-------
Reviewed by (initial)
PHab: CHANNEL/RIPARIAN TRANSECT FORM - RIVERS
SITE NAME:
DATE: g I 5 I 98 VISIT DOJg|lD2D3
SITE ID: ORRV _3_ _|5_ -_35_2
TEAM ID (X):
D2 D3 D4 D5 D6 D7 D8
TRANSSECT(X): DA DB DC Efo
DH PI DJ DK
"LITTORAL" SUBSTRATE INFORMATION
SLOPE / BEARING / DISTANCE
SHORE
IOM SEC
BL
GF
SA
FN
HP
WD
OT
RS
CB
GC
GF
SA
FN
HP
WD
OT
BOTTOM
BL
GC
GF
SA
FN
HP
WD
OT
GF
SA
FN
HP
WD
OT
CLASS
BOTTOM SUBSTRATE FROM (X ONE):
JUDGEMENT -OR- OBS. @ 5 LITTORAL DEPTHS
INTENDED TRANSECT SPACING xxx (mH
RS - BEDROCK (SMOOTH) • (LARGER THAN A CAR)
SLOPE
xx.x %
RR - BEDROCK (ROUGH) • (LARGER THAN A CAR)
BL - BOULDER (250 TO 4000 MM) • (BASKETBALLTO CAR)
CB - COBBLE (64 TO 250 MM) • (TENNIS BALL TO BASKETBALL)
GC - COARSE GRAVEL (16 TO 64 MM) • (MARBLE TO TENNIS BALL]
BEARING
0 - 360°
H-OO
DISTANCE
xxx (m)
FLAG
Fl
GF - FINE GRAVEL (2 TO 16 MM) • (LADYBUG TO MARBLE)
SA- SAND (0.06TO2MM) • (GRITTY-UP TO LADYBUG SIZE)
CANOPY DENSITY @ BANK
- DENSIOMETER (OTO 17 MAX)
FN - SILT/CLAY/MUCK* (NOTGRITTV)
HP - HARDPAN • (FIRM, CONSOLIDATED FINE SUBSTRATE)
UP
WD-WOOD- (ANY SIZE)
LEFT
OT- OTHER © (COMMENT)
FLAG
DOWN
RIGHT
BANK CHARACTERISTICS
CIRCLE ONE
V
G
F
Near Vertical/Undercut (>75°)
Steep (30-75°)
Gradual (5-30°)
Flat(<5°)
WETTED WIDTH
BANKFUU. WIDTH
BANKFUU. HT.
INCISED HT.
XXX(m)
O
75-
0.7
FLAG
K
BANK
ANGLES
Rat (<5'
LARGE WOODY DEBRIS in Wet Channel (10 x 20m Plot)
DIAMETER
0.3 - 0.6 m
0.6 - 0.8 m
0.8-1.0m
>1.0m
WOOD ALL/PART IN WETTED CHANNEL
LENGTH5-15m 15-30m
DRY OUT ALL/PART IN BANKFULL CHANNEL
0.3 - 0.6 m
0.6 - 0.8 m
0.8-1.0m
>1.0m
DEPTH
SONAR (It) xx
POLE (m) x.x)
0,6
0.6
0-6
IN SITU WATER
MEASUREMENTS
WATER TEMPERATURE °C (xxxx)
CONDUCTIVITY nS/CM (xxxx]
(OS
FLAG
COMMENTS
&& TKEP
K=
Flag Codes: K - no measurement made; U = suspect measurement; F1, F2, etc. = misc. flags assigned by each field crew. Explain all flags in comments section
on this side or on Side 2 of this form.
Rev. 05/29/98 (_rvphc_.98)
PHab: CHANNEL/RIPARIAN TRANSECT FORM - RIVERS -1
Figure 6-4. Channel/Riparian transect form - page 1 (front side).
-------
interval between thalweg profile measure-
ments, measure the wetted channel width with
a laser range finder at several locations near
the upstream end of the reach and multiply it
by 40 (100) to set the river sample reach
length. Then divide that reach length by 100
(or 200) to set the thalweg increment distance.
Following these guidelines, you will be mak-
ing 100 or 200 evenly-spaced thalweg pro-
file measurements, 10 or 20 between each
detailed channel cross section where littoral/
riparian observations are made. The number
and spacing of measurements are as follows
for the two different sample reach lengths:
40 Ch-W
number spacing
100 Ch-W
number spacing
Transects and 10 4 Ch-W 10 10 Ch-W
Riparian Plots
Thalweg 100 0.4 Ch-W 200 0.5 Ch-W
Depth
measurements
Thalweg
Substrate,
Habitat Class
100 0.4 Ch-W 100 1.0 Ch-W
6.3 Logistics, Work
Flow, and Defining
Sample Locations
The two-person habitat assessment team
uses the most nimble of the selection of wa-
tercraft judged capable of navigating the river
reach. In a single midstream float down the
40 or 100 Channel-width reach, the team ac-
complishes a reconnaissance, a sonar/pole
depth profile, and a pole-drag to tally snags
and characterize mid-channel substrate. The
float is interrupted by stops at 11 transect lo-
cations for littoral/riparian observations. They
determine (and mark ~ optional) the position
of each successive downstream transect us-
ing a laser range finder to measure out and
mentally note each new location 4 (or 10)
channel-width's distance from the preceding
transect immediately upstream. The crew then
floats downstream along the thalweg to the
new transect location, making thalweg pro-
file measurements and observations at 10 (or
20) evenly-spaced increments along the way.
When they reach the new downstream transect
location, they stop to do cross-section, littoral,
and riparian measurements. Equipping the
boat with a bow or stern anchor to stop at
transect locations can greatly ease the shore
marking operation and shoreline measure-
ment activities. In addition, while they are
stopped at a cross-section station, the crew
can fill out the habitat "typing" entries retro-
spectively and prospectively for the portion
of the stream distance that is visible up- and
downstream. They can also record reconnais-
sance and safety notes at this time. While
stopped at the transect location, the crew
makes the prescribed measurements and ob-
servations, collects biological samples,
backsites slope and bearing towards the pre-
vious upstream transect, and sets or mentally
notes eye-level flags or reference points on
shore for subsequent backsites. The habitat
crew also assists the electrofishing boat crew
over jams and helps to conduct shuttles (this
can take considerable time where put-ins and
take-outs are distant).
6.4 Reconnaissance
and Reach Marking
The purpose of the reconnaissance is to
locate (and optionally mark) the reach sam-
pling location and to inform the second boat
of the route, craft, and safety precautions
needed during its subsequent electrofishing
activities. After finding adequate put-in and
take-out locations, the team may opt to mark
the upstream end of the sample reach end with
-------
colored flagging. Based on several channel
width measurements using a laser range finder,
they determine the sample reach length (40 x
or 100 x Channel Width), the transect spac-
ing (4 x or 10 x Channel Width) and thalweg
sampling interval (0.5 x Channel Width). As
the crew floats downstream, they stop (and
optionally flag) 11 transect locations along the
riverbank in the process of carrying out slope,
bearing, and distance backsites. As the team
floats downstream, they may choose andcom-
municate to the electrofishing crew the most
practical path to be used when fishing with a
less maneuverable boat, taking into consider-
ation multiple channels, blind channels, back-
waters, alcoves, impassible riffles, rapids,
jams, and hazards such as dams, bridges and
power lines. They determine if and where
tracking or portages are necessary.
6.5 Thalweg Profile
"Thalweg" refers to the flow path of the
deepest water in a river channel. The thalweg
profile is a longitudinal survey of maximum
depth and several other selected characteris-
tics at 100 (or 200) near-equally spaced points
along the centerline of the river between the two
ends of the river reach (Figure 6-1). For practi-
cal reasons, field crews will approximate a thal-
weg profile by sounding along the river course
that they judge is deepest, but also safely navi-
gable. Data from the thalweg profile allows cal-
culation of indices of residualpool volume, river
size, channel complexity, and the relative pro-
portions of habitat types such as riffles and pools.
The procedure for obtaining thalweg profile
measurements is presented in Table 6-2. Record
data on the Thalweg Profile Form as shown in
Figure 6-3.
6.5.1 Thalweg Depth
Mh mmm *^
Profile
A thalweg depth profile of the entire 40
or 100 Channel-width reach shall be approxi-
mated by a sonar or sounding rod profile of
depth while floating downstream along the
deepest part of the channel (or the navigable
or mid-channel path). In the absence of a re-
cording fathometer (sonar depth sounder with
strip-chart output or electronic data recorder),
the crew records depths at frequent, relatively
evenly-spaced downstream intervals while
observing a sonar display and holding a
surveyor's rod off the side of the boat (see
subsection 6.5.2, below). The sonar screen is
mounted so that the crew member can read
depths on the sonar and the rod at the same
time. The sonar sensor may need to be
mounted at the opposite end of the boat to
avoid mistaking the rod's echo for the bot-
tom, though using a narrow beam (16 degree)
Sonar transducer minimizes this problem. It
is surprisingly easy to hold the sounding rod
vertical when you are going at the same speed
as the water. In our river trials, one measure-
ment every half-channel-width (10 to 15 m)
in current moving at about 0.5 m/s resulted in
one measurement every 20 to 30 seconds. To
facilitate accomplishing this work fast enough,
the field form only requires "checks" for any
observations other than depth measurements.
To speed operations further, it may also be
advantageous to mount a bracket on the boat
to hold the clipboard.
6.5.2 Pole Drag for
Snags and Substrate
Characteristics
The procedure for obtaining pole drags
for snags and substrate characteristics is pre-
sented in Table 6-2. While floating down-
stream, one crew member holds a calibrated
PVC sounding tube or fiberglass surveying
rod down vertically from the gunwale of the
boat, dragging it lightly on the bottom to si-
multaneously "feel" the substrate, detect
-------
snags, and measure depth with the aid of so-
nar. The number of large snags hit by this rod
shall be recorded as an index of fish cover
complexity (modification of Bain's "snag
drag"). While dragging the sounding rod
along the bottom, the crew member shall
record the dominant substrate type sensed by
dragging the rod along the bottom (bedrock/
hardpan, boulder, cobble, gravel, sand, silt &
finer) (Figure 6-3). In shallow, "wild," fast-
water situations, where pole-dragging might
be hazardous, prews will estimate bottom con-
ditions the best they can visually and by us-
ing paddles and oars. If unavoidable, suspend
measurements until out of Whitewater situa-
tions, but make notes and appropriately flag
observations concerning your best judgements
of depth and substrate.
6.5.3 Channel Habitat
Classification
The crew will classify and record the
channel habitat types shown in Figure 6-3
(fall, cascade, rapid, riffle, glide, pool, dry)
and check presence of off-channel and back-
water habitat at a spatial resolution of about
0.4 channel-widths on a 40 Channel-width
reach. On a 100 Channel-width reach habitat
classifications are made every 1.0 channel-
widths and off-channel and backwater habi-
tat presence is checked every 0.5 channel-
width distance — the same interval as thalweg
'depths. The resulting database of traditional
visual habitat classifications will provide a
bridge of common understanding with other
studies. The procedures for classifying chan-
nel habitat are presented in Table .6-2. The
designation of side channels, backwaters and
other off-channel areas is independent of the
main-channel habitat type. Main channel
habitat units must meet a minimum size crite-
ria in addition to the qualitative criteria listed
in Table 6-3. Before being considered large
enough to be identified as a channel-unit scale
habitat feature, the unit should be at least as
long as the channel is wide. For instance, if
there is a small, deep (pool-like) area at the
thalweg within a large riffle area, don't record
it as a pool unless it occupies an area about as
wide or long as the channel is wide.
Mid-Channel Bars, Islands, and Side
Channels pose some problems for the sam-
pler conducting a thalweg profile and neces-
sitate some guidance. Mid-channel bars are
defined here as channel features below the
bankfull flow level that are dry during
baseflow conditions (see Section 6.6.4 for
definition of bankfull channel). Islands are
channel features that are dry even when the
river is at bankfull flow. If a mid-channel fea-
ture is as high as the surrounding flood plain,
it is considered an island. Both mid-channel
bars and islands cause the river to split into.
side channels. When a bar or island is encoun-
tered along the thalweg profile, choose to
navigate and survey the channel that carries
the most flow.
When side channels are present, the com-
ments column of the Thalweg Profile form
should reflect their presence by checking the
"Off-Channel" column., These checkmarks
will begin at the point of divergence from the
main channel, continuing downstream to the
point of where the side channel converges with
the main channel. In the case of a slough or
alcove, the "off-channel" checkmarks should
continue from the point of divergence.
6.6 Channel Margin
("Littoral") And Riparian
Measurements
Components of this section include slope
and bearing, channel margin depth and sub-
-------
Table 6-2. Thalweg Profile Procedure.
1. Determine the interval between measurement stations based on the wetted width used to determine
the length of the sampling reach.
2. Complete the header information on the Thalweg Profile Form, noting the transect pair (upstream to
downstream).
3. Begin at the upstream transect (station"!" of "20" or station"!" of "10").
Thalweg Depth Profile
a) While floating downstream along the thalweg, record depths at frequent, approximately even-
spaced downstream intervals while observing a sonar display and holding a surveyor's rod off the
side of the boat.
b) A depth recording approximately every 0.4 (or 0.5) channel-width distance is required, yielding
10 (or 20) measurements between channel/riparian cross-section transects.
c) If the depth is less than approximately 0.5 meters, or contains a lot of air bubbles, the sonar
fathometer will not give reliable depth estimates. In this case, record depths using a calibrated
measuring rod. In shallow, "wild," fast-water situations depths may have to be visually estimated
to the nearest 0.5 meter.
d) Measure depths to nearest 0.1 m and record in the "SONAR" or "POLE" column on the Thalweg
Profile Form.
Pole Drag for Snags and Substrate Characteristics
a) From the gunwale of the boat, hold a fiberglass surveying rod or calibrated PVC sounding tube
down vertically into the water.
b) Lightly drag the rod on the river bottom to "feel" the substrate and detect snags.
c) Observations are taken at half the frequency as depth measurements (i.e., at every other depth
measurement point on 100 Channel-Width reaches).
d) Record the number of snags hit by the rod and the dominant substrate type sensed by dragging
the rod along the bottom.
e) On the Thalweg Profile Form, circle the appropriate "SUBSTRATE" type and tally the number of
"SNAGS".
Channel Habitat Classification
a) Classify and record the channel habitat type at increments of every 1.0 channel width.
b) Check for off-channel and backwater habitat at increments of every 0.4 (or 0.5) channel width.
c) If channel is split by a bar or island, navigate and survey the channel with the most discharge.
d) When a side channel is encountered, check the "OFF-CHANNEL" column beginning with the
point of divergence from the main channel, continuing downriver until the side channel
converges with the main channel.
e) On the Thalweg Profile Form, circle the appropriate "CHANNEL HABITAT" and check the off-
channel column as described in (d) above.
4. Proceed downriver to the next station ("2"), and repeat the above procedures.
5. Repeat the above procedures until you reach the next transect. Prepare a new Thalweg Profile Form,
then repeat the above procedures for each of the reach segments, until you reach the downriver end of
the sampling reach (Transect "K").
-------
; Table 6-3. Channel Unit Categories.
Class (Code)
Channel Unit Habitat Classes"
Description
Pools (PO):
? Plunge Pool
Trench Pool
Lateral Scour Pool
• Backwater Pool
Dam Pool
Glide (GL)
piffle (RI)
tr
feapid (RA)
i
^Cascade (CA)
!Fails(FA)
«-._-.
f Dry Channel (DR)
Ipff-Channel Areas
f--
Still water, low velocity; smooth, glassy surface, usually deep compared to other
parts of the channel:
Pool at base of plunging cascade or falls.
Pool-like trench in the center of the stream
Pool scoured along a bank.
Pool separated from main flow off the side of the channel.
Pool formed by impoundment above dam or constriction.
Water moving slowly, with a smooth, unbroken surface. Low turbulence.
Water moving, with small ripples, waves and eddies -- waves not breaking,
surface tension not broken. Sound: "babbling", "gurgling".
Water movement rapid and turbulent, surface with intermittent Whitewater and
breaking waves. Sound: continuous rushing, but not as loud as cascade.
Water movement rapid and very turbulent over steep channel bottom. Most of
the water surface is broken in short, irregular plunges, mostly Whitewater.
Sound: roaring.
Free falling water over a vertical or near vertical drop into plunge, water
turbulent and white over high falls. Sound: from splash to roar.
No water in the channel
Side-channels, sloughs, backwaters, and alcoves that are separated from the
main channel.
? Note that in order for a channel habitat unit to be distinguished, it must be at least as wide or long as the
t:ehannel is wide.
strata, large woody debris, bank angle and @ Q 1 SIODG
channel cross-section morphology, canopy "
cover, riparian vegetation structure, fish cover, BSSFIFIQ
and human influences. All measurements are The slop£j or gradient5 of the stream
recorded on the two-sided Channel/Riparian reach is useful in three different ways. First,
Transect Form (Figures 6-4 and 6-5). the overall stream gradient is one of the ma_
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Reviewed by (initial)
PHab: CHANNEL/RIPARIAN TRANSECT FORM - RIVERS (continued)
SITE NAME: BEAVER RlV£/? DATE: # I 5 / 98 VISIT DO $10203
SITE ID:
ORRV °l 8 -_3^_9_ TEAMID(X): 01 Q2 Q3 D4 O5 D6 Q7
D8
TRANSSECT(x>: QA DB DC HD DE DF DG DH DI DJ DK
VISUAL RIPARIAN
ESTIMATES
RIPARIAN
VEGETATION COVER
(10mx20mPlot)
LEFT
BANK
RIGHT
BANK
FLAG
0. ABSENT ( 0%) D = DECIDUOUS
1- SPARSE ( C.CONFEROUS .
