorsolateral spots (Figure 4-23B). The
Appalachians are home to the greatest
salamander diversity, and larvae of
Desmognathus spp., Gyrinophilus spp. (Figure
4-24), and Pseudotriton spp. are also
frequently encountered in streams of this
region. Amphibian diversity is lower in the
western United States, but species often
encountered in streams include the giant
salamanders (Dicamptodon spp.) and tailed
frogs (Ascaphus spp.). Petranka (1998)
provides a larval key and distribution map for
salamanders of the United States. Other
regional keys and distribution maps may be
available for your area (e.g., Green and Pauley
1987, Pfmgsten and Downs 1989, Minton
2001) and many larval descriptions can be
found in the primary literature.
4.5.1 Time-constrained sampling
Time-constrained sampling is an effective way
of sampling salamander larvae from a variety
of habitats, typically with minimal cost, effort,
and stream disturbance. While the timed-
search method makes density determinations
difficult, it can be used to estimate relative
abundance of species. Timed sampling has an
additional advantage in that it increases the
chance of collecting rare taxa, or rare
individuals when salamanders are scarce
(Crump and Scott 1994, Barr and Babbitt
2001). Chalmers and Droege (2002)
additionally found that timed-search sampling
was more effective than use of leaf litter bags
in estimating abundance of larval E.
bislineata. We chose the time-constrained
approach for this protocol because it is robust
and best suited for collection of larvae from
streams across multiple regions where species
composition and densities may be highly
variable. The method is also effective over a
wide range of stream size, hydroperiod, and
condition.
Figure 4-23 Northern tow-lined
salamander, Eurycea cirrigera, from
Robinson Forest, KY: A) egg clutch; B)
larva; and C) adult.
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Figure 4-24 Larval spring salamander,
Gyrinophilus porphyriticus, from Robinson
Forest, KY.
Procedure
When possible, sampling should be conducted
with clear skies to maximize visibility. The
amphibian survey reach and primary sampling
reach should have similar discharge and
habitat characteristics. Sampling typically
begins ca. 10-m upstream of the primary 30-m
sampling reach and progresses in the upstream
direction. If there is an obvious change in
habitat or discharge (i.e., a tributary
confluence or headcut) above the primary
reach, then sampling should be conducted
downstream of the primary sampling reach.
Sampling begins after noting the starting time
and then continues for exactly 30 minutes.
Only one person should conduct the sampling
to standardize the level of effort and, if
possible, the same person should conduct the
survey at each site to minimize sampling
variability.
Sampling is confined to the wetted area of the
stream because the survey focuses on the
larval stage. One crew member moves
carefully upstream (or downstream in some
cases) turning loose cover objects (leaves,
cobble, woody debris, etc.) in all available
habitat types (shallow, deep, fast, slow, etc.).
Salamander larvae are often found in isolated
backchannels or at the stream margins where
there may only be a thin surface film. Cover
objects are turned individually by hand rather
than using kick nets or other more destructive
sampling methods. This saves time in sorting
through debris and allows for the survey to
cover a greater stream area. The investigator
should slowly and methodically turn random
cover objects from bank to bank while moving
along the stream. Salamanders encountered
are carefully collected into the hand net for
identification. It should not be the intent of
the investigator to turn over every object in
the stream and there is no distance objective.
Surveys in larger perennial streams with a
greater wetted area will therefore cover less
stream length than surveys done in small
intermittent stream reaches. The objective is
to keep the level of effort the same in every
study reach, regardless of stream size or
habitat types. Approximate length (m) of
stream surveyed should be noted at the end of
the 30-minute survey.
Larval salamanders observed during the
survey are identified in the field when possible
and recorded on the amphibian survey field
sheet along with corresponding life stage
(larva, juvenile, adult) (Figure 4-25). Mean
snout-vent length (SVL) for each cohort
should be visually estimated (mm) for each
species and recorded on the field form to help
determine when larvae may have hatched.
Presence of larger/older larvae of species with
multi-year larval stages (e.g., Desmognathus
quadramaculatus, Gyrinophilus porphyriticus,
northern populations of E. bislineata) should
be noted as this can be an important indicator
of stream permanence. Unknown species and
voucher specimens should be recorded
photographically, showing top and side views
at a minimum. Each species should be
125
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vouchered for each area sampled (i.e.,
ecoregion, national forest, etc.). Salamanders
should not be collected and returned to the
laboratory without appropriate collection
permits. Salamanders collected for vouchers
or for species confirmation should be
anaesthetized with 0.1% MS-222 (tricaine
methylsulfonate) (Beachy 1994) and then
preserved with a 10% formalin solution.
Preserved specimens should be placed in vials
labeled with the site name, date, and name of
the collector. Photos and preserved specimens
may be sent to regional experts for species
confirmation. Though the survey is aimed at
the larval stage, adults and egg clutches found
during the survey should also be noted on the
field form and photographed. Any fish
observed during sampling should also be
recorded.