2 -MODERATE (10-40%) E . BROADLEAF EVERGREEN
3 - HEAVY (40 - 75%) M - MIXED
4 -VERY HEAVY 1 >75%) N-NONE
CANOPY (>5m HIGH)
VEGETATION TYPE
BIG TREES (TRUNK > 0.3 m DBH)
SMALLTREES (TRUNK > 0 Z m DBH)
D C E M C£fl
0 1234
©1234
D C E MOO
( ON BANK
2) P C B
(3) P C B
(o) P C B
0 Q5 C B
Q) P C B
(2? PCS
(a) PCS
@ P C B
(jj) P C B
(^ P C p
(p) P ' C B
_. . — '""i'mViVmr^^s--;;^
yy PCS
©PCS
£5) P C B
0. (PJ C B
©PCS
O P C B
(o) P C B
O P C B
(a) P OB
(5J P C B
© P C B
COMMENTS (A
FLAG
F7
ddition
FISH
COVER/
OTHER
(10 mX 20m Plot)
FILAMENTOUS ALGAE
MACROPHYTES
WOODY DEBRIS (> 0.3 m - BIG)
BRUSH/WOODY DEBRIS (-= 0.3 m - SMALL)
.OVERHANGING VEG. (s 1 m OF SURFACE)
UNDERCUT BANKS
BOULDERS/LEDGES
ARTIFICIAL STRUCTURES
COVER IN-CHAN
0 = ABSENT (
1 = SPARSE <
2. MODERATE 10
3 = HEAVY 40
4 = VERY HEAVY
CIRCLE ONE
@ 1234
0 Q 2 3 4
(5) 1 2 3 4
Q^ 1234
(0\ 1234
00234
00234
(J5 1234
CHANNEL CONSTRAINT
DISTANCE FROM SHORE TO RIPARIAN VEGETATION (XX] xxx*- £
CIRCLE ONE
NEL
0%)
10%)
- 40%)
-75%)
>75%)
FLAG
D
(Q CHANNEL IS CONSTRAINED.
U CHANNEL IS UNCONSTRAINED IN BROAD VALLEY.
B CHANNEL IS IN BROAD VALLEY BUT CONSTRAINED BY INCISION.
N CHANNELISIN NARROW VALLEY BUT NOT VERY CONSTRAINED.
CIRCLE ONE
^E^ 1 COULD READILY SEE OVER THE BANK.
NO 1 COULD NOT READILY SEE OVER THE BANK.
al space available on Side 1)
RAfLRoAp GRADE
.................. ..............J
Flag Codes: K - no measurement made; U - suspect measurement; F1, F2, etc. = misc. flags assigned by each field crew. Explain all flags in comments section
on mis side or on Side 1 of this form.
Rev. 05/29«8 <_rvphc_.98)
PHab: CHANNEL/RIPARIAN TRANSECT FORM - RIVERS - 2
Figure 6-5. Channel/Riparian transect form-page 2 (backside).
•fit
-------
jor stream classification variables, giving an
indication of potential water velocities and
stream power; both of which are in turn im-
portant controls on aquatic habitat and sedi-
ment transport within the reach. Second, the
spatial variability of stream gradient is a mea-
sure of habitat complexity, as reflected in the
diversity of water velocities and sediment sizes
within the stream reach. Lastly, using meth-
ods described by Stack (1989), Robison and
Kaufmann (1994), and Kaufmann et al.,
(1999), the water surface slope will allow us
to compute residual pool depths and volumes
from the multiple depth and width measure-
ments taken in the thalweg profile (Subsec-
tion 6.5). Compass Bearings between cross
section stations, along with the distance be-
tween stations, will allow us to estimate the
sinuosity of the channel (ratio of the length of
the reach divided by the straight line distance
between the two reach ends).
Measure slope and bearing by
"backsiting" upstream from cross-section sta-
tion B to A, C to B, D to C, etc., down to the
llth cross section (Figure 6-1). To measure
the slope and bearing between adjacent sta-
tions, use an Abney Level (or clinometer), and
a bearing compass following the procedure
presented in Table 6-4. Record data for slope
and bearing in the Slope/Bearing/Distance
section of the Channel/Riparian Transect
Form (Figure 6-4).
It may be necessary to set up intermedi-
ate slope and bearing stations between the
normal 11 stations if you do not have direct
line-of-site along (and within) the channel
between stations. This can happen if brush is
too heavy or if there are tight meander bends
or sharp slope breaks. To backsite upstream
from supplemental stations, treat them just as
you do a normal transect location in steps 1
to 6 of Table 6-4. Record supplemental slope,
bearing, and distance backsites sequentially
in the spaces provided on the field form.
6.6,2 Channel Margin
Depth and Substrate
Substrate size is one of the most impor-
tant determinants of habitat character for fish
and macroinvertebrates in streams. Along with
bedform (e.g., riffles and pools), substrate in-
fluences the hydraulic roughness and conse-
quently the range of water velocities in the
channel. It also influences the size range of
interstices that provide living space and cover
for macroinvertebrates, salamanders, and
sculpins (as well as other benthic fishes). Sub-
strate characteristics are often sensitive indi-
cators of the effects of human activities on
streams. Decreases in the mean substrate size
and increases in the percentage of fine sedi-
ments, for example, may destabilize channels
and indicate changes in the rates of upland
erosion and sediment supply.
Channel margin depths are measured
along the designated shoreline at each transect
within the 10m swath of the 20m channel mar-
gin length that is centered on the transect lo-
cation. Dominant and sub-dominant bottom
substrates are determined and recorded at 5
systematically-spaced locations that are lo-
cated by eye within the 1 Om x 20m plot. These
methods are an adaptation of those used by
the U.S.EPA for evaluating littoral substrates
in lakes (Kaufmann and Whittier 1997),
where the substrate size may be visually as-
sessed or estimated by "feel" using the sur-
veyors rod or PVC sounding tube in deep,
turbid water. The procedure for obtaining
channel margin depth and substrate measure-
ments is described in more detail in Table 6-
5. Record these measurements on the Chan-
nel/Riparian Transect Form as shown in
Figure 6-4.
-------
Table 6-4. Procedure for Obtaining Slope and Bearing Data.
1. Set eye-level flagging atupstream transect: Place flagging or mentally note a landmark at a standardized
eye level along the shoreline at Transect A while doing shoreline measurements. To accomplish this, sit
in the boat with your clinometer or Abney level held against your measuring rod at a comfortable,
standardized height above the water surface (or designated place on bottom of boat). This shall be the
same height you plan to use for all slope backsites from downstream. Site towards the nearby bank with
the clinometer or Abney level indicating 0% slope. Note the level on the object sited and place flagging
on it (optional). Accuracy of the clinometer measurements can be checked occasionally against a
surveyors level.
2. Using the laser rangefinder. determine and record the intended location and distance of the next
downstream Transect.
3. Float downstream (doing your thalweg profile measurements at 10 or 20 increments) to Transect B,
where the next channel/riparian station is located.
4. Measure (w/ laser rangefrndert and record the distance back to the flagged upstream transect. (Note
that, because of hazards and maneuvering problems, this distance may unavoidably differ from the
"intended transect spacing" that is set at 4 (or 10) times the wetted width in the near vicinity of the
furthest upstream transect (A).
5. Backsite the river gradient: While at the bank at Transect B, hold your Abney or clinometer at the same
level on your measuring rod that you used at the previous station when you set up the eye-level
flagging. Site back upstream at your flagging at Station A; read and record percent Slope on the field
form. Be careful, the clinometer reads both percent slope and degrees of the slope angle. Percent slope
is the scale on the right hand side as you look through most clinometers. If using an Abney Level,
insure that you are reading the scale marked " PERCENT."
6. Backsite the compass bearing: From the bank at Station B, site back with your compass to the flagging
you placed at Station A and record your compass bearing ("Azimuth"). It does not matter for these
measurements whether or not you adjust your compass bearings for magnetic declination, but it is
important that you are consistent in the use of magnetic (unadjusted) or true (adjusted) bearings
throughout all the measurements you make on a given reach. Write on the field form which type of
bearings you take. Also guard against recording "reciprocal" bearings (erroneous bearings 180 degrees
from what they should be). The best way to do this is to know where the primary (cardinal) directions
are in the field - north (0 degrees), east (90 degrees), south (180 degrees), and west (270 degrees) -- and
insure that your bearings "make sense."
7. Repeat step 1, setting your eye-level flagging at Transect B before floating down to a new downstream
transect. Then repeat steps 2 through 7.
Again adapting methods developed for
lake shorelines by Kaufmann and Whittier
(1997), identify the dominant and subdomi-
nant substrate present along a shoreline swath
20 meters long and 1 meter back from the
waterline. The substrate size class choices are
as shown in Table 6-5.
6.6.3 Large Woody
Debris
Methods for tallying large woody debris
(LWD) are adapted from those described by
Kaufmann and Robison (1998). This com-
ponent of the EMAP Physical Habitat proto-
col allows estimates of the number, size, and
total volume of large woody debris within the
river reach. LWD is defined here as woody
material with small end diameter of at least
30 cm (1ft) and length of at least 5 m (15 ft).
These size criteria are larger than those used
by Kaufmann and Robison (1998) in wade-
able streams because of the lesser role that
small wood plays in controlling velocity and
morphology of larger rivers.
The procedure for tallying LWD is pre-
sented in Table 6-6. The tally includes all
pieces of LWD that are at least partially in the
baseflow channel (Wetted Channel). Sepa-
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Table 6-5. Channel Margin Depth and Substrate Procedure.
If not already done, fill in the header information on page 1 of a Channel/Riparian Transect Form. Be
sure to indicate the letter designating the transect location.
Measure depth and observe bottom substrates within a 10m swath along the 20m of the channel
margin that is centered on each transect location.
Determine and record the depth and the dominant and subdominant substrate size class at 5
systematically-spaced locations estimated by eye within this 10m x 20m plot and 1m back from the
waterline. If the substrate particle is " artificial" (e.g. concrete or asphalt), choose the appropriate
size class, flag the observation and note that it is artificial in the comment space.
Code Size Class
RS Bedrock (Smooth)
RR Bedrock (Rough)
HP Hardpan
BL Boulders
CB Cobbles
GC Gravel (Coarse)
GF Gravel (Fine)
SA Sand
FN Fines
WD Wood
OT Other
Size Range (mm)
>4000
>4000
:>250 to 4000
>64 to 250
>16 to 64
> 2 to 16
>0.06 to 2
<0.06
Regardless of Size
Regardless of Size
Description
Smooth surface rock bigger than a car
Rough surface rock bigger than a car
Firm, consolidated fine substrate
Basketball to car size
Tennis ball to basketball size
Marble to tennis ball size
Ladybug to marble size
Smaller than ladybug size, but visible as
particles - gritty between fingers
Silt Clay Muck (not gritty between fingers)
Wood & other organic particles
Concrete, metal, tires, car bodies etc.
(describe in comments)
4.
On page 1 of the Channel/Riparian Transect Form, circle the appropriate shore and bottom substrate
type and record the depth measurements ("SONAR" or "POLE" columns).
5. Repeat Steps 1 through 4 at each new cross section transect.
rately tally wood that is presently dry but con-
tained within the "BankfuH" or active chan-
nel (flood channel up to bankroll stage). In-
clude wood that spans above the active
channel or spanning above the active chan-
nel with the "Dry but within Bankfull" cat-
egory. For each tally (Wetted Channel and
Dry but within Bankfull), the field form (Fig-
ure 6-4) provides 12 entry boxes for tallying
debris pieces visually estimated within three
length and four diameter class combinations.
Each LWD piece is tallied in only one box.
Woody debris is not tallied in the area be-
tween channel cross, sections, but the pres-
ence of large debris dams and accumulations
should be mapped and noted in the comments.
For each LWD piece, first visually esti-
mate its length and its large and small end di-
ameters in order to place it in one of the di-
ameter and length categories. The diameter
classes on the field form (Figure 6-4) refer to
the large end diameter. The diameter classes
are 0.3m to <0.6m, 0.6m to <0.8m, and 0.8m
to <1 .Om and >1 .Om. The length classes are
5m to < 15m, 15m to <30m, and >30m. Some-
times LWD is not cylindrical, so it has no clear
"diameter". In these cases visually estimate
what the diameter would be for a piece of
wood with circular cross section that would
have the same volume. When evaluating
length, include only the part of the LWD piece
that has a diameter greater than 0.3m (1 ft).
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Table 6-6. Procedure for Tallying Large Woody Debris.
Note: Tally pieces of large woody debris (LWD) within the 11 transects of the river reach at the same
time the shoreline measurements are being determined. Include all pieces whose large end is
located within the transect plot in the tally.
1. LWD in the active channel is tallied in 11 "plots" systematically spaced over the entire length of the
stream reach. These plots are each 20 m long in the upstream-downstream direction. They are
positioned along the chosen bank and extend from the shore in 10m towards mid-channel and then all
the way to the bankfull margin.
2. Tally all LWD pieces within the plot that are at least partially within the baseflow channel. Also tally
LWD that is dry but contained within the active channel. First, determine if a piece is large enough to
be classified as LWD (small end diameter 30 cm [1 ft.]; length 5 m [15 ft.])
3. For each piece of LWD, determine its diameter class based on the diameter of the large end (0.3 m to <
0.6 m, 0.6 m to <0.8 m, 0.8 m to <1.0 m, or >1.0 m), and the length class of the LWD pieces based on the
part of its length that has diameter 30 cm. Length classes are 5m to < 15m, 15m to <30m, or >30m.
• If the piece is not cylindrical, visually estimate what the diameter would be for a piece of wood
with circular cross section that would have the same volume.
• When estimating length, include only the part of the LWD piece that has a diameter greater than
0.3m (1ft.)
4. Place a tally mark hi the appropriate diameter °° length class tally box in the "WOOD All/Part in
WETTED .Channel1' section of the Channel/Riparian Transect Form.
5. Tally all shoreline LWD pieces along the littoral plot that are at least partially within or above
(bridging) the bankfull channel, but not in the wetted channel. For each piece, determine the diameter
class based on the diameter of the large end (0.3 m to < 0.6 m, 0.6 m to <0.8 m, 0.8 m to <1.0 m, or >1.0
m), and the length class based on the length of the piece that has diameter 30 cm. Length classes are 5m
to <15m, 15m to <30m, or >30m.
6. Place a tally mark for each piece in the appropriate diameter =» length class tally box in the "DRY BUT
ALL/PART IN Bankfull Channel" section of the Channel/Riparian Transect Form.
7. After all pieces within the segment have been tallied, write the total number of pieces for each diameter
oo length class in the small box at the lower right-hand corner of each tally box.
8. Repeat Steps 1 through 7 for the next river transect, using a new Channel/Riparian Transect Form.
Count each of the LWD pieces $s one tally en-
try and include the whole piece when assessing
dimensions, even if part of it is outside of the
bankfull channel. If you encounter massive,
complex debris jams, estimate their length,
width, and height Also estimate the diameter
and length of large "key" pieces and judge the
average diameter and length of the other pieces
making up the jam. Record this information in
the comments section of the form.
6.6.4 Bank Angle and
Channel Cross-Section
Morphology
Undercut, vertical, steep, and gradual bank
angles are visually estimated as defined on the
field form (Figure 6-4). Observations are made
from the wetted channel margin up 5 m (a
canoe's length) into the bankfull channel mar-
gin on the previously chosen side of the stream.
The channel dimensions to be measured
or estimated are the wetted width, mid-chan-
nel bar width, bankfull height and width, the
amount of incision, and the degree of chan-
nel constraint. These shall be assessed for the
whole channel (left and right banks) at each
of the 11 cross section transects. Each are re-
corded on the Channel/Riparian Transect
Form (Figure 6-4). The procedure for obtain-
ing bank angle and channel cross-section
morphology measurements is presented in
Table 6-7.
-------
Table 6-7. Procedure for Bank Angle and Channel Cross-Section.
Visually estimate the bank angle (undercut, vertical, steep, gradual), as defined on the field form.
Bank angle observations refer to the area from the wetted channel margin up 5 m (a canoe's length)
into the bankfull channel margin on the previously chosen side of the river. Circle the range within
.which the observed band angle falls on the "Bank CHARACTERISTIC" section of the Channel/
Riparian Transect Field Form. .
With a laser rangefinder at a cross-section transect, measure and record the wetted width value in the
"Wetted Width" field in the bank characteristics section of the field data form. Also determine the
bankfull channel width and the width of exposed mid-channel bars (if present) with the laser
rangefinder and surveyor's rod. Record these values in the "Bank CHARACTERISTIC" section of the
field data form.
To estimate bankfull height, hold the surveyor's rod vertical, with its base planted at the water s edge.
-Using the rod as a guide while examining both banks, estimate (by eye) the height of bankfull flow
above the present water level. Look for evidence on one or both banks such as:
• An obvious slope break that differentiates the channel from a relatively flat floodplain terrace
higher than the channel.
• A transition from exposed river sediments to terrestrial vegetation.
• A transition from sorted river sediments to unsorted terrestrial soils.
• Transition from bare rock to moss growth on rocks along the banks.
• Presence of drift material caught on overhanging vegetation.
• Transition from flood- and scour-tolerant vegetation to that which is relatively intolerant of these
conditions.
Hold the surveyor's rod vertical, with its base planted at the water's edge. Using the surveyor s rod as
a guide while examining both banks, estimate (by eye) the channel incision as the height up from the
water surface to the elevation of the first terrace of the valley floodplain (Note this is at or above the
bankfull channel height). Record this value in the "Incised Height" field of the Bank Characteristic
section on the field data form.
Repeat Steps 1 through 4 at each cross-section transect. Record data for each transect on a separate
field data form.
Wetted width refers to the width of the
channel as defined by the presence of free-
standing water; if greater than 15m, it can be
measured with the laser range finder. Mid-
channel bar width, the width of exposed mid-
channel gravel or sand bars in the channel, is
included within the wetted width, but is also
recorded separately. In channel cross-section
measurements, the wetted and active channel
boundaries are considered to include mid-
channel bars. Therefore, the wetted width
shall be measured as the distance between
wetted left and right banks. It is measured
across and over mid-channel bars and boul-
ders. If islands are present, treat them like bars,
but flag these measurements and indicate in
the comments that the "bar" is an island. If
you are unable to see across the full width of
the river when an island separates a side chan-
nel from the main channel, record the width
of the main channel, flag the observation, and
note in the comments section that the width
pertains only to the main channel.
Bankfull height and width shall be esti-
mated with the aid of the surveyor's rod and
laser range finder. The "bankfull" or "active"
channel is defined as the channel that is filled
by moderate sized flood events that fill the
channel to its flood banks. Measure bankfull
width over and across mid-channel bars.
Bankfull flows typically recur every 1 to 2
years and do not generally overtop the chan-
nel banks to inundate the valley floodplain.
They are believed to be largely responsible
for the observed channel dimensions in most
-------
rivers and streams. If the channel is not greatly
incised, bankfull channel height and the
amount of incision will be the same. How-
ever, if the channel is incised greatly, the
bankfull level will be below the level of the
first terrace of the valley floodplain, making
"Bankfull Height" smaller than "Incision"
(Figure 6-6). You will need to look for evi-
dence of recent flows (within about 1 year)
to distinguish bankfull and incision heights,
though recent flooding of extraordinary mag-
nitude may be misleading.
Estimating the level of bankfull flow dur-
ing baseflow conditions requires judgement
and practice; even then it remains somewhat
subjective. In many cases there is an obvious
slope break that differentiates the channel from
a relatively flat floodplain terrace higher than
the channel. Because scouring and inunda-
tion from bankfull flows are often frequent
enough to inhibit many types of terrestrial
vegetation, the bankfull channel may be evi-
dent by a transition from exposed river sedi-
ments and water-loving plants to upland ter-
A. Channal not "Incised"
Downcutting over Geologic Time
Stream - No recent incision.,,
Bankfull Level at Valley
Bottom
, First Terrace on
Valley Bottom
Second Terrace
Valley Fill
B. Channal "Incised"
Downcutting over Geologic Time
Recent incision: Bankfull
Level below first terrace of
Valley Bottom
First Terrace on
Valley Bottom
Second Terrace
Valley Fill
Figure 6-6. Schematic showing bankfull channel and incision for channels. (A) not recently incised
and (B) recently incised into valley bottom. Note level of bankfull stage relative to elevation of first
terrace on valley bottom (Stick figure included for scale).
-------
restrial vegetation. Similarly, it may be iden-
tified by noting where moss growth on rocks
along the banks has been removed by flood-
ing. The bankfull flow level may also be seen
by the presence of drift material caught on
overhanging vegetation.
As described in Table 6-7 and shown in
Figure 6-6, examine both banks and estimate
(by eye) the amount of channel incision from
the water surface to the elevation of the first
terrace of the valley floodplain. In cases where
the channel is^utting a valley sideslope and
has oversteepened and destabilized that slope,
the bare "cutbank" is not necessarily an indi-
cation of recent incision. Examine both banks
to make a more accurate determination of
channel downcutting. Finally, assess the de-
gree of river channel constraint by answering
the four questions on the form (Figure 6-5)
regarding the relationships among channel
incision, valley sideslope, and width of the
valley floodplain.
6.6.5 Canopy Cower
(Densiometer)
Riparian canopy cover over a river is
important not only for its role in moderating
water temperatures through shading, but also
as riparian wildlife habitat, and as an indica-
tor of conditions that control bank stability and
the potential for inputs of coarse and fine par-
ticulate organic material. Organic inputs from
riparian vegetation become food for river or-
ganisms and structure to create and maintain
complex channel habitat.
Vegetative cover over the river margin
shall be measured at the chosen bank at each
of the 11 transect locations (A-K). This mea-
surement employs the Convex Spherical
Densiometer, model B (Lemmon, 1957). The
densiometer must be taped exactly as shown
in Figure 6-7 to limit the number of square
grid intersections to 17. Densiometer readings
can range from 0 (no canopy cover) to 17
(maximum canopy cover). Four measurements
are obtained at each cross-section transect
(upriver, downriver, left, and right). Concen-
trate on the 17 points of grid intersection on
the densiometer. If the reflection of a tree or
high branch or leaf overlies any of the inter-
section points, that particular intersection is
counted as having cover. The measure to be
recorded on the form is the count (from 0 to
17) of all the intersections that have vegeta-
tion covering them. Therefore, a higher num-
ber indicates greater canopy extent and den-
sity. In making this measurement, it is
important that the densiometer be leveled us-
ing the bubble level (Figure 6-7).
The procedure for obtaining canopy
cover data is presented in Table 6-8. These
bank densiometer readings complement your
visual estimates of vegetation structure and
cover within the riparian zone (Section 6.6.6).
For each of the four directions, count the num-
ber of covered densiometer intersection points.
Record these counts in the "Canopy Density
@ Bank" section of the Channel/Riparian
Transect Form as shown in Figure 6-4.
6.6.6 Riparian
Vegetation Structure
The previous section (6.6.5) described
methods for quantifying the cover of canopy
over the river margin. .The following visual
estimation procedures, adapted from
Kaufmann and Robison (1998), are a semi-
quantitative evaluation of riparian vegetation
structure, the type and amount of different
types of riparian vegetation. These field char-
acterizations shall be used to supplement in-
terpretations of riparian vegetation from aerial
photos and satellite imagery. Together, they
-------
Tape
Bubble Leveled-
Flgure6-7. Schematic of modified convex spherical canopy densiometer (From Mulvey et al.,
1992). In this example, 10 of the 17 intersections show canopy cover, giving a densiometer reading
of 10. Note proper positioning with the bubble leveled and face reflected at the apex of the "V".
Table 6-8. Procedure for Canopy Cover
Measurements.
1. Take densiometer readings at a cross-section
transect while anchored or tied up at the river
margin.
2. Hold the densiometer 0.3 m (1 ft) above the
surface of the river. Holding the densiometer
level using the bubble level, move it in front of
you so your face is just below the apex of the
taped 'V.
3. At the channel margin measurement locations,
count the number of grid intersection points
within the "V" that are covered by either a tree,
a leaf, a high branch, or the bank itself.
4. Take 1 reading each facing upstream (UP),
downstream (DOWN), left bank (LEFT), and
right bank (RIGHT). Right and left banks are
defined with reference to an observer facing
downstream.
5. Record the UP, DOWN, LEFT, and RIGHT
values (0 to 17) in the "CANOPY COVER @
BANK" section of the Channel/Riparian
Transect Form.
6. Repeat Steps 1 through 5 at each cross-section
transect. Record data for each transect on a
separate field data form.
are used to evaluate the health and level of
disturbance of the river/riparian corridor. They
also indicate the present and future potential
for various types of organic inputs and shad-
ing. The cover and structure of riparian veg-
etation is estimated in three riparian layers
within 1 Om x 20m plots along the river shore-
line that are centered on the transect location
with boundaries estimated by eye. As em-
ployed by Allen-Gill (unpublished manu-
script), these plots shall be set back from the
channel so that they describe vegetation above
bankfull flow. As a result, gravel bars within
the bankfull channel are not included in the
vegetation plot (Figure 6-2).
Observations to assess riparian vegeta-
tion apply to the riparian area upstream 10
meters and downstream 10 meters from each
of the 11 cross-section stations (Figure 6-2).
They include the visible area from the river
-------
bankfull margin back a distance of 10m (30
ft) shoreward from both the left and right
banks, creating a 10m X 20m riparian plot on
each side of the river (Figure 6-2). The ripar-
ian plot dimensions are estimated, not mea-
sured. On steeply sloping channel margins,
the 10m X 20m plot boundaries are defined
as if they were projected down from an aerial
view. If the wetted channel is split by a mid-
channel bar, the bank and riparian measure-
ments shall be for each side of the channel,
not the bar. If an island obscures the far bank
of the main channel, assess riparian vegeta-
tion on the bank of the island.
Table 6-9 presents the procedure for
characterizing riparian vegetation structure
and composition. Figure 6-5 illustrates how
measurement data are recorded in the "Visual
Riparian Estimates" section of the field form.
Conceptually divide the riparian vegetation
into three layers: a CANOPY LAYER (>5m
high), an UNDERSTORY (0.5 to 5m high),
and a GROUND COVER layer (<0.5 high).
Note that several vegetation types (eg. grasses
or woody shrubs) can potentially occur in
more than one layer. Similarly note that some
things other than vegetation are possible en-
tries for the "Ground Cover" layer (eg. bar-
ren ground and duff, which includes fallen
leaves, needles and twigs),
Before estimating the areal coverage of
the vegetation layers, record the type of veg-
etation (Deciduous, Coniferous, Broadleaf
Evergreen Mixed, or None) in each of the
two taller layers (Canopy and Understory).
Consider the layer "Mixed" if more than 10%
of the areal coverage is made up of the alter-
nate vegetation type.
You will estimate the areal cover sepa-
rately in each of the three vegetation layers.
Note that the areal cover can be thought of as
the amount of shadow cast by a particular layer
alone when the sun is directly overhead. The
maximum cover in each layer is 100%, so the
sum of the areal covers for the combined three
layers could add up to 300%. The four entry
choices for areal cover within each of the three
vegetation layers are "0" (absent: zero cover),
"1" (sparse: <10%), "2" (moderate: 10-40%),
"3" (heavy: 40-75%), and "4" (very heavy:
>75%). These ranges of percentage areal
cover corresponding to each of these codes
are also shown on the Field Form. When rat-
ing vegetation cover types, mixtures of two
or more subdominant classes might all be
given sparse ("1") moderate ("2") or heavy
("3 ") ratings. One very heavy cover class with
no clear subdominant class might be rated' '4''
with all the remaining classes either moder-
ate ("2"), sparse ("1") or absent ("0"). Two
heavy classes with 40-75% cover can both
be rated "3".
As an additional assessment of the "old
growth" character of riparian zones, search
for the largest riparian tree visible on either
side of the river from the littoral-riparian sta-
tion. Identify if possible the species or the
taxonomic group of this tree and estimate its
height, diameter (Dbh), and distance from the
wetted river margin.
6.6.7 Fish Cover,
Algae, Aquatic
Macrophytes
This portion of the EMAP physical habi-
tat protocol is a visual estimation procedure
modified from methods developed for lake
shorelines (Kaufmann and Whittier 1997) and
for wadeable streams (Kaufmann and Robison
1998). The aim is to evaluate, semi-quantita-
tively, the type and amount of important types
of cover for fish and macroinvertebrates. Over
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Table 6-9. Procedure For Characterizing Riparian Vegetation Structure.
1. Anchor or tie up at the river margin at a cross-section transect; then make the following observations
to characterize riparian vegetation structure.
2. Estimate the distance from the shore to the riparian vegetation plot; record it just below the title
"Channel Constraint" on the field form.
3. Facing the left bank (left as you face downstream), estimate a distance of 10 m back into the riparian
vegetation, beginning at the bankfull channel margin. Estimate the cover and structure of riparian
vegetation in 3 riparian layers along the river shoreline within an estimated 10m x 20m plot centered
on the transect, and beginning at the bankfull river margin along the river shoreline.
|,
I:
• On steeply-sloping channel margins, estimate the distance into the riparian zone as if it were pro-
jected down from an aerial view.
4. Within this 10 m <» 20 m area, conceptually divide the riparian vegetation into three layers: a CANOPY
LAYER (>5m high), an UNDERSTORY (0.5 to 5 m high), and a GROUND COVER layer (<0.5 m high).
5. Within this 10 m °° 20 m area, determine the dominant vegetation type for the CANOPY LAYER
(vegetation > 5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None.
Consider the layer "Mixed" if more than 10% of the areal coverage is made up of the alternate
vegetation type. Indicate the appropriate vegetation type in the "Visual Riparian Estimates" section of
the Channel/Riparian Cross-section and Thalweg Profile Form.
6. Determine separately the areal cover class of large trees (> 0.3 m [1 ft] diameter at breast height [DBH])
and small trees (< 0.3 m DBH) within the canopy layer. Estimate areal cover as the amount of shadow
that would be cast by a particular layer alone if the sun were directly overhead. Record the appropriate
cover class on the field data form ("0"=absent: zero cover, "1 "=sparse: <10%, "2"=moderate: 10-40%,
"3"-heavy: 40-75%, or "4"=very heavy: >75%).
7. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the dominant
vegetation type for the undersfory layer as described in Step 5 for the canopy layer.
8. Determine the area! cover clasjs for woody shrubs and saplings separately from non-woody vegetation
within the understory, as described in Step 6 for the canopy layer.
9. Look at the GROUND COVER layer (vegetation < 0.5 m high). Determine the areal cover class for
woody shrubs and seedlings, non-woody vegetation, and the amount of bare ground present as
described in Step 6 for large canopy trees.
1 ' ;..•;; , , :'!
10. Repeat Steps 1 through 9 for the opposite bank.
11. Repeat Steps 1 through 10 for all cross-section transects, using a separate field data form for each
transect.
a defined length and distance from shore at 11
systematically spaced plot locations, crews shall
estimate by eye and by sounding the propor-
tional cover offish cover features and trophic
level indicators including large woody debris,
rootwads and snags, brush, undercut banks,
overhanging vegetation, rock ledges, aquatic
macrophytes, filamentous algae, and artificial
structures. Alone and in combination with other
metrics, this information is used to assess habi-
tat complexity, fish cover, and channel distur-
bance.
-------
The procedure to estimate the types and
amounts of fish cover is outlined in Table 6-
10. Data are recorded in the "Fish Cover/
Other" section of the Channel/Riparian
Transect Form as shown in Figure 6-5. Crews
will estimate the areal cover of all of the fish
cover and other listed features that are in the
water and on the banks within the 1 Om x 20m
plot (refer to Figure 6-2).
Observations to assess fish cover and
several other in-channel features apply to a
10 m x 20 m inundated area adjacent to the
selected bank extending 10 m out from the
channel margin, and then upstream 10m and
downstream 10m from each of the 11 transect
cross-sections (Figure 6-2). These plot dimen-
sions are estimated by eye. The ranges of per-
centage afeal cover corresponding to each of
these codes are the same as for riparian veg-
etation cover (Section 6.6.6) and are also
shown on the Field Form.
I Table 6-10. Procedure For Estimating Fish Cover.
P=.l.
2.
. -.. . .. • .... ,.. . ...
Stop at the designated shoreline at a cross- .
section transect and estimate a 10m distance ]
upstream and downstream (20m total length), '
and a 10m distance out from the banks to
define a 20m x 10m littoral plot.
Examine the water and the banks within the :
20m x 10m littoral plot for the following
features and types of fish cover: filamentous
algae, aquatic macrophytes, large woody ,
debris, brush and small woody debris, over- j
hanging vegetation, undercut banks, boulders,
and artificial structures.
For each cover type, estimate its areal cover ;
by eye and/or by sounding with a pole. "
:~z Record the appropriate cover class in the
"Fish Cover/Other" section of (he Channel/ ]
Riparian Transect Form ("0"=abserit: zero 1
cover, "l"=sparse: <10%, "2"=moderate: 10-"
., "3"=heavy: 40-75%, or "4"=very heavy: "
"''
-4. Repeat Steps 1 through 3 at each cross-
section transect, recording data from each
f-™™ transect on a separate field data form.
P; _.. _;, f • '.-„ .:.,._„.;•...'. . . '... -.' i,.~...
Filamentous algae pertains to long
streaming algae that often occur in slow mov-
ing waters. Aquatic macrophytes are water
loving plants in the river, including mosses,
that could provide cover for fish or
macroinvertebrates. If the river channel con-
tains live wetland grasses, include these as
macrophytes. Woody debris includes the
larger pieces of wood that can provide cover
and influence river morphology (i.e., those
pieces that would be included in the large
woody debris tally [Section 6.6.3]). Brush/
woody debris pertains to the smaller wood
that primarily affects cover but not morphol-
ogy. The entry for trees or brush within one
meter above the water surface is the amount
of brush, twigs, small debris etc. that is not in
the water but is close to the river and pro-
vides cover. Boulders are typically basketball
to car sized particles. Many streams contain
artificial structures designed for fish habitat
enhancement. Streams may also have in-chan-
nel structures discarded (e.g. cars or tires) or
purposefully placed for diversion, impound-
ment, channel stabilization, or other purposes.
Record the cover of these structures on the
form.
6.6.8 Human Influences
Field characterization of the presence
and proximity of various important types of
human activities, disturbances, and land use
in the river riparian area is adapted from meth-
ods developed by Kaufmann and Robison
(1998) for wadeable streams. This informa-
tion shall be used in combination with ripar-
ian and watershed landuse information from
aerial photos and satellite imagery to assess
the potential degree of disturbance of the
sample river reaches.'
For the left and right banks at each of
the 11 detailed Channel/Riparian Cross-Sec-
-------
tions, evaluate the presence/absence and the
proximity of 11 categories of human influ-
ences outlined in Table 6-11. Confine your
observations to the river and riparian area
within 10m upstream and 10m downstream
from the cross-section transect (Figure 6-2).
Four proximity classes are used: On the
riverbank within 10m upriver or downriver
of the cross-section transect, present within
the 10m x 20m riparian plot, present outside
of the riparian plot, and not present. Record
human influences on the Channel/Riparian
Transect Form (Figure 6-5).
You may mark "P" more than once for
the same human influence observed outside
of more than one riparian observation plot
(e.g. at both Transect D and E). The rule is
that you count human disturbance items as
often as you see them, BUT NOT IF you have
to site through a previously counted transect
or its 1 Ox20m riparian plot.
6.7 Summary of
Workflow
Table 6-12 lists the activities performed
at and between each transect for the physical
habitat characterization. The activities are
performed along the chosen river bank and
mid-channel (thalweg profile).
6.8 Equipment and
Supplies
Figure 6-8 lists the equipment and sup-
plies required to conduct all the activities de-
scribed for characterizing physical habitat.
This checklist is similar to the checklist pre-
sented in Appendix A, which is used at the
base location (Section 3) to ensure that all of
the required equipment is brought to the river.
Use this checklist to ensure that equipment
and supplies are organized and available at
,
i Table 6-1 1. Procedure for Estimating Human Influence.
- -
1 .
2.
3.
*
i-
p5.
I !. 6.
1
.
Stop at the designated shoreline at a cross-section transect, look toward the left bank (left when facing
downstream), and estimate a lOm distance upstream and downstream (26m total length). Also, ]
estimate a distance of 10m back into the riparian zone to define a riparian plot area.
Examine the channel, bank and riparian plot area adjacent to the defined river segment for the *
following human influences: (1) walls, dikes, revetments, riprap, and dams; (2) buildings; (3)
pavement (e.g., parking lot, foundation); (4) roads or railroads, (5) inlet or outlet pipes; (6) landfills or
trash (e.g., cans, bottles, trash heaps); "(7) parks or maintained lawns;~(8) row crops; (9) pastures, '
rangeland, or hay fields; (10) logging; and (11) mining (including gravel mining).