4.5.2 Alternative methods
Salamanders have been collected from streams
using a wide variety of sampling methods
(reviewed by Heyer et al. 1994). Larvae may
be qualitatively sampled using kick-nets or
conventional dredge nets typically used in
benthic macroinvertebrate sampling. Such
collection methods; however, can be
destructive and time-consuming and may not
adequately represent rare taxa. Typical
quantitative approaches include: other benthic
sampling devices (e.g., benthic corers, Surber
samplers), quadrats (e.g., Welsh and Lind
1996, Rocco and Brooks 2000), transects (e.g.,
Resetarits 1997, Welsh and Oliver 1998), and
artificial habitats (Pauley 1998). Bury and
Corn (1991) provide an example of a more
intensive sampling method for western
streams, whereby all moveable objects are
removed from a 10-m stream section by a two
people over ca. 5 hours.
126
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AMPHIBIAN ASSESSMENT FORM
Page 4 of4
SITE NAME:
DATE:
COLLECTION TIME:
to
REACH LENGTH COVERED (m):
SURVEYOR
SPECIES / LIFE STAGE
SPECIES
#LARVAE ftlUVENILE #ADULT
TOTAL
VOUCHER?
De&nognathus monticola
De&nognathus ochrophaeiis
De&nognathus -welteri
Ewycea bislineata
Eurycea longicauda
GyrinopHlus porphyriticus
Psedotriton montanus
Pseudotriton ruber
OTHER
NOTES ON AMPHIBIANS
NOTES ON FISH (include species present)
Figure 4-25 Amphibian survey field form.
References
Barr, G. E., and K. J. Babbitt. 2001. A
comparison of 2 techniques to sample larval
stream salamanders. Wildlife Society
Bulletin 29:1238-1242.
Beachy, C. K. 1994. Community ecology of
streams: effects of two species of predatory
salamanders on a prey species of
salamander. Herpetologica 50:129-136.
127
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Bury, B. R., and P. S. Corn. 1991. Sampling
Methods for Amphibians in Streams of the
Pacific Northwest. Gen. Tech. Rep. PNW-
GTR-275. Department of Agriculture,
Forest Service, Pacific Northwest Research
Station. Portland, Oregon.
Chalmers, R. 1, and S. Droege. 2002. Leaf
litter bags as an index to populations of
northern two-lined salamanders (Eurycea
bislineatd). Wildlife Society Bulletin 30:71-
74.
Crump, M. L., and N. J. Scott, Jr. 1994.
Standard techniques for inventory and
monitoring: visual encounter surveys. Pages
84-92 in W. R. Heyer, M. A. Donnelly, R.
W. McDiarmid, L. C. Hayek, and M. S.
Foster (editors). Measuring and Monitoring
Biological Diversity: Standard Methods for
Amphibians. Smithosonian Institution,
Washington, D.C.
Davic, R. D. and H. H. Welsh, Jr. 2004. On
the ecological roles of salamanders. Annual
Review of Ecology, Evolution and
Systematics 35:405-434.
Green, N. B., and T.K.Paul ey. 1987.
Amphibians and Reptiles in West Virginia.
University of Pittsburgh Press, Pittsburgh,
Pennsylvania.
Heyer, W. R., M. A. Donnelly, R. W.
McDiarmid, L. C. Hayek, and M. S. Foster
(editors). 1994. Measuring and Monitoring
Biological Diversity: Standard Methods for
Amphibians. Smithosonian Institution,
Washington, D.C.
Johnson, B. R. and J. B. Wallace. 2005.
Bottom-up limitation of a stream salamander
in a detritus-based food web. Canadian
Journal of Fisheries and Aquatic Sciences
62:301-311.
Minton, S. A. 2001. Amphibians and Reptiles
of Indiana. Indiana Academy of Science,
Indianapolis, Indiana.
Pauley, T. K. 1998. A new technique to
monitor larval and juvenile salamanders in
stream habitats. Banisteria 12:32-36.
Petranka, J. W. 1998. Salamanders of the
United States and Canada. Smithsonian
Institution Press. Washington, D.C.
Pfingsten, R. A, and F. L. Downs (editors).
1989. Salamanders of Ohio. Ohi o
Biological Survey Bulletin Vol. 7(2), Ohio
Biological Survey, Columbus, Ohio.
Resetarits, W. J. Jr. 1997. Differences in an
ensemble of streamside salamanders
(Plethodontidae) above and below a barrier
to brook trout. Amphibia-Reptilia 18:15-25.
Rocco, G. L., and R. P. Brooks. 2000.
Abundance and Distribution of a Stream
Plethodontid Salamander Assemblage in 14
Ecologically Dissimilar Watersheds in the
Pennsylvania Central Appalachians. Final
Technical Report No. 2000-4 of the Penn
State Cooperative Wetlands Center,
University Park, Pennsylvania.