For each type of influence, determine if it is present and what its proximity is to the river and riparian
plot area. Consider human disturbance items as present if you can see them from the cross-section "',
trarisect"Do not 'include fliem'ir' you have 'to site'ftrough"' another transect or its 1pm o° 20rn riparian '
• """"''"' " "'" '" " "'""'""' ................. '''" ''
. . ... ..._ .. .. -,..
For each type of influence, record the appropriate proximity class in.the "Human Influence" part of the
"Visual Riparian Estimates" section of the Channel/Riparian Transect Form. Proximity classes are: ~\
• B("Bank") Present wimin the defined 20m river segment and located in the stream or on ;
................................ ............ ........... the wetted or bankfull bank. . _ ./ _. . ........ c_^ ........ - _ •' ...... • ..... ' ....... ; 'j
• C ("Close") Present within the 10 °° 20m riparian plot area, but above the bankfull level.
• P ("Present") ............ Pjesent,iibiut_obseryedi gutside^e^riparian plot' area. '• ' - '. ....... ^ }
• O(" Absent") Not present within or adjacent to the 20m river segment or the riparian plot area '
at the transect ................. ...,„'
Repeat Steps 1 through 4 for the opposite bank. •
Repeat Steps 1 through 5 for each cross-section transect, recording data for each transect on a separate
field form. . ' . . ...... ' ....... '-..-' :' *
-------
of Workflow - River Physical Habitat Characterization.
tgrA. At the chosen bank on first transect (farthest upstream):
:1.
Move boat in a "loop" within 10 x 20 meter littoral plot, measuring five littoral depths and probing '
substrate. . .......... ' " "- . .';:-.-. .; . -.'•-. '•'.'". .'• :•-.-.' . ' ''t
, Estimate dominant and subdominaiit littpral substrate, based on probing the five locations. :
Estimate areal cover of fish concealment features ;in 10 Jt20 meter littoral plot. : r
Tally LWD wimin or partially within the 10 x 20 meter littoral plot. '
Measure water conductivity and temperature. :
Do densiometer measurements at bank (facing upstream, downstream, left, right). ]
Choose bank angle class, estimate bankfull height, width and channel incision. (Note that width and •
incision estimates incorporate both left and right banks.). :
Tally LWD entirely outof water but at least partially within the bankfull channel. . . *
Estimate and record distance to riparian vegetation on the chosen bank.
Make visual riparian vegetation cover estimates for the 10 x 20 meter riparian plot on both sides of the '\
channel. (Note that riparian plot starts at bankfuli and continues back 10m away from the bankfull
' ' "' - ••• ~ ----••••• ••••
|11. Identify species, height, Dbh, and distance from riverbank of largest riparian tree within your vision. ;
S12. Make visual human disturbance tally. It has the same plot dimensions as the riparian vegetation — «
"""except if a disturbance item is observed in the river or within the bankfull channel, then the proximity j
^ code is "B", the closest rating. Disturbances within the plot get a rating of "C"; those visible beyond..,
""7 the plot are rated T". : \ '"
3. Siting clinometer level (0%) towards the near or far bank at the current transect, mark or remember an I
eyeTlevel point to which you will be siting when backsiting from the next downstream transect. >
4^ Get out far .enough from the bank so you can see downstream. Then use the'laser rahgefinder to site and •
record the distance to the intended position of the next downstream transect. :
LB.Thalweg Profile:
1 . As soon as you get out from the bank after doing transect activities, take the first of 20 thalweg depth *
5 measurements and substrate/snag probes using sonar and pole — also classify habitat type.
t 2. Estimate thalweg measurement distance increments by keeping track of boat lengths or channel- i
ft- -------- width distances traversed; each increment is I/ 10th (or l/20th) the distance between transects. ]
_3. At the 20th thalweg measurement location, you are one increment upstream of the next transect. 1
Backsite compass bearing mid-channel, then measure the distance and % slope back to your visual «
"mark" on the bank at the previous transect. ••'•?>
-. -:• ... - \ '^-\(; ,•••• -:••:•••••:. •'• ^,-..:::- •'.;•,-:•;. v.-.r-:V- '• •••••••••\
the Whole Process (for the remaining 1.0 transects and spaces in between). .. i
|L~.."
the river site in order to conduct the activities
efficiently.
6.9 Literature Cited
Bain, M.B., J.T. Finn, and H.E. Booke.
1985. Quantifying stream substrate
for habitat analysis studies. Nor.
Amer. Jour, of Fish. Man. 5:499-500.
Allen-Gil, S., M. Green, and D. H. Landers. Frissell, C.A., W.J. Liss, C.E. Warren,
Unpublished manuscript. Fish abun-
dance, instream habitat and the effects of
historical landuse practices in two large
alluvial rivers on the Olympic Penninsula,
Washington. U.S. EPA, WED.
and M.D. Hurley. 1986. A hierarchi-
cal framework for stream habitat
classification: viewing streams in a
watershed contest. Environ. Mgmt.
10(2): 199-214.
-------
Equipment and Supplies for Physical Habitat
Qty. Item
1
1
1
1
Irollea.
2
2pair
1
1
1
1
1 1 plus
extras
1 1 plus
extras
1 copy
1 set
Surveyor's telescoping leveling rod (round profile, fiberglass, metric scale, 7.5m
extended)
Clinometer (or Abney level) with percent and degree scales.
Convex spherical canopy densiometer (Lemmon Model B), modified with taped
"V
Bearing compass (Backpacking type)
Colored surveyor's plastic flagging (2 colors)
Covered clipboards (lightweight, with strap or lanyard to hang around neck)
Soft (#2) lead pencils (mechanical are acceptable)
Chest waders with felt-soled boots for safety and speed if waders are the
neoprene "stocking" type
Camera- waterproof 35mm with standard and wide angle lens
Film - 35mm color slide film, ASA 400 and 100
Fiberglass Tape and reel (50m metric) with good hand crank and handle
SONAR depth sounder - narrow beam (16 degrees)
Laser rangefinder - 400 ft. distance range - and clear waterproof bag
Channel/Riparian Transect Forms
Thalweg Profile Forms
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for physical
habitat characterization
Figure 6-8. Checklist of equipment and supplies for physical habitat
-------
Kaufmann, P.R. (ed.) 1993. Physical Habitat.
Pages 59-69 in R.M. Hughes, ed. Stream
Indicator and Design Workshop.. EPA/
600/R-93/138. U.S. Environmental Pro-
tection Agency, Corvallis, OR.
Kaufmann, P.R., P. Levine, E.G. Robison, C.
Seeliger, and D.V. Peck (1999). Quanti-
fying Physical Habitat in Wadeable
Streams. EPA 620/R-99/003. U.S. Envi-
ronmental Protection Agency, Washing-
ton, D.C. 102 pp + Appendices.
Kaufmann, P.R. and E.G. Robison. 1998.
Physical Habitat Assessment, pp 77-118
In: Lazorchak, J.L., Klemm, D.J., 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.
Kaufmann, P.R. and T.R. Whittier. (1997).
Habitat Assessment. Pages 5-1 to 5-25 In:
J.R. Baker, G.D. Merritt, and D.W.
Sutton (eds) Environmental Monitoring
and Assessment Program — Lakes Field
Operations Manual. EPA/600/R-97/003.
U.S. Environmental Protection Agency,
Las Vegas, NV, Corvallis, OR, and
Cincinnati, OH.
Lemmon, P.E. 1957. A new instrument for
measuring forest overstory density. J. For.
55(9):667-669.
Mulvey, M., L. Caton, and R. Hafele. 1992.
Oregon nonpoint source monitoring
protocols stream bioassessment field
manual: for macroinvertebrates and
habitat assessment. Oregon Department
of Environmental Quality Laboratory
Biomonitoring Section. 1712 S.W. llth
Ave. Portland, OR, 97201. 40 p.
Plafkin, J.L., M.T. Barbour, K.D. Porter,
S.K. Gross, R.M. Hughes. 1989. Rapid
bioassessment protocols for use in
streams and rivers: benthic
macroinvertebrates and fish. EPA/440/4-
89/001. U.S. Environmental Protection
Agency, Assessment and Watershed
Protection Division, Washington, DC.
Platts, W.S., W.F. Megahan, and G.W.
Minshall. 1983. Methods for evaluating
stream, riparian, and biotic conditions.
USDA For. Serv., Gen. Tech. Rep. INT-
183, 71 p.
Robison, E.G. and R.L. Beschta. 1990.
Characteristics of coarse woody, debris
for several coastal streams of'southeast
Alaska, USA. 47(9): 1684-1693.
Robison, E.G. and P.R. Kaufmann. 1994.
Evaluating two objective techniques to
define pools in small streams, pgs 659-
668, In R.A. Marston and V.A.
Hasfurther (eds.) Effects of Human
Induced changes on hydrologic systems.
Summer Symposium proceedings, Ameri-
can Water Resources Association,. June
26-29, 1994, Jackson Hole, WY. 1182
pp.
Stack, BLR. 1989. Factors influencing pool
morphology in Oregon coastal streams.
M.S. Thesis, Oregon State University.
109 p.
-------
-------
Section 7
Periphyton
Brian H. Hill1 and Alan T. Herlihy2
Periphyton are algae, fungi, bacteria,
protozoa, and associated organic matter asso-
ciated with channel substrates. Periphyton are
useful indicators of environmental condition
because they respond rapidly and are sensi-
tive to a number of anthropogenic distur-
bances, including habitat destruction, con-
tamination by nutrients, metals, herbicides,
hydrocarbons, and acidification.
Periphyton samples are collected at the
near-shore shallows when stopped at each of
the cross-section transects (transects "A"
through "K") established within the sampling
reach (Section 4). Periphyton samples are
collected at each transect at the same time as
sediment samples (Section 8) and benthic
macroinvertebrate samples (Section 9). One
composite "index" sample of periphyton is
'U.S. EPA, National Exposure Research Laboratory, Eco-
logical Exposure Research Division, 26 W. Martin L. King
Dr., Cincinnati, OH 45268
Department of Fisheries and Wildlife, Oregon State
Univeristy, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR
97333
prepared for each river site. At the comple-
tion of the day's sampling activities, but be-
fore leaving the river, four types of labora-
tory samples are prepared from the composite
periphyton sample.
7.1 Sample Collection
The general scheme for collecting per-
iphyton samples from the sampling reach at
each river is illustrated in Figure 7-1. At each
transect, samples are collected from the shore-
line assigned during the layout of the reach
(Section 4). The substrate selected for sam-
pling should be collected from a depth no
deeper than can be reached by submerging
your arm to mid-bicep depth. If a sample can-
not be collected because the location is too
deep, skip the transect. The procedure for
collecting samples and preparing a compos-
ite sample is presented in Table 7-1. One
sample is collected from each of the transects
and composited in one bottle. The volume of
the sample is recorded on the Sample Collec-
tion Form as shown in Figure 7-2.
-------
K J
Cross Section Transects (A to K)
Transect Samples (11 Total)
Composite Transect Samples
ID/Enumeration Sample
Acid/Alkaline
Phos
50-mL aliquot
Preserve with 10% formalin
50-mL aliquot
Store at -20 °C
Chlorophyll Sample
o
Biomass Sample
Filter 25-mL aliquot
lass-fiber filter)
re at -20 °C
Filter 25-mL aliquot (glass-fiber filter)
Store filter at -20 °C
Figure 7-1. Index sampling design for periphyton.
7.2 PrGDaratiOn Of ^ex samPl£s: an ID/enumeration sample (to
. . ^ - determine taxonomic composition and rela-
L3DOratOry oampieS tiveabundances),achlorophyllsample,abio-
Four different types of laboratory mass sample (for ash-free dry mass [AFDM]),
samples are prepared from the composite in- and an acid/alkaline phosphatase activity
-------
= TabIe 7-1. Procedure for Collecting Composite Index Samples of Periphyton.
;: 1, Starting with Transect "A", collect a single sample from the assigned shoreline using'the procedure
^ below.
•- (a) Collect a sample of substrate: (rock or wood) that is small enough (< 15 cm diameter) and can be
r easily removed from the river. Place the substrate in a plastic funnel which drains into a 500-mL
'"'"'™ plastic bottle with volume graduations marked on it.
; V (b) Use the area delimiter to define a 12-cm2 area on the upper surface of the substrate. Dislodge
e attached periphyton from the substrate within the delimiter into the funnel by brushing with a
;rf stiff-bristled toothbrush for 30 seconds. Take care to ensure that the upper surface of the sub-
; : strate is the surface that is being scrubbed, and that the entire surface within the delimiter is
pe—-= - -scrubbed.
EL (c) Fill a wash bottle with river water. Using a minimal volume of water from this bottle, wash the
B dislodged periphyton from the funnel into the 500-mL bottle.
; If no coarse sediment (cobbles or larger) are present:
''-•'-: (d) Use the area delimiter to confine a 12-cm2 area of soft sediments.
S - (e) Vacuum the top 1 cm of sediments from within the delimited area into a de-tipped 60-mL
i; syringe.
; (f) Empty the syringe into the same 500-mL plastic bottle as above.
" 2. Repeat Step 1 for transects "B" through "K". Place the sample collected at each sampling site into the
HE single 500-mL bottle to produce the composite index sample.
3. After samples have been collected from all 1-1 transects, thoroughly mix the 500-mL bottle regardless
of substrate type. Record the total estimated volume of the composite sample in the periphyton
section of the Sample Collection Form.
(APA) sample. All the sample containers re-
quired for an individual river should be sealed
in plastic bags until use (see Section 3) to
avoid external sources of contamination (e.g.,
dust, dirt, or mud) that are present at river
shorelines.
A set of completed periphyton sample
labels is shown in Figure 7-3. All labels in a
set have the same sample ID number. Circle.
the habitat type of the composite index sample
and the appropriate type of sample (chloro-
phyll, biomass, etc.) on each label. Attach
completed labels to the appropriate contain-
ers and cover with clear tape. When attach-
ing the completed labels, avoid covering any
volume graduations and markings on the con-
tainer.
7.2.1 ID/Enumeration
Sample
Prepare the ID/Enumeration sample as
a 50-mL aliquot from the composite index
sample, following the procedure presented in
Table 7-2. Preserve each sample with 2 mL
of 10% formalin. Record the ID number
(barcode) from the container label and the total
volume of the sample in the appropriate fields
on the Sample Collection Form as shown in
Figure 7-2. Store the preserved samples up-
right in a container cpntaining absorbent ma-
terial, according to the guidelines provided for
handling formalin-preserved samples.
7.2.2 Chlorophyll
Sample
Prepare the chlorophyll sample by fil-
tering a 25-mL aliquot of the composite in-
dex sample through a glass fiber filter (0.45
m nominal pore size). The procedure for pre-
paring chlorophyll samples is presented in
Table 7-3. Chlorophyll can degrade rapidly
when exposed to bright light. If possible, pre-
pare the samples in subdued light (or shade),
filtering as quickly as possible after collec-
-------
Reviewed by (initial)
SAMPLE COLLECTION FORM - RIVERS
SITE NAME: BEAVER RlV£/f
DATE: g IS 1 98 VISIT DO J3 1 D2 D3
SITE ID: ORRV
5 -
TEAM ID (X):
D2 D3 D4 H5 D6 D7 D8
COMPOSITE BENTHOS SAMPLES
BANK SAMPLED (LOOKING DOWNSTREAM): ]S LEFT
RIGHT
SAMPLED
(BARCODE)
HABITAT
(CIRCLE ONE)
NO.
OF JARS
FLAG
S = SHORE
M = MID-CHANNEL
D = DRIFT
COMMENTS
24-56 I Z
RlfiKTE4Mr-S4MPL£D AT LAUtJCff
HO SHORE KICK AT -nf?AwsEcrs
=> TOO DEEP
STATION (CIRCLE IF DONE)
SHORE KICK
MID-CHANNEL
s
M
s
M
M
s
M
(£>
M
S
M
DRIFT
NET
TRANSECT
TIME (24-HR CLOCK)
START
VELOCITY
(FT/SEC)
XX.XX
IA DB DC Do DE DF DG DH Di DJ DK
3. 0
COMPOSITE PERIPHYTON SAMPLES
SAMPLE ID (BARCODE)*
2. QfO 1 3
COMPOSITE VOLUME xxxx mL *•
32.5
ASSEMBLAGE ID
(50-mLTUBE)
CHLOROPHYLL
(GF/F FILTER
BIOMASS
(TARED FILTER)
APA SAMPLE
(50-mLTUBE)
SUB. SAMPLE VOL.
xx ml
VOL. FILTERED
xxxxmL
FILTER NO.
VOL. FILTERED
xxxxmL
SUB. SAMPLE VOL.
xxxx mL
2.5
£-0
COMMENTS:
Flag codes; K- Sample no* collected; U» Suspect sample; F1.F2, etc. - misc. flag assigned by field crew. Explain all flags in Comments sections.
Rav. OS/29/98 (jvscmb.SS) SAMPLE COLLECTION FORM - RIVERS -1
Figure 7-2. Sample Collection Form (pagel) showing data recorded for periphyton samples.
-------
PERIPHYTON
^PA) BIOMASS CHLA ID
SITE ID: ORRV j? £ - 3. 9 9.
DATE: JL/5/J.8
HABITAT: POOL <§IFFLE/RUN^
SUBSAMPLE VOLUME: 50 mL
COMPOSITE VOLUME:32£.mL
229000
PERIPHYTON
APA <@OMAS§> CHLA ID
SITE IDTSHHV 3 JL - 3. 5L 3.
DATE: JL/ 5/_98 _
HABITAT: POOL
SUBSAMPLE VOLUME: 25" mL
COMPOSITE VOLUME:£2£.mL
229000
PERIPHYTON
APA BIOMASS (gR
SITE ID: ORRV j?
DATE: JL/ 5/98
HABITAT: POOL
SUBSAMPLE VOLUME:^£_mL
COMPOSITE VOLUME:32£.mL
229000
PERIPHYTON
APA BIOMASS CHLA (5)
SITE ID: ORRV J? £ - 9_ 9. 37
DATE: JL / jS / 98 _
HABITAT: POOL (RIFFLE/RUN)
SUBSAMPLE VOLUME: 5"O mL
COMPOSITE VOLUME:
229000
Figure 7-3. Completed set of periphyton sample labels.
I: Table7-2. Preparation of ID/Enumeration Samples for Periphyton.
JT1. Prepare abarcoded sample label and circle the sample type ("ID") on the label. Record the volume of
|~= the subample (typically 50 mL) and the volume of the composite index sample on the label. Attach the
C completed label to a 50-mL centrifuge tube; avoid covering the volume graduations and markings.
JT~" Cover the label completely with a clear tape strip.
| 2. Record the sample ID number (barcode) of the label and the total volume of the composite index
!„, " sample on the form.
£v3. Rinse a 60-mL syringe with deionized water.
K;-"'~ - ••..••. • • • : .
I 4. Thoroughly mix the bottle containing the composite sample.
F • . ' . • .
:l . - • • . . • • ' :
fe 5. Withdraw 50 mL of the mixed sample into the syringe. Right after mixing, place the contents of the
JET syringe sample into the labeled 50-mL centrifuge tube.
|; 6r Wearing gloves and safety glasses, use a syringe or bulb pipette to add 2 mL of 10% formalin solution
I to the tube. Cap the tube tightly and seal with plastic electrical tape. Shake gently to distribute
£•- preservative.
7. Record the volume of the sample in the centrifuge tube (excluding the volume of preservative) in the
"Assemblage ID Subsample Vol." field of the Sample Collection Form.
-------
Table 7-3. Procedure for Preparing Chlorophyll Samples for Periphyton.
1.
2.
3.
4.
5.
6.
Using clean forceps, place a glass fiber filter on the filter holder. Use a small amount of deionized water
from a wash bottle to help settle the filter properly. Attach the filter funnel to the filter holder and filter
chamber, then attach the hand vacuum pump to the chamber.
Rinse the sides of the filter funnel and the filter with a small volume of deionized water.
Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water.
Mix the composite sample bottle thoroughly.
Measure 25 mL (±1 mL) of sample into the graduated cylinder.
• NOTE: For a composite sample containing fine sediment, (e.g., the "DEPOSITIONAL" sample),
allow grit to settle for 10 - 20 seconds before pouring the sample into the graduated cylinder.
Pour the 25-mL aliquot into the filter funnel, replace the cap, and pull the sample through the filter
using the hand pump. NOTE: Vacuum pressure from the pump should not exceed 15 psi to avoid
rupture of fragile algal cells.
• If 25 mL of sample will not pass through the filter, discard the filter and rinse the chamber
thoroughly with deionized water. Collect a new sample using a smaller volume of sample, measured
to ±1 mL. Be sure to record the actual volume sampled on the sample label and the Sample
Collection Form.
Remove both plugs from the filtration chamber and pour out the filtered water in the chamber.
Remove the filter funnel from the filter holder. Remove the filter from the holder with clean forceps.
Avoid touching the colored portion of the filter. Fold the filter in half, with the colored sample
(filtrate) side folded in on itself. Wrap the folded filter in a small piece of aluminum foil.
Complete a periphyton sample label for chlorophyll, including the volume filtered, and attach it to
the foil. Cover the label completely with a strip of clear tape. Place the foil packet into a self-sealing
plastic bag.
Record the sample ID number (barcode) of the label and the total volume of the composite index
sample on the form. Record the volume filtered in the "Chlorophyll" field on the Sample Collection
Form. Double check that the volume recorded on the collection form matches the total volume
recorded on the sample label.
10. Place the plastic bag containing the filter into a portable freezer or between two sealed plastic bags of
ice or frozen gel packs in a cooler.
11. Rinse the filter funnel, filter holder, filter chamber, and graduated cylinder thoroughly with deionized
water.
7.
8.
9.
tion to minimize degradation. The filtration
apparatus is illustrated in Figure 7-4. Rinse
the filtration chamber with deionized water
each day before use at the base site and then
seal in a plastic bag until use at the stream
(see Section 3). Keep the glass fiber filters in
a dispenser inside a sealed plastic bag until
use.
It is important to measure the volume of
the sample being filtered accurately (±1 mL)
with a graduated cylinder. During filtration,
do no exceed 15 pounds per square inch (psi)
to avoid rupturing cells. If the vacuum pres-
sure exceeds 15 psi, prepare anew sample. If
the filter clogs completely before all the
sample in the chamber has been filtered, dis-
card the sample and filter, and prepare a new
sample using a smaller volume of sample.
After filtering each sample, wrap the fil-
ter in aluminum foil. Complete a sample la-
bel (Figure 7-3) and check it to ensure that all
written information is complete and legible.
Affix the label to the foil packet and cover it
completely with a strip of clear tape. Record
the barcode assigned to the sample on the
Sample Collection Form (Figure 7-2). Make
™|,'!:V "ijK .-• v
-------
Hand
Vacuum Pump
Clear
Plastic
Tubing
Figure 7-4. Filtration apparatus for preparing chlorophyll and biomass subsamples for periphyton.
Modified from Chaloud et al. (1989).
sure the volume recorded on each sample la-
bel matches the corresponding volume re-
corded on the Sample Collection Form.
Record a flag and provide comments on the
Sample Collection Form if there are any prob-
lems in collecting the sample or if conditions
occur that may affect sample integrity. Store
each foil packet in a self-sealing plastic bag.
Store the sample frozen until shipment to the
laboratory (Section 3).
7.2.3 Biomass Sample
Prepare the biomass sample from a 25-
mL aliquot of the composite index sample.
As with the chlorophyll sample, it is impor-
tant to measure the volume to be filtered ac-
curately (±1 mL).
After filtering each sample, complete a
sample label as shown in Figure 7-3. Check
each sample label to ensure that all written
information is complete and legible. Affix the
label to the filter container and cover it com-
pletely with clear tape. Record the bar code
assigned to the sample, the container num-
ber, and the volume filtered on the Sample
Collection Form as shown in Figure 7-2.
Make sure the information recorded on each
sample label and filters container matches the
corresponding values recorded on the Sample
Collection Form. Record a flag and provide
comments on the Sample Collection Form if
there are any problems in collecting the
sample or if conditions occur that may affect
sample integrity. Store each labeled filter con-
-------
Table7-4. Procedure For Preparing Biomass Samples For Periphyton.
1. Using clean forceps, remove a glass-fiber filter and place it on the filter holder. Use a small amount of
deionized water from a wash bottle to help settle the filter properly. Attach the filter funnel to the filter
holder and filter chamber, then attach the hand vacuum pump to the chamber.
Rinse the filter chamber and filter with a small volume of deionized water.
Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water.
Mix the composite sample bottle thoroughly.
Measure 25 mL (±1 mL) of composite sample into the graduated cylinder.
• NOTE: For a composite sample containing fine sediment, allow grit to settle for 10 - 20 seconds
before pouring the sample into the graduated cylinder.
Pour the 25-mL aliquot into filter funnel, replace the cap, and pull the sample through the filter using
the hand pump. NOTE: Filtration pressure should not exceed 15 psi to avoid rapture of fragile algal
2.
3.
4.
5.
6.
7.
8.
9.
If 25 mL of sample will not pass through the filter, discard the filter and rinse the chamber
thoroughly with deionized water. Collect a new sample using a smaller volume of sample, measured
to ±l mL- Be sure to record the actual volume filtered on the sample label and the Sample Collection
Form.
Remove both plugs from the filtration chamber and pour out the filtered water in the chamber.
Remove the filter funnel from the filter holder. Remove the filter from the holder with clean forceps
Avoid touching the colored sample portion of the filter.
Complete a periphyton sample label for biomass, including the volume filtered, and attach it to the
foil. Cover the label completely with a strip of clear tape. Place the foil packet into a self-sealing
plastic bag.
Record the sample ID number (barcode) of the label and the total volume of the composite sample on
the form. Record the volume filtered in the "Biomass" portion on the Sample Collection Form. Double
check that the volume recorded on the collection form matches the total volume recorded on the
sample label.
10. Place the labeled filter container into a cooler containing two sealed plastic bags of ice.
1 1 . Rinse the filter funnel, filter holder, filter chamber, and graduated cylinder thoroughly with deionized
water.
tainer frozen until shipment to the laboratory
(Sections).
7.2.4 Acid/Alkaline
Phosphatase Activity
Sample
The Acid/Alkaline phosphatase activity
(APA) sample is prepared from a 50-mL
subsample of the composite index sample.
Table 7-5 presents the procedure for prepar-
ing APA samples. No field treatment (i.e., fil-
tration, preservation) of the APA sample is
necessary. Complete a label for each sample
as shown in Figure 7-3 and affix it to a 50-
mL centrifuge tube. Record the ID number
(barcode), and the volume of the subsample
on the Sample Collection Form (Figure 7-2).
Check to ensure that the information recorded
on the Sample Collection Form matches the
corresponding information recorded on the
sample label. Store APA samples frozen until
shipment to the laboratory (Section 3).
7.3 Equipment and
Supplies
Figure 7-5 is a checklist of equipment
and supplies required to conduct periphyton
sample collection and processing activities.
This checklist is similar to the checklist pre-
sented in Appendix A, which is used at the
base location (Section 3) to ensure thatall of
-------
|Table7-5. Procedure for Preparing Add Alkaline Phosphatase Activity Samples for Periphyton.
i£i
Prepare a barcoded sample label. Circle the sample type ("APA") and the habitat type ("Riffle/
Run" or "Pool") on the label. Record the volume of the sample (typically 50 mL) and the
volume of the composite index sample on the label. Attach the completed label to a 50-mL
centrifuge tube; avoid covering the volume graduations and markings. Cover the label
completely with a clear tape strip. ..".,.
Rmse a 60-mL syringe with deionized water.
Thoroughly mix the bottle containing the composite sample.
Withdraw 50 mLof the mixed sample into the syringe. Place the contents of the syringe sample
into the labeled 50-mL centrifuge tube. Cap the tube tightly and seal with plastic electrical
tape. '-''"•'.'''- ' : .• . " '''"''. ' -'-•.-
Record the sample ED number (barcode) of the label and the total volume of the composite
index sample on the form.
Record the volume of the sample in the centrifuge tube in the "APA Sample" field of the
Sample Collection Form.
Equipment and Supplies for Periphyton
Qty. Item
1
1
1
1
1
1
1
4
1 box
1 pair
1
1
1
2
2
4mL
1
1 pair
1 pair
1 set
1
1 pkg.
1
1 copy
1 set
Large funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent at 90° angle
1 -L wash bottle for stream water
1-L wash bottle containing deionized water
500-mL plastic bottles for the composite sample.
60 mL plastic syringe with 3/8" hole bored into the end
50-mL screw-top centrifuge tubes (or similar sample vials)
Glass-fiber filters for chlorophyll and biomass samples
Forceps for filter handling.
25-mL or 50-mL graduated cylinder
Filtration unit, including filter funnel, cap, filter holder, and receiving chamber
Hand-operated vacuum purnp and clear plastic tubing
Aluminum foil squares (3" x 6")
Self-sealing plastic bags for chlorophyll samples
10% formalin solution for ID/Enumeration samples
Small syringe or bulb pipette for dispensing formalin
Chemical-resistant gloves for handling formalin
Safety glasses for use when handling formalin
Sample labels (4 per set) with the same barcode ID number
Sample Collection Form for river
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Clear tape strips for covering labels
Cooler with bags of ice to store frozen samples
Field operations and method manual
Laminated sheets of procedure tables and/or quick reference guides for periphyton
Figure 7-5. Checklist of equipment and supplies for periphyton
-------
the required equipment is brought to the river.
Use this checklist to ensure that equipment
and supplies are organized and available at
the river site in order to conduct the activities
efficiently.
7.4 Literature Cited
Handbook of Methods for Acid Deposi-
tion Studies: Field Methods for Surface
Water Chemistry. EPA 600/4-89-020.
U.S. Environmental Protection Agency,
Washington, D.C.
Chaloud, D. J., J.M. Nicholson, B.P. Baldigo,
C.A. Hagley, and D.W. Sutton. 1989.
-------
Section 8
Sediment Community Metabolism
Brian H. Hill and Alan T. Herlihy
This section describes procedures to
collect a composite sediment sample from the
sampling reach. Sediment samples are col-
lected from each transect at the same time as
periphyton samples (Section 7) and benthic
macroinvertebrate samples (Section 9). At
each river, a composite "index" sample of sedi-
ment is prepared and used in the determina-
tion of sediment community metabolism.
The method outlined here for determin-
ing sediment community metabolism is de-
signed for headwater to mid-order streams,
and has been adapted for larger rivers or lakes.
The method measures changes in dissolved
oxygen (DO) concentrations of the overlying
water within microcosms containing small
amounts (ca. 10 mL) of sediments as a means
of assessing benthic microbial community
activity. Sediments are collected from depo-
sitional habitats along the study reach. Fol-
'U.S. EPA, National Exposure Research Laboratory, Eco-
logical Exposure Research Division, 26 W. Martin L. King
Dr., Cincinnati, OH 45268.
Department of Fisheries and Wildlife, Oregon State Uni-
versity, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
lowing incubation, the DO is re-measured and
the sediments are saved for ash-free dry mass
(AFDM) analysis. Respiration rate, estimated
as the change in DO concentration per hour
within each microcosm, is adjusted for
AFDM, yielding a measure of community
respiration per gram of AFDM. Organic car-
bon turnover time can be calculated from the
empirical relationship between the organic
carbon content of the sediment (estimated as
0.5 oo AFDM) and oxygen consumption.
8.1 Sample Collection
Table 8-1 describes the procedure for
collecting the composite sediment sample.
Collect sediment from depositional areas (e.g.,
pools, eddies, and backwaters) located at or
near each of the cross-section transects within
the sampling reach. If soft sediments are
scarce, collect them from wherever you can
within the sampling reach. At each sampling
point, use a small plastic scoop to collect the
top 2 cm ( 1 inch) of soft surface sediment.
Combine sediments from different sampling
points into a single jar or self-sealing plastic
bag to prepare a single composite index sample
-------
Table8-l. Sediment Collection Procedure.
1. At the first cross-section, locate a depositional
habitat (a pool, eddy, or backwater).
• If soft sediments are scarce, collect them
wherever you can within the reach
2. Use a plastic scoop to collect a sample of
surficial sediment (top 2 cm). Remove any
visible organisms from the sediment. Place the
sample in aplastic jar with volume graduations
labeled "SEDIMENT SAMPLE":
• Approximately 250 mL of sediment (~ 30
mL per transect) is required for sediment
metabolism.
3. Repeat Steps 1 through 2 for Transects "B"
through "K".
for the river reach. A composite sample vol-
ume of 250-mL is sufficient to prepare sedi-
ment metabolism samples.
8.2 Determining
Sediment Respiration
The procedure to measure sediment res-
piration in presented in Table 8-2. A dissolved
oxygen meter, equipped with a biological
oxygen demand (BOD) probe and stirrer, is
used for the determination of respiration rates.
This may or may not be the same meter used
Table 8-2. Procedure To Measure Sediment Respiration.
1.
i 2.
3.
I *
l- 5.
S 6.
I :
=
L 7.
.
I:
i
I 8.
\ 9.
f,-,
L, ia
Inspect the probe of the dissolved oxygen meter for outward signs of fouling and for an intact
membrane. Do not touch the electrodes inside the probe with any object. Always keep the probe moist
by keeping it inside its calibration chamber. Check the batteries and electronic functions of the meter as
described in the meter's operating manual.
Calibrate the oxygen probe in water-saturated air as described in the operating manual. Allow at least 15
minutes for the probe to equilibrate before attempting to calibrate.
• NOTE: Try to perform the calibration as close to river temperature as possible (not air temperature)
by using river water to fill the calibration chamber prior to equilibration.
• NOTE: For doing the elevation correction, the elevation of the sample site is provided on the site
information sheet in the dossier for the site^ Alternatively, obtain the elevation from a topographic
map. . ' ' !' '. -.-•••
Prepare a set of five sediment metabolism sample labels. Note that each label will have a different sample
JD number (barcode). Attach each completed label to a 50-mL screw-cap centrifuge tube.
• NOTE: Avoid covering volume gradations on the tube with the label. Cover each label with a strip
of clear tape.
Fill a small insulated cooler full with river water. Measure the dissolved oxygen and temperature of the
water in the cooler. Record the values in the "Initial O2" and "Initial Incubation Temp." fields in the
metabpHsm section of the .Field Measurement Form. ' ___ _ _ r -
Thoroughly mix the composite sediment sample. Use a small plastic spoon to transfer 10 mL of sediment
from the composite sample container to each of the five labeled tubes.
Fill each tube to the top (no head space) with river water from the cooler and seal the tube. Fill a
centrifuge tube labeled "BLANK" with river water from the cooler and seal. This tube serves as a
control for changes in ambient conditions during the incubation period.
Place the six tubes in a 1 -L plastic beaker and place the beaker inside the cooler. Record the start time in
the "Incubation Time" area of the Field Measurement Form. Close the cooler and incubate the sediment
samples for 2 hours.
After incubation, re-calibrate the oxygen probe (i.e., the meter was turned off or you have moved to a
different elevation during the incubation) before the end of the incubation period.
At the end of the incubation period, record the end time in the "Incubation Time" area of the Field
Measurement Form. Measure the DO in each tube, including the blank. Record the sample ID number of
each tube and its measured DO concentration on the Field Measurement Form.
Decant the overlying water from each labeled tube, retaining the sediment. Tightly seal each tube and
place in a cooler with bags of ice as soon as possible. Keep the samples frozen until they can be
shipped. Discard the water from the "BLANK" tube.
-------
to determine in situ dissolved oxygen concen-
tration (Section 5). If a separate meter is used
to measure sediment respiration, check the
probe membrane and the meter's batteries and
electronics according to the instrument's op-
erating manual (see Sections 3 and 5, also).
Calibrate the meter as directed in the
instrument's operating manual.