Welsh, H. H., and A. J. Lind. 1996. Habitat
correlates of the southern torrent
salamander, Rhyacotriton variegates
(Cauadata: Rhyacotritonidae), in
northwestern California. Journal of
Herpetology 30:385-398.
Welsh, H.H., and L.M.Oliver. 1998. Stream
amphibians as indicators of ecosystem
stress: a case from California's redwoods.
Ecological Applications 8:1118-1132.
128
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Equipment and supplies 0.1% MS-222 (optional)
Aquarium dip net (approximately 15.5 x 12 10% formalin (optional)
cm, 1-mm mesh) Digital camera
Wristwatch or stopwatch Field forms
Specimen containers (optional)
129
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5 APPENDIX FIELD FORMS
130
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HEADWATER STREAM ASSESSMENT FORM
Pagel of 4
STREAM NAME (& ABREV):
COUNTY: STATE: USGS QUAD NAME:_
SITE#
DATE: /
REACH LENGTH (m):
LATITUDE:
LONGITUDE:
DIRECTIONS TO STREAM SITE
STUDY REACH HYDROLOGIC CONDITION
D Visible surface flow continuous (4)
D Surface water in pools only (1)
D Visible flow interstitial (3) D Surface water present but no visible flow (2)
D No surface water (0)
MAX. POOL DEPTH (cm)
DEPTH TO BEDROCK /
GROUNDWATER (m)
(3 measures in depositional habitat)
SINUOSITY
(number of bends)
DISTANCE TO NEAREST
SURFACE WATER (m)
CHANNEL SLOPE (%)
(for three 10m sections of study reach)
% CANOPY COVER
(facing upstream, downstream, right & left banks)
0 <100 100-500 >500
PRESENCE OF HEADCUT
IN REACH
ALGAL COVER INDEX
# CORES FOR SUBSTRATE MOISTURE
(depositional)
I1"
Terrestrial Herbaceous Vegetation in Active Channel?
Base Flow Conditions?(Y/N) Date of Last Precipitation?
Roots of Riparian Vegetation in Active Channel?
IN SITU WATER QUALITY MEASUREMENTS
Location of
Measurements
Cond
(^S/cm)
Temp
°
DO
(mg/1)
Comments
BIOTIC SAMPLES (ALL TAKEN IN THALWEG & LABEL SAMPLES COMPLETELY)
# BUCKET SAMPLES: Depositional Habitats
INVERTEBRATE
Erosional Habitats TOTAL AREA SAMPLED: m2 (each ~ 0.05 m2)
SAMPLE ID
HABITAT TYPE
NUMBER OF
BAGS
COLLECTED
BY:
COMMENTS
# STONES SAMPLED: Depositional Habitats_
ALGAE
Erosional Habitats
TOTAL AREA SAMPLED:
cm (each 12)
BRYOPHYTES SAMPLED: Y
SAMPLE ID
COMMENTS
131
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* MEASURES TAKEN IN THALWEG § Where EPA Width > 2.2X BE Width then indicate: >2.2 BE Page 2 of 4
Meter #
0
i
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
Modal
Sediment
Particle
Size
(mm) *
Water
Depth
(cm)*
Habitat
Type
(E/D)
Notes
(e.g., LWD,
Leafpack)
Velocity
(m/s)*
Wetted
Width
(m)
BE
Width
(m)
BE
Depth
(m)*
EPA
width
(m)§
132
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STREAM DISCHARGE
Page 3 of 4
Wetted Width (m)
CELL WIDTH (m):
Depth(cm)
Velocity(m/s)
Q (m3s ') =
Discharge procedure:
Velocity procedure/meter model:
DRAWING OF STREAM REACH
FLOW-*
30
— 15
NOTES & SITE PHOTOS
Wentworth scale (mm): <2, 2-4, 4-8, 8-16, 16-32, 32-64, 64-128, 128-256, 256-512, >512
ACI: l=substrate is rough with no apparent growth 1.5=substrate is slimy, but biofilm not visible 2=thin layer
visible (0.5-1mm thick) 3=algal mat 1-5 mm thick & filamentous algae rare 4=algal mat 5-20 mm thick &
filamentous algae common 5=algal mat >2 cm thick &/or filamentous algae dominate
133
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AMPHIBIAN ASSESSMENT FORM
Page 4 of 4
SITE NAME:
DATE:
COLLECTION TIME:
to
REACH LENGTH COVERED (m):
SURVEYOR:
SPECIES / LIFE STAGE
SPECIES
#LARVAE
#JUVENILE
#ADULT
TOTAL
VOUCHER?
Desmognathus fuscus
Desmognathus monticola
Desmognathus ochrophaeus
Desmognathus welteri
Eurycea bislineata
Eurycea longicauda
Gyrinophilus porphyriticus
Psedotriton montanus
Pseudotriton ruber
OTHER
NOTES ON AMPHIBIANS
NOTES ON FISH (include species present)
134
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