A small cooler filled with stream water
is used as an incubation chamber. The initial
dissolved oxygen concentration and tempera-
ture of the water in the cooler are measured
and recorded on the Field Measurement Form
as shown in Figure 8-1. This concentration is
assumed to be the initial concentration of all
subsamples. Five sediment subsamples (10-
mL ±1 mL) are prepared from the composite
sediment sample. A set of completed sample
labels for these subsamples is shown in Fig-
ure 8-2. A 10-mL subsample of water from
the incubation cooler is used as a control for
changes in ambient conditions during the
incubation. The subsamples are incubated in
the cooler for 2 hours. After the incubation,
the final DO concentration of each tube is de-
termined and recorded on the Field Measure-
ment Form (Figure 8-1). The sediment in each
tube is retained and stored frozen until it .can
be shipped to the laboratory,(Section-3) to
determine the AFDM.
8.3 Equipment and
Supplies
Figure 8-3 is a checklist of equipment
and supplies required to conduct sediment
sampling and to determine sediment commu-
nity respiration. This checklist is similar to.the
checklist presented in Appendix A, which is
used at the base location (Section 3) to en-
sure that all of the required equipment is
brought to the river.-Use this checklist to en-
sure that equipment and supplies are orga-
nized and available at the river site in order to
conduct the activities efficiently.
-------
Reviewed by (initial)
FIELD MEASUREMENT FORM - STREAMS/RIVERS
SITE NAME: BEAVE"R R»V£j?
DATE: g I 5 I 98 VISIT DO Jfl 1 D2 D3
SITEID: ORRV
TEAM ID (X):
D2 D3 D4 D5 D6 D7 D8
WEATHER CONDITIONS (X)
CLOUD COVER
D 5-25%
D 25-50% .
50-75%
>75%
PRECIPITATION
. NONE
D LIGHT
D MODERATE
D HEAVY
PREVIOUS PRECIPITATION (24 H)
NONE
LIGHT
n MODERATE Q HEAVY
AIR TEMPERATURE XX
°C
IN SITU MEASUREMENTS
STATION ID:
K
Assume X-site unless marked
QCCS COND U.S/CM XXXX
75-
Ar
STREAM/RIVER COND U.S/CM XXXX
TRAK/5ECT *£"
STREAM/RIVER DO MG/LXX.X
STREAM/RIVER TEMP °C XX.X
IT.
Ar TRANSECT
•FISH TISSUE SAMPLES - SECONDARY SAMPLE (where available; 5 individuals)
,
(MOIL)3
xxx
INITIAL
INCUBATION
TEMP. I'd
XX.X
INCUBATION TIME
(24-HR TIME)
START
(KH:MM)
DURATION OF
INCUBATION
(HH:MM)
lg.'3Q ZZ
oo
SAMPLE ID
(BARCODE)
FINAL O,
(MG/LT
XXJ<
243663
7,5
INITIAL P.O. AT 1000'ELEVATION
7.2
FINAL 0.0. AT 2200' - METER CALIBRATED
243665
r.i
FOR EACH ELEVATION
2^3667
7.0
CO//TROL
OXYGEN METER CALIBRATION INFORMATION
MEMBRANE CHECK
ELECTRONIC ZERO
CALIBRATION CHAMBER TEMPERATURE:
SATURATED O2 @ @ TEMP.:
STATION ELEVATION (FROMTOPO. MAP OR ALTIMETER):
ELEVATION CORRECTION FACTOR:
Th» caSbtatlcxi value Is obtained by multiplying the saturated DO concentration times an
dotation axrect'on laaor (obtained from the tables on the back ol the YSI meter). Adjust the
meter reading to the calibration value.
CALIBRATION VALUE:
Flag codes: K - no measurement or observation made; U = suspect measurement or observation; Q = unacceptable QC check associated with measurement; F1, F2, etc. =
miscellaneous (lags assigned by each field crew. Explain all flags in comments section.
Rev. 05/29/98 (strvlldm.98)
FIELD MEASUREMENT FORM - STREAMS/RIVERS - 1
Figure 8-1. Field Measurement Form (page 1), showing data for sediment metabolism samples.
-------
SEDIMENT METABOLISM
SITE ID:
DATE: j I S 198
SAMPLE TYPE: (m) R2 R3 R4 R5
229000
SEDIMENT METABOLISM
SITE ID: ORRV_9 5.-_2._9_5.
DATE: JJ_/_S_/98
SAMPLE TYPE: R1 (p3 R3 R4 R5
229001
SEDIMENT METABOLISM
SITE ID: ORRV _i JL-.13_ 3_
DATE: g / £~I98
SAMPLE TYPE: R1 R2
-------
Jl -
-------
Section 9
Benthic Macroinverfebrates
Donald J. Klemm1, James M. Lazorchak1, and David V. Peck2
Benthic macroinvertebrates inhabit the sedi-
ment or live on the bottom substrates of lakes,
streams, andrivers. Themacroinvertebrate assem-
blages in rivers reflect the overall biological integ-
rity of the benthic community such thatmonitoring
these assemblages is useful in assessing the status
of the water body and monitoring trends. Benthic
communities respond differently to a wide array of
stressors. As aresult of this, it is often possible to
determine the type of stress that has affected a
benthic macroinvertebrate community (Plafkin et
aL, 1989; Klemm etal., 1990; BarbouretaL, 1999
). Because many macroinvertebrates have relatively
long life cycles of ayear or more and are relatively
immobile, macroinvertebrate community structure
isafunction of past environmental conditions.
EMAP scientists are currently evaluating
two different approaches to developing ecologi-
cal indicators based on benthic invertebrate as-
'U.S. EPA, National Exposure Research Laboratory, Eco-
logical Exposure Research Division, Ecosystems Research
Branch, 26 W. Martin Luther King Dr., Cincinnati, OH
45268.
2U.S. EPA, National Health and Environmental Effects Re-
search Laboratory, Western Ecology Division, Regional Ecol-
ogy Branch, Corvallis, OR 97333.
semblages. The first is a multimetric approach,
where different structural and functional at-
tributes of the assemblage are characterized as
"metrics". Individual metrics that respond to dif-
ferent types of stressors are scored against ex-
pectations under conditions of minimal human
disturbance. The individual metric scores are
then summed into an overall index value that is
used to judge the overall level of impairment of
an individual stream reach. Examples of
multimetric indices based on benthic invertebrate
assemblages include Kerans and Karr (1994),
Fore etal. (1996), Barbour etal. (1995; 1996),
and Karr and Chu (1999).
The second approach being investigated
is to develop indicators of condition based on
multivariate analysis of benthic assemblages and
associated abiotic variables. Examples of this
type of approach as applied to benthic inverte-
brate assemblages include RIVPACS (Wright
1995), and BEAST (Reynoldson et aL, 1995).
Rosenberg and Resh (1993) present various
approaches to biological monitoring .using
benthic invertebrates, and Norris (1995) briefly
summarizes and discusses approaches to ana-
-------
lyzing benthic macroinvertebrate community
data.
Field procedures for collecting and pro-
cessing benthic invertebrate samples from non-
wadeable streams are presented in Section 9.1.
These procedures are based upon draft proce-
dures developed for the Mid-Atlantic Integrated
Assessment (MAIA) study conducted in the
eastern U.S. Section 9.2 contains an equipment
and supply checklist for benthic invertebrate
sampling.
9.1 Sampling Procedures
for Non-wadeable Streams
The length of river reach established for
larger non-wadeable streams and rivers is much
larger than for wadeable streams, making a vi-
sual estimate of the number of riffle and pool
macrohabitat units impossible. In addition, mid
channel depths of larger streams and rivers will
make it impractical to collect kick net samples
from mid-river habitats. In non-wadeable
streams and rivers, samples are collected at each
of eleven transects established for physical habi-
tat characterization. At each transect, two kick
net samples are obtained from shallow area (<
1m) near the bank of the river. Kick net samples
collected from each transect are composited into
a single sample for the river; samples collected
from different macrohabitat types are not
composited separately. Akick net modified for
use by one person is shown in Figure 9-1. In
addition to the mesh size used in the two EMAP
studies other mesh sizes such as 250 - 800 m
can be used depending upon the objectives of
1.5 m long, 2-piece detachable handle
Mesh = 595/600
Sewed End
30cm
Canvas Bottom Panel
Figure9-1. Modified kick net. (Not drawn to scale.).
-------
the program and potential for clogging. For deep
rivers that are extremely difficult or hazardous
to obtain benthic samples with a kick net, a
ponar or core grab sample could be used in-
stead.
In addition, two daytime drift net samples
are collected from as near to the downstream
end of the defined reach as is practical. The drift
net assemblies are positioned when the crew
drops off a vehicle at the take-out point, and
are retrieved when the crew reaches the sam-
pling point in the boat.
9.1.1 Sample
Collection Using Kick
Nets
9.1.1.1 Selection of
Sampling Points
Samples are collected from non-wadeable
streams during a downstream traverse of the
sample reach. At each transect location, locate
a suitable sampling point on the same side of
the river as fish sampling is conducted (Section
10). Locate the sampling point in an area away
from the river margin, but at a depth less than or
equal to 1 m.
9.1.1.2 Sample Collection
At each sampling point, obtain TWO kick
net samples using the procedures presented in
Table 9-1 (if the sampling point is located in a
riffle or glide macrohabitat) or Table 9-2 (if the
sampling point is located in a pool macrohabitat).
If there is insufficient flow to sample a transect
with the modified kick net following this proto-
col, spend about 60 seconds hand picking a
sample from approximately 0.25 m2 of substrate
at the station and combine it with samples from
other transects in the bucket. If there is too little
water to collect the sample with the kick net,
randomly pick up 10 rocks, and pick and wash
the organisms off of them into the bucket. Keep
a note of this on the field sheets and in all data-
bases generated from sites where more than one
transect has to be sampled in this manner. Re-
sults may show a bias due to the larger organ-
isms picked in this approach.
9.1.2 Sample
Processing: Kick Net
Samples
After all transects have been sampled, the
composite sample is processed as described in
Table 9-3. Sample labels to put on and in the
jar are shown in Figures 9-2 and 9.5, respec-
tively, and the sample collection form is shown
in Figure 9-4. Ensure the sample is preserved
and that the jar is completely sealed. Place the
sealed sample jar upright in a cooler or plastic
bucket for transport. Blank labels for use inside
of sample jars are presented in Figure 9-5. These
can be copied onto waterproof paper.
9.1.3 Description of
Drift Nets and Habitat
Sampled
Drift nets are stationary nets designed to
sample organisms from flowing waters such as
streams and rivers. The drift net sampler is de-
signed to obtain qualitative and quantitative
samples of macroinvertebrates which either ac-
tively or passively enter the water column from
all types of substrates in flowing water with a
velocity of not less than 0.05 m/s. They can be
used to capture organisms at and below the sur-
face of the water. Drift nets can be used indi-
vidually or in groups with nets strung out side
by side or arranged vertically.
-------
Table 9-1. Collecting Kick Net Samples From Non-wadeable Streams: Riffle/Run Macrohabitats.
1. Attach the four foot handle to the kick net. Care should be exercised to be sure the handle is on tight or
the net might become twisted hi strong current or while dragging it through the water causing the loss
of part of the sample.
2. Position the sampler quickly and securely on the river bottom with the net opening upstream so as to
eliminate gaps under the frame. Reposition the sampling point to avoid large rocks that prevent the
sampler from seating properly.
3. The sampling area (or quadrate) has a width and length equal to the width of the net frame (0.5 m) or a
total area-0.25m2.
4. Hold the sampler in position on the substrate and check the quadrat directly in front of the net for heavy
organisms, such as mussels and snails. Place these organisms into the net.
5. Continue to hold the sampler securely while vigorously kicking the substrate within the quadrat for 20
seconds (use stopwatch).
6. After 20 seconds, hold the net in place with the knees. Pick up any loose rocks in the quadrat and scrub
off organisms in front of the net. Place any additional mussels and snails found in the quadrat in the net.
7. Remove the net from the water with a quick upstream motion to wash the organisms to the bottom of the
net. Immerse the net several times to remove fine sediments and to concentrate organisms at the end of
the net. Avoid having any water or additional material enter the mouth of the net during this operation.
8. Transfer the contents of the net into a plastic bucket half filled with water by inverting the net into the
bucket.
9. Inspect the net for clinging organisms. Use forceps and remove any organisms found and place them
into the bucket.
10. Carefully inspect large objects (rocks, sticks, leaves, etc.) in the bucket. Wash off any organisms, then
discard the objects. Remove as much detritus, sediment, and debris as possible without losing any
organisms.
11. See Table 9.3 for processing kick net samples.
The drift net consists of a bag of nylon or
nylon monofilament frame. The standard drift
net is approximately 1 m (39.3") long and has a
closed end. The drift net open end is 30.48 cm
(12") x 45.72 cm (18"). The net frame is made
of stainless steel rods or P VC pipe. The frame
of the drift net is anchored into the river bed by
a pair of steel rods, 15.46 cm (18") long or can
be attached to a "floating drift assembly" device
(Figure 9-3), Wildlife Supply Co., 1999-2000.
Drift net frames can also be fitted anteriorly with
a mouth reducing rectangular plexiglass enclo-
sure (Rutter and Ettinger, 1977;Wefring, 1976)
to increase filtration efficiency. For EMAP,
MAIA sampling in Regions 2,3, and 4 rivers, a
drift net with 600 m mesh openings has been
used in conjunction with the floating drift assem-
bly device (other mesh sizes such as 250 - 800
m can be used depending upon the objectives
of the program and potential for clogging).
The drift collection usually represents a
wide spectrum of the habitats found in a river.
Drift nets are effective for the collection of emi-
grating and dislodged benthic macroin-verte-
brates drifting in the water column of flowing
streams and rivers. Sampling efficiency of this
gear is a function of current velocity and sam-
pling period. Data collected can be used to es-
timate macroinvertebrate drift densities and rates
(individuals per unit volume of water per unit
time passing through the net). However, this re-
quires an estimate of the volume of water pass-
ing through the sampling nets. This is accom-
plished by averaging repeated measures of the
water velocity at the mouth of the drift net and
recording the total time the drift net is set in the
-------
I Table9-2. CollectingKickNetSamplesFromNon-wadeableStream:PooI\GlideMacrohabitats.
Attach the four foot handle to the kick net. Care should be exercised to be sure the handle is on tight or
the net might become twisted in strong current or while dragging it through the water causing the loss
~;jaf part of the sample.
_ The sampling area (or quadrate) has a width and length equal to the width of the frame (0.5 m) or a total
" area=0.25m 2.
Inspect the river bottom within the quadrat for any heavy organisms, such as mussels and snails.
Remove and place these organisms into the net.
Disturb the substrate within the quadrat by kicking vigorously with the feet. Drag the net repeatedly
and continuously through the disturbed area just above the bottom whole continuing to kick for 20
seconds (use a stopwatch). Keep moving the net so that the organisms trapped in the net will not
escape.
_ Remove the net from the water with a quick upstream motion to wash the organisms to the bottom of the
~ net. Immerse the net several times to remove fine sediments and to concentrate organisms at the end of
- the net. Avoid having any water or additional material enter the mouth of the net during this operation.
Hold the net so that the mouth is out of the water and the net is partially submerged. Pick up any loose
"rocks hi the quadrat and rub or brush any organisms found on them into the net. Also recheck the
quadrat for any additional snails or clams and place them in the net.
Transfer the contents of the net into a bucket half filled with water by inverting the net into the bucket.
Inspect the net for clinging organisms. Use forceps and remove any organisms found and place them in
-- the bucket
Carefully inspect large objects (rocks, sticks, leaves, etc.) in the bucket. Wash any organisms found
— into the bucket, then discard the objects. Remove as much detritus, sediment, and debris as possible
_._ without losing any organisms.
. See Table 9.3 for processing kick net samples.
water column. Repeated measures of the water
velocity are most representative of the sample
period if they are taken when the nets are first
set and just prior to removal of the net from the
system.
Limitations and hazards of daytime driftnet
sampling include:
• Unknown where organisms come
from; terrestrial species may make up
part of sample in summer and periods
of wind and rain. It is not the ideal time
of day for sampling drifting benthic
macroinvertebrates.
. • Installing and retrieving drifts from
areas of swift current can be hazardous.
Placing nets in highly visible areas may
result in tampering or theft. Boats and
skiers may damage drift net assemblies.
9.1.3.1 Drift Net Sampling
Procedures
ForEMAP, MAIApilot studies of Regions
2,3, and 4 rivers, install two drift nets at transect
K (See Table 9-4), one about 25 cm from the
bottom substrate and one about 10 cm below
the surface in water not exceeding 3 m in depth,
using cable and anchor attached to a "floating
drift assembly" device. The installation proce-
dure for drift nets is presented in Table 9-4. In-
stall the net in an area of river that is receiving
part of the main channel flow, but that can be
safely accessed by wading. A location that you
would consider to provide a representative
water chemistry sample is probably also suit-
able for positioning a drift net. Do not use drift
nets if the current velocity at the sampling point
is less than 0.05 m/s or more than same rate.
-------
Table9-3. ProcessingKickNetSamples: Non-wadeableStreams.
1. Fill out a sample label for the composite samples. Attach the label to a 500-mL (or 1 -L) plastic jar. If the
sample contains alarge volume of material, complete a sample label for additional containers and attach
them. Make sure the barcode numbers on each label are identical.
2. Hand-pick large organisms from the bucket containing the composite sample and place them into the
appropriately labeled jar.
3. Hand-pick large rocks and sticks remaining in the bucket. Use 'a small brush to scrub debris from them
back into the bucket. Discard the rock or stick.
4. Empty the contents of the bucket into a sieve (600 m) mesh, and then transfer into the labeled jar. NOTE:
Do not fill the jar more than ° full of material. If necessary, use a larger jar and/or distribute the sample
among two or more labeled jars. Rinse residue from the bucket into the jar using a wash bottle and a small
volume of water.
5. Add 95% ethanol to each labeled jar so that the final concentration of ethanol is at least 70%. If there is
a small amount of water in the sample, it may not be necessary to fill the jar entirely full to reach a 70%
concentration. It is very important that sufficient ethanol be used to reach a 70% concentration.
Otherwise, the organisms will not be properly preserved.
6. Place the waterproof label with the following information inside each jar:
• Stream Number • Date of collection
• Type of sampler and mesh size used • Collectors initials
• Habitat type (riffle/run, pool/glide) • Number of transects composited
• Name of stream
7. Rinse the bucket well to eliminate any residue.
8. Complete the Sample Collection Form and on the jars (1 of 2,2 of 2, etc). Record the barcode number of
the composite sample, and the habitat type (shore). If more than one container was required for a
sample, record the number of containers on the collection form. Replace the lid on the jar. Seal the
container lid(s) with plastic or electrician's tape. Also note any peculiarities associated with a particular
samples by using a flag code and/or a written comment on the collection form.
COMPOSITE BENTHOS
SITE ID: ORRV_?; Z_-4_3L
DATE: JL/_j5L/98_
HABITAT: Shore (Drift"
229001
COMPOSITE BENTHOS
SITE ID: ORRV .^ _S-_3.£?_i.
DATE:
HABITAT:
Drift
229000
COMPOSITE BENTHOS
SITE ID: ORRVJ? JL--3.33.
DATE: g/S'/QS
HABITAT: (Shore) Drift
BARCODE: 22100Q
Figure 9-2. Completed labels for benthic macroinvertebrate samples. The bottom label is used if
more than one jar is required for a composite sample.
-------
Net
Shallow Set
Pentagonal Floating PVC Frame
- Chain
Flow
U-Shaped
PVC Pipe
Net
Deep Set
Figure 9-3. Shallow and deep set drift net assemblies. (Not drawn to scale).
Measure the current velocity at the entrance to
each net at the time the net is installed and again
when it is retrieved. Velocity is determined by
timing a neutrally buoyant object over a known
distance or using a flow meter.
9.1.3.2 Processing and
Preservation of Drift Samples
After retrieving the drift nets from the
stream or river, process the sample as described
in Table 9-5. Sample labels are shown in Figure
9-2, and the sample collection form is shown in
Figure 9-4. Note that the material from the two
nets is combined to yield a single composite
sample of drift for the stream or river. Blank
labels for use inside of sample jars are presented
in Figure 9-5. These can be copied onto water-
proof paper.
9.1.3.3 Maintenance of the
Drift Nets
After the drift sample has been processed
and preserved, thoroughly wash the drift nets
with water from the stream or river to remove
all debris, etc.
9.2 Equipment Checklist
A list of all equipment and supplies required
to conduct benthic invertebrate sampling is pre-
sented in Figure 9-6.
-------
Reviewed by (initial)
SAMPLE COLLECTION FORM - RIVERS
SITE NAME: BEAVE"R
DATE:
/ 5 1 98 VISIT DO gl 1 D2 D3
SITE ID: ORRV ^ £ - *? *? 9
TEAM ID (X):
D2 D3 D4 D5 D6 D7 D8
COMPOSITE BENTHOS SAMPLES
BANK SAMPLED (LOOKING DOWNSTREAM): §3 LEFT Q ™GHT
SAMPLED
(BARCODE)
HABITAT
(CIRCLE ONE)
NO.
OF JARS
FLAG
S = SHORE
M = MID-CHANNEL
D « DRIFT
COMMENTS
2436 IZ
M
£D AT LAUhlCft
24-36
NO SHORE KICK AT TffAMSECTS
=>7DO DEEP
STATION (CIRCLE IF DONE)
SHORE KICK
MID-CHANNEL
s
M
s
M
S
M
DRIFT
NET
TRANSECT
TIME (24-HR CLOCK)
START
• VELOCITY
• (FT/SEC)
XX.XX
DB-DC DP DE DF DG DH Pi DJ DK pCf ;
)
-------
BENTHOS IDENTEEICATION
Site Number
Stream
Collection Date.
Sampler
Habitat Type
Collector(s) _
Number of Transects
BENTHOS IDENTIFICATION
Site Number
Stream
Collection Date.
Sampler
Habitat Type
Collector(s) _
Number of Transects
BENTHOS IDENTIFICATION
Site Number
Stream
Collection Date.
Sampler
Habitat Type
Collector(s) _
Number of Transects
BENTHOSIDENTIFICATION
Site Number
Stream
Collection Date
Sampler
Habitat Type
Collector(s) '
Number of Transects
Figure 9-5. Blank labels for benthic invertebrate samples.
-------
Table9-4. CollectionProceduresforDriftNetSamples: Non-wadeableStreams.
NOTE: Do not use drift nets for large rivers with currents less than 0.05 m/s.
Installation and Retrieval of Drift Nets:
1. Ideally, the net should be installed at the downriver end of the designated sampling reach (transect K in
non-wadeable streams). In practice, the net is installed at either the takeout point (1 st choice) or put-in
point, whichever is located closer to the designated sampling reach. Mark the nearest transect on the
Sample Collection Form and note if the drift net location is outside of the designated sampling reach in
the Comments section of the collection form.
2 Locate the drift net assembly in an area receiving the main flow of the river (i.e., avoid backwaters, river
margins, eddies, etc.)
3. Anchor the net assembly using anchors and cables.
4. RecordtheSTARTTIMEofsamplingontheSampleCollectionForm.
5. Measure the current velocity at the entrance of the net, using the neutrally buoyant object technique (or
a flow meter) as follows:
A. Measure out a straight segment of the river reach just upstream of the drift net location in which
an object can float relatively freely and passes through within about 10 to 30 seconds.
B. Select an object that is neutrally buoyant, like a small rubber ball or an orange; it must float, but
very low in the water. The object should be small enough that it does not "run aground" or drag
bottom.
C Time the passage of the object through the defined river segment 3 times. Record the length of
the segment and each transit time in the Comments section of the Sample Collection Form.
6. The net assembly should be left in the river for at least 3 hours or as long as possible at the site. Upon
return to the net location after floating the designated sampling reach, retrieve the net assembly from the
water, taking care not to disturb the bottom upstream of the net.
7. Record the END TIME on the Sample Collection Form.
8. Determine the current velocity again as described in Step 5 above. Record the three "final" velocity
estimates in the Comments section of the collection form. Calculate the average velocity from the initial
and final values (6 measurements). Record the average velocity in the "Velocity" field of the Sample
Collection form. Exclude any gross outlier values from the computation of the average velocity.
9. Note in the comments section if the net is badly clogged, which may occur at locations with high
discharge and/or where the float time of the sampling reach is long.
-------
|i Table 9-5. Procedures for Processing Drift Net Samples: Npn-wadeable Streams.
(;: 1. Fill out a sample label for the composite drift sample. Attach the label to a 500-mL or 1 -L plastic jar. If the ;
i- sample contains a large volume of material, complete a sample label for additional containers and attach \
t _ it to a second jar. Make sure the barcode numbers on each label agree. ]
}|:2. Concentrate the material in each net in one corner by swishing up and down in the stream or river. Wash ;
IL the material into a bucket half-filled with water. Use a wash bottle and/or forceps to remove as much
|E material as possible from the net. • . - i
|- 3. Repeat Step 1 for the second drift net. The contents of both nets are combined into a single bucket. i
p,4. After the two net samples are combined into a single bucket, pour the composite sample into a sieving ;
1%1 bucket (595 micron mesh).
US Hand-pick large organisms from the sieve bucket containing the composite sample and place them into '
C . the appropriately labeled jar. !
p,6 Hand-pick large rocks and sticks remaining in the bucket. Use a small brush to scrub debris from them '.
]g; back into the bucket. Discard the rock or stick. , j
p7. Lightly "tapping" the bottom of the sieve bucket on the surface of the stream or river helps to remove ;
|~ fine material. Remove as much material as possible using the sieve bucket.
I- 8. Empty the contents of the bucket into the labeled jar. If necessary, distribute the sample among two or
g^r: more labeled jars. Rinse residue from the bucket into the jar using a wash bottle and a small volume of
f--~~ water. ' . '- •'"'".'• -'•'/• " .-.....•. .. ., • :. .:. .. . ...... _, • ,
jr::- — - . - . ' - •..-.• (.,-. . . • •••-.'---' '•'...- . .. ' . : • .-•.--..- 1
p9. Add 95% ethanol to each labeled jar so that the final concentration of ethanol is at least 70%. If there is ^
jEc:- a small amount of water in the sample, it may not be necessary to fill the jar entirely full to reach a 70%
|^ concentration. It is very important that sufficient ethanol be used to reach a 70% concentration.
Otherwise, the organisms will not be properly preserved.
Place a waterproof label with the following information inside each jar:
lO.
Date of collection
Collectors initials
Number of transects composited
Stream Number
• Type of sampler and mesh size used
• Habitat type (drift net)
• Name of stream
Rinse the bucket well to eliminate any residue.
Complete the Sample Collection Form. Record the barcode number of the composite sample, and the
habitat type (drift). If more than one container was required for a sample, record the number of
containers on the collection form and on the jars. Replace the lid on the jar, and seal the container lid(s)
with plastic or electrician's tape. Also note any peculiari ties associated with a particular samples by
using a flag code and/or a written comment on the collection form.
' •" "
-------
Qty.
Equipment and Supplies for Benthic Macroinvertebrates
Item
1
1
1
2
1
2pr.
1
1
1
12
2
Igal
2pr.
1
6
6
2
1
Ipkg.
4rolls
1
1
1
Ipkg.
1 copy
1 set
Modified kick net with 595 m mesh openings and closed bag (Wildco #425-150-595)
Handle for Kick Net Sampler, four foot length
Floating drift net assembly (PVC frame, chains, snap-clips, and carabineers)
Drift nets, 595 m mesh, closed end
Sieve-bottomed bucket, 595- m mesh openings (optional)
Watchmakers' forceps
Wash bottle, 1-L capacity
Small spatula, spoon, or scoop to transfer sample
Funnel, with large bore spout
Sample jars, plastic with screw caps, 500 mL and 1 L capacity, suitable for use with
ethanol
Buckets, plastic, eight to ten quart capacity
95% ethanol, in a proper container
Rubber gloves, heavy rubber
Coolef (with suitable absorbent material) for transporting ethanol and samples
Sample labels, pre-numberedbarcoded, stick-on type
Sample labels, blank, stick-on type (for additional containers)
Sample Collection Form for site
Field check list sheet
Soft (#2) lead pencils
Clear waterproof tape strips
Plastic electrical tape
Knife, pocket, with at least two blades
Stopwatch
Pocket-sized field notebook (optional)
Kim wipes in small self-sealing plastic bag
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for benthic
macroinvertebrates
Figure 9-6. Equipment and supply checklist for benthic macroinvertebrates.
9.3 Literature Cited
Allan J.D. and E. Russek. 1985. The
quantification of stream drift. Can J. Fish.
Aquat. Sci. 42(2)210-215.
ASTM.I996a. Standard practice for collecting
benthic macroinvertebrates with driftnets.
ASTM Designation: D 455-85. Annual
Books of ASTM Standards. Pesticides;
Resource Recovery; Hazardous Sub-
stances and Oil Spill Responses; Waste
Disposal; Biological Effects. Vol.1104,
ASTM, 100 Barr Harbor Drive, West
Conshohocken, PA. pp.89-91.
ASTM. 1996b. Standard guide for selecting
stream-net sampling devices for collecting
-------
benthic macroinverte-brates. ASTM Des-
ignation: D 4556-85. Annual Books of
ASTM Standards. Pesticides; Resource
Recovery; Hazardous Substances and Oil
Spill Responses; Waste Disposal; Biologi-
cal Effects. Vol.1104, ASTM, 100 Barr
Harbor Drive, West Conshohocken, PA.
pp.78-88.
Barbour, M.T., J.B. Stribling, and J.R. Karr.
1995. The multimetric approach for
establishing biocriteria and measuring
biological condition. Pages 69-80 In: W.S.
Davis and T.P. Simon (eds.) Biological
assessment and criteria: tools for water
resource planning and decision-making.
Lewis Publishers, Chelsea, MI
Barbour, M.T., J. Gerritsen, G.E. Griffith, R.
Frydenborg, E. McCarron, J.S. White, and
M.L. Bastian. 1996. A framework for
biological criteria for Florida streams using
benthic macroinvertebrates. Journal of the
North American benthological Society
15(2): 185-211.
Barbour, M.T., J. Gerritsen, B.D. Snyder, and
J.B. Stribling. 1999. Rapid bioassessment
protocols for use in wadeable streams and
rivers: periphyton, benthic macroinverte-
brate, and fish. EPA/841/B-99/002. U.S.
Environmental Protection Agency, Office of
Water, Washington, D.C.
Fore, L.S., J.R. Karr, and R.W. Wisseman.
1996. Assessingn invertebrate responses to
human activities, evaluating alternative
approaches. Journal of the North American
Benthological Society 15(2):212-231.
Karr, J.R. andE.W. Chu. 1999. Restoring life in
running waters. Island Press, Washington,
D.C., 206 pp.
Kerans, B.L. and J.R. Karr. 1994. A benthic
index of biotic integrity (B-IBI) for rivers of
the Tennessee Valley. Ecological Applica-
tions 4:768-785.
Klemm, D.J., P.A. Lewis, F. Fulk, 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 Pro-
tection Agency, Cincinnati, OH 45268.
Norris, R.H. 1995. Biological Monitoring: the
dilemma of data analysis. Journal of the
North American Benthological Society
14:440-450
Oregon Department of Environmental Quality.
1997. Biological Assessment of Wadeable
Streams of the Upper Deschutes River
Basin: Quality AssuranceProjectPlan
Plafkin, J.L., M.T. Barbour, K.D. Porter, S.K.
Gross, and R.M. Hughes. 1989. Rapid
Bioassessment Protocols for Use in
Streams and Rivers: Benthic Macro-
invertebrates, and Fish. EPA/440/4-89/
001. U.S. Environmental Protection Agency,
Washington, D.C.
Reynoldson, T.B., R.C. Bailey, K.E. Day, and
R.H. Norris. 1995. Biological guidelines for
freshwater sediment based on Benthic
Assessment SedimenT (the BEAST) using
a multivariate approach for predicting
biological state. Australian Journal of
Ecology 20:198-219.
Rosenberg, D.M. and V.H. Resh. 1993.
Freshwater biomonitoring and benthic
macroinvertebrates. Chapman and Hall,
New York.
Rutter, R.P.M. and Ettinger, W.S. 1977.
Method for sampling invertebrate drift from
a small boat. Progressive Fish-Culturist
39(l):49-52.
Washington Department of Ecology. 1997.
Biological Assessment ofWadeable Streams
-------
of the Chehalis River Basin: Quality
Assurance Project Plan.
Wefring, D.R. 1976. A method for the
collection of invertebrate drift from large
rivers. M.A. Thesis, St. Cloud State
University, St. Cloud, MN.
Wildlife Supply Company. 1999. Drift Net
Floating Assemblies. 301 CassSt,Saginaw,
MI, Page 7. (E-Mail: goto@wildco.com;
WebSite: http://www.wildco.com), (phone:
800-799-8301, FAX: 800-799-8115).
Wright, J.F. 1995. Development and use of a
system for predicting the macroinvertebrate
fauna in flowing waters. Australian Journal of
Ecology 20:181-197.
9-11:
-------
Section 10
Aquatic Vertebrates
Frank H. McCormick1 and Robert M. Hughes2
Vertebrate sampling is intended to collect
all but the rarest fish and aquatic amphibian spe-
cies in a reach and their abundances in the col-
lection should be relative to their proportionate
abundance in the water body. Data on species
richness, species guilds, abundance, size and
anomalies are used to assess ecosystem condi-
tion. In rivers, vertebrates are collected first.
Boat electrofishing equipment is used as the prin-
cipal sampling gear (Section 10.1.1), and only
the boat personnel are involved in collecting
aquatic vertebrates. In addition to gathering data
on the assemblage, fish specimens are retained
for analysis of tissue contaminants (Section 11).
10.1 Sample Collection
Depending on the survey region, rivers are
sampled along one bank for a distance equal to
either 40 or 100 times the wetted width in the
'U.S. EPA, National Exposure Research Laboratory,
Ecological Exposure Research Division, 26 W. Martin
Luther King Dr., Cincinnati, OH 45268.
2Dynamac Internationa! Corp., 200 SW 35th St.,
Corvallis, OR 97333.
vicinity of the point of entry. The mean channel
width is measured with a laser range finder and
estimated from maps and at the top of the reach.
In the relatively fast, cold, oligotrpphic, or spe-
cies-depauperate rivers of some regions, lower
total fish catches and efficiency of capture (com-
pared with those in relatively slow, warm,
eutrophic, or species-rich rivers) necessitate a
greater sampling reach length. To capture a suf-
ficient number offish in rivers of some regions,
sample reaches 100 Channel-Widths long may
be specified for regional surveys, based largely
upon fish capture requirements. River reaches
40 channel widths long are specified in the Mid-
Atlantic region, for example, whereas 100 chan-
nel-width long reaches are specified in Pacific
Northwest rivers.
10.1.1 Electrofishing
Because vertebrates are collected using
electrofishing units, safety procedures must be
followed at all times (refer to Section 2). Pri-
mary responsibility for safety while electrofishing
rests with the crew leader. Electrofishing units
have a high voltage output and may deliver a
fatal electrical shock. While electrofishing, avoid
-------
contact with the water unless sufficiently insu-
lated against electrical shock and, do not touch
objects outside the boat. Use watertight rubber
linesman's gloves. If gloves develop a leak or
become wet inside, use another pair or stop fish-
ing until they are repaired and thoroughly dry.
Avoid contact with the anode and cathode at all
times due to the potential shock hazard. At no
time while the electrofisher is on should a team
member reach into the water for any reason. If
it is necessary for a team member to reach into
the water to pick up a fish or something that has
been dropped, do so only after the electrical
current has been turned off. Do not electrofish
when navigating major rapids and wait for the
second boat to clear them. Do not resume
electrofishing until allnontarget individuals are
clear of the electroshock hazard or obstacle.
The boat units have three kill switches. Insure
that these workbefore fishing. Do not make any
modifications to the electrofishing unit that in-
terrupt the current or that would make it impos-
sible to turn off the electricity.
Crew members must complete CPR and
first aid courses. They should be strong swim-
mers and, as appropriate, complete a white
water rescue course. Wear a life jacket when in
a boat and avoid operating electrofishing equip-
ment within 20 feet of nontarget organisms. Dis-
continue activity during thunderstorms, heavy
rain or if the top or inside of the boat is wet.
Crew members should keep each other in con-
stantview or communication while electrofishing.
For each site, know the location of the nearest
hospital with a defibrillation unit. Although the
crew leader has authority, each crew member
has the responsibility to question and modify an
operation or decline participation if it is unsafe.
Use hand signals to communicate direction and
power on or off because of generator noise, and
avoid colliding with obstacles overhead and in
the water. Rest if the team becomes fatigued,
and drink lots of water.
Gasoline is extremely volatile and flam-
mable. Its vapors readily ignite on contact with
heat, spark or flame. Never attempt to refill the
generator while it is running. Always allow the
generator to cool before refilling. Keep
gasoline out of direct sunlight to reduce volatil-
ization and vapor release. Always wear gloves
and safety glasses when handling gasoline. Keep
gasoline only in approved plastic containers and
store in a tightly closed container.
Boat electrofishing sampling procedures
are presented in Table 10-1. Record informa-
tion on the Vertebrate Collection Form as shown
in Figure 10-1. If the river cannot be sampled
by electrofishing, complete the "NOT FISHED"
field on the form. Select the initial voltage based
on the measured conductivity of the river (see
Section 5). Select the initial frequency based on
the expected size offish. If fishing success is
poor, increase the pulse width first and then the
voltage. Increase the frequency last to minimize
mortality or injury to large fish.
The electrofishing boat is a 14 -16 ft. in-
flatable raft or John boat modified for two per-
sons and all fishing equipment Boat configura-
tion consists of a frame mounted generator and
electrofishing control box, port and starboard
cathodes, and two anodes extending out over
the bow. Alternatively, the John boat itself may
be used as the cathode. The boat is maneuvered
by one operator seated near the stern, and the
vertebrates are collected and identified by one
netter operating from the bow. Prior to fishing,
determine that the netter is wearing gloves and
both team members are clear of all electrodes.
Wear polarized sunglasses to aid vision. Start
the electrofisher, set the timer to zero, and de-
press the foot pedal to begin fishing. Starting at
the top of the reach and along the designated
-------
I Table 10-1. Procedure to Collect Aquatic Vertebrates by Boat Electrofishing.
|p Onshore at launch site
t a. Check generator oil and fill tank with gas (wipe up any spillage).
f b. Clip cathodes to sides of frame & connect their cables to the cathode outlet (if the fishing site is
ig distant, keep electrodes in boat).
c. Connect anode cables to outlets (if the fishing site is distant, keep anodes on poles in the boat).
d. Connect generator and pulsator.
e. Confirm that all gear for the day and a spare vehicle key are in the boat.
f. Put on a life jacket and gloves.
g. Go to step 2 & 3 below to assess electrofisher performance.
fn.
"
*_..
s"
r.2.
£[:_-
ir
jfi
pr-
Complete the header information on a copy of the Vertebrate Collection Form. Indicate the transect
being sampled in the "TRANSECT" field on the form.
Select river bank for fishing (left for odd numbered sites, right for even) unless immediate hazards or
obstructions preclude this. Stay along the selected bank throughout the day's fishing to the degree it is
safely navigable; do not switch back and forth between banks unless the river aspect is unchanging
and the selected side is not representative (e.g., very sunny and shallow) of both. Using the
rangefinder, determine a downstream point that is 10 mean channel widths distant (this is the profile
length). Record this distance on the field sheet.
Check all electrical connections and potential conductors and place the anodes and cathodes in the
water. Fill livewell and put on linesman gloves. Verify that all electrical switches are off, that all non-
target organisms are clear of the water or two boat lengths away, and that boat surfaces are dry.
Start generator, switch to pulsed DC, a frequency of 30 pps, low range and 40%. Increase % (voltage) as
needed to roll fish. If success is poor, reduce %, switch to high range, and again increase % as needed.
If effectiveness is still low, switch to 60 pps and repeat the process. If the current (amperage needle) is
reduced, switch back to low range to avoid overloading the generator. Switching should occur when
power is off. Netter activates safety switches and insures that when either is employed current ceases.
Verify that fish are rolled and relaxed but not rigid. Record settings on field sheet in comments section
and start cleared clocks.
With system activated and safety switches on, begin fishing downstream near shore. Maneuver the
boat or anode to cover a swath two-three meters wide, at an oars length from shore, near cover, and at
depths less than three meters wherever possible. Do not place the boat in danger in order to fish
particular habitats; cut the generator and stow the gear before negotiating hazards.
Place fish directly in livewell as soon as possible; do not hold them in the electrical field. Pay special
attention to netting small and benthic fishes as well as fishes that respond differently to the current-not
just the big fish that move to the surface. Try to net all fish seen, but in productive systems this will be
impossible. Do not chase individual fish with the boat or lean far out from the boat to net them. If benthic
fish are being missed, pivot the boat occasionally or hold the net behind the anode and along the
bottom so some are collected.
Cease sampling at the end of the profile. Process the fish quickly and carefully, returning them to the
water unless they are vouchered. Be sure that the data sheet is completed accurately and completely,
and that voucher specimens are taken. Record the "Total Shock Time," "TOTAL FISHING TIME," and
"SHOCK DISTANCE" on the Vertebrate Collection Form. If no aquatic vertebrates were collected,
complete the "NONE COLLECTED" field on the Vertebrate Collection Form.
Complete field collections and field sheets for other indicators taken at the end of the profile. Return to
step 1 for each of the subsequent 9 profiles, but begin downstream from where fish were released.
shoreline, fish in a downriver direction. Adjust
voltage and output according to sampling ef-
fectiveness and incidental mortality to specimens.
The netter uses an insulated dip net to re-
trieve stunned individuals, which are then de-
posited into a livewell for later processing (Sec-
tion 10.2). Change the water in the livewell
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-------
periodically to minimize mortality prior to pro-
cessing. If individuals show signs of stress (loss
of righting response, gaping, gulping air, exces-
sive mucus), stop and process them. This should
only be necessary on very warm days, in long
reaches, or if very large numbers of individuals
are collected. Electrofishmg may also need to
cease at times to immediately process and re-
lease specimens (e.g., listed species or large
game fish or if fish appear to be stressed due to
temperature and/or low DO) as they are netted
(see Section 10.2). If periodic processing is re-
quired, be sure to release individuals upriver and
away from the shoreline to reduce the likelihood
of collecting them again.
At the completion of electrofishing each
profile, record the total operating time (shock
time) and total fishing time shown on the
electrofisher timer and the distance sampled by
electrofishing on the Vertebrate Collection Form
(Figure 10-2). If no aquatic vertebrates were
collected, indicate this on the form as shown in
Figure 10-2. During this project, specimens
should be processed after completion of every
transect when possible to provide data on catch
per unit effort.
10.2 Sample
Processing
Sample processing involves tallying and
identifying fish, examining individual specimens
for external anomalies, obtaining length measure-
ments from selected specimens, preparing
voucher specimens for taxonomic confirmation
and archival at a museum, and selecting speci-
mens to prepare samples for fish tissue contami-
nants (see Section 11). Process collections as
quickly as possible to minimize stress to live
specimens. The netter is responsible for identi-
fying, measuring, and examining aquatic verte-
brates contained in the livewell. At the end of
each profile, the netter processes fish from the
livewell while the operator records information
on the field data forms.
10.2.1 Taxonomic
Identification and Tally
Table 10-2 presents the procedure for
identifying and tallying aquatic vertebrates.
Record identification and tally data for each
species on the Vertebrate Collection Form as
shown in Figure 10-1. Also record comments
and data for additional species on the Verte-
brate Collection Form. The team is to be pro-
vided with a list of standardized names (re-
quired) and species codes (optional) for aquatic
vertebrate species that are expected to be col-
lected (see Appendix C for an example).
Taxonomic identification should be per-
formed only by trained ichthyologists familiar
with the fish species and other aquatic verte-
brate taxa of the region. Use taxonomic refer-
ence books and other materials that contain
species descriptions, ranges, and identification
keys to make species identifications in the field.
Where there are many individuals of easily iden1
tified species, processing may be facilitated by
keeping a tally count of the number of individu-
als of each species as it is taken from the livewell
and totaling the tally once processing is com-
plete.
To minimize handling, process threatened
and endangered species first, and immediately
return all individuals to the river. If conditions
permit and stress to individuals will be minimal,
photograph such fish for voucher purposes
(Section 10.2.3). Photographs offish, fish too
large to voucher, fish anomalies, and the sites
themselves are very informative to those of us
who cannot be in the field. Be sure to photo-
graph the site number so we can link photos
and places. Indicate if photographed with an "F"
series flag for the species on page 1 of the Ver-
-------
Figure 10-2. Fish length measurements (modified from Lagler, 1956).
I Table 10-2. Procedure To Identify, Tally, And Examine Aquatic Vertebrates.
1. Complete the header information on the form, then record the common name (from a standardized list)
i andspeciescodeonthefirstblanklineinthe"SPECIMENS"sectionoftheVertebrateCollectionForm.
If a species cannot be positively identified, assign it an' 'unknown'' species code from the list provided.
: 2. Examine each fish individually; small-sized fish species may be handled in small manageable groups to
speed processing.
1 3. To minimize handling, threatened and endangered species should be identified, counted, and returned
! immediately to the stream. If conditions permit and stress to individuals will be minimal, photograph fish
: for voucher purposes. Indicate if photographed on data sheet with flags and comments. If protected
fish have died, they should be vouchered in formalin. At the earliest possible time, the appropriate state
officials should be notified. ;
^ 4 Sport fish and very large specimens should be identified, measured for total length to the nearest mm,
examined for external anomalies, and released. Record all information on the vertebrate collection form.
« Keep voucher specimens (up to 5) of smaller individuals of each species. If no smaller individuals are
collected, photograph each species and indicate so on the data form. Large, questionable species
should be placed on ice and then frozen.
5. Identify all other species in the livewell.
6. Tally the number of individuals collected (use the "Tally" box on the Vertebrate Collection Form if
necessary) and record the total number in the "Count" field on the form.
7. Measure the total length of the largest and smallest individual to provide a size range for the species.
Record these values in the' 'Length'' area of the Vertebrate Collection Form.
8. Measure the total length of each individual (up to 30) and record the lengths in the boxes on the
Vertebrate Length Recording Form (2 lines of boxes per species). For smaller species, measure and
record lengths of a random set (up to 30) of the individuals collected.
9. Examine each individual for external anomalies and note the types of anomalies observed. After all of
the individuals of a species have been processed, record the anomaly code and the total number of
individuals affected in the "Anomalies" area of the Vertebrate Collection Form.
10. If individuals have died due to the effects of electrofishing or handling, record the total number of
mortalities on the Vertebrate Collection Form.
11. Follow the appropriate procedure to prepare voucher specimens and/or to select specimens for tissue
samples. Release all remaining individuals into the river.
12. Repeat Steps 1 through 11 for all other profiles.
-------
tebrate Collection Form (Figure 10-1) and
record a notation in the comments section. If
protected fish have died, they should be pre-
pared as voucher specimens and preserved in
formalin. Notify the appropriate state officials
as soon as possible.
If a species cannot be confidently identi-
fied in the field (e.g., small individuals or sus-
pected hybrids), record it as an "unknown" spe-
cies on the Vertebrate Collection Form, using
one of the names (and code) provided for un-
knowns from the standardized list (see Figure
10-1 for an example). If possible, flag unknown
species with an "F" series flag and provide your
best guess at an identification in the comments
section of the form.
10,2.2 External
Examination and
Length Measurements
During the tallying procedure for each spe-
cies (Table 10-2), examine each individual for
the presence of external anomalies. External
anomalies may resultfrom sublethalenvironmen-
tal or behavioral stress, diseases, and toxic
chemicals. Readily identified external anomalies,
include deformities, eroded fins, lesions, tumors,
diseases and parasites. Codes for different types
of anomalies are presented in Table 10-3.
Record the types of anomalies observed and
the number of individuals affected on the Verte-
brate Collection Form as shown in Figure 10-
1.
Blackening and exopthalmia may occa-
sionally resultfrom electrofishing. Injuries due
to sampling are not included in the tally of ex-
ternal anomalies, but should be noted in the com-
ments section of the Vertebrate Collection Form
(Figure 10-1). Care should be taken in the early
stages of electrofishing to use the most effective
combination of voltage and pulse width while
minimizing injury to fish. Blackening from
electrofishing usually follows the myomeres or
looks like a bruise. If fish die due to the effects
of sampling or processing, record the number
for each species on the Vertebrate Collection
Form (Figure 10-1).
For each species, use a measuring board
or ruler to determine the total length (Figure 10-
2) of the largest and smallest individuals (this is
a check on your measurements of total lengths
recorded on the Vertebrate Length Recording
Form [Figure 10-3]). Use of "tick marks" on
the length form will aid you in determining maxi-
mum and minimum lengths for a profile. Mea-
sure individuals on the right side, and slide fish
to touch the "Bump Board" on the measuring
board. Measure total length to the nearest milli-
meter (mm) and record these values on the Ver-
tebrate Collection Form as shown in Figure 10-
1. Measure the total lengths of up to 30
individuals and record these values on the Ver-
tebrate Length Recording Form as shown in
Figure 10-3.
10.2.3 Preparing
Voucher Specimens
With the exception of very large individu-
als and protected species, collect vouchers of
all species allowed by collecting permits to pro-
vide a permanent, archived, historical record of
fish collections. Prepare the voucher sample for
a site according to the procedure presented in
Table 10-4. Retain additional specimens of the
appropriate species for the fish tissue contami-
nants sample (Section 11). For each species,
voucher specimens take priority over specimens
for the tissue contaminants sample.
Voucher specimens for each species are
counted and placed into individual nylon mesh
bags (1 bag per species). Nylon stockings or
-------
Table 10^3. External Anomaly Categories and Codes.
Categories Code
Definition
Absent AB
Blisters BL
Blackening BK
Extensive black spot disease BS
Cysts CY
Copepod CO
Deformities DE
Eroded fins EF
Eroded gills B3
Fungus FU
Fin anomalies FA
Grubs WG
Hemorrhaging HM
Ich K
Lesions IE
Lice U
Mucus MU
None NO
Other OT
Scale anomalies SA
Shortened operculum • SO
Tumors TU
Leeches WL
Exophthalmia EX
Absent eye, fin, tail.
In mouth, just under skin.
Tail or whole body with darkened pigmentation.
Small black cysts (dots) all over the fins and body.
Fluid-filled swellings; may be either small or large dots.
A parasitic infection characterized by a worm-like copepod
embedded in the flesh of the fish; body extends out and
leaves a sore/discoloration at base, may be in mouth gills,
fins, or anywhere on body.
Skeletal anomalies of the head, spine, and body shape;
amphibians may have extra tails, limbs, toes.
Appear as reductions or substantial fraying of fin surface
area.
Gill filaments eroded from tip.
May appear as filamentous or "fuzzy" growth on the fins,
eyes, or body.
Abnormal thickenings or irregularities of rays
White or yellow worms embedded in muscle or fins.
Red spots on mouth, body, fins, fin bases, eyes, and gills.
White spots on the fins, skin or gills.
Open sores or exposed tissue; raised, granular, or warty
outgrowths.
Scale-like, mobile arthropods.
Thick and excessive on skin or gill, or as long cast from vent.
No anomalies present.
Anomalies or parasites not specified (Please comment).
Missing patches, abnormal thickenings, granular skin
Leaves a portion of the gill chamber uncovered
Areas of irregular cell growth which are firm and cannot be
easily broken open when pinched. (Masses caused by
parasites can usually be opened easily.)
Annelid worms which have anterior and posterior suckers.
They may attach anywhere on the body.
Bulging of the eye.
parity hose may substitute for nylon bags. Each
bag contains a numbered tag (Figure 10-4).
Record the tag number and the number of indi-
viduals vouchered for each species on the Ver-
tebrate Collection Form as shown in Figure 10-
1. Single specimens of easily identified and
distinct species (e.g., sandroller, smallmouth
bass) may be placed directly in the jar.
The preceding steps are critical to en-
able us to link a species' field and lab identi-
fications with the number of individuals so
named. If done correctly, we can estimate the
number of individuals collected from the pro-
portions in the bag, even if a presumed single
species turns out to be 2 or 3 species (this is
one reason we voucher as many specimens
of a species as possible). It is useful to pre-
_
-------
Page
EAMS/RIVERS
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'. Table 10-4. Guidelines and Procedures for Preparing Aquatic Vertebrate Voucher Specimens.
Determine the voucher category of a species and the number of specimens to include in the voucher
sample based on the following guidelines. NOTE: Category 3 species should be processed first.
A Category 1 — Large easily identified species OR adults may be difficult to identify OR the
species is uncommon in that region. Examples include:
Centrarchids
Salmonids
Catostomids
Cyprinids
Ictalurids
1. Preserve 1-2 small (<150 mm total length) adult individuals per site plus 2-5 juveniles. If only
large adults are collected, reserve smallest individuals until voucher procedure is complete
and preserve ONLY if space is available.
• NOTE: Individuals with a total length > 160 mm should be slit on the lower abdomen of
the RIGHT side before placing them into the container.
2. Photograph if considered too large for the jar and place in bag on ice for freezing (Do not
retain large gamefish). All photographs should include (1) a card with the stream ID, date,
species code, and common name, and (2) a ruler or some other object of known length to
provide some indication of the size of the specimen.
3. Retain additional individuals for the tissue contaminant sample.
B. Category 2—Small to moderate-sized fish OR difficult to identify species. Examples include:
Lampreys Minnows Sculpins Sticklebacks
Sunfish
1. Preserve up to 20 adults and juveniles (ideally several per profile). If fewer than 20
individuals are collected, voucher all of them. Voucher samples take priority over tissue
contaminant sample.
2. Retain additional individuals for tissue contaminants sample.
C Category 3 — Species of "special concern." These are state or federally listed species.
1. Photograph as in Step 1.A.2 and then release immediately.
2. If specimens have died, proceed to Step 2 and include them as part of the voucher sample.
Flag the species with an "F" series flag on the Vertebrate Collection Form and note it is a listed
species in the comments section of the form. Notify the appropriate state officials as soon as
possible. :
3. Place the voucher specimens in a bucket with two carbon dioxide tablets (e.g., Alka Seltzer®),
and a small volume of water. When specimens are anaesthetized, transfer them to a nylon
mesh bag. Record the number of individuals included in the voucher sample in the
"Vouchered Count" field for the species on the Vertebrate Collection Form.
4. Select a "FISH-BAG" tag that has the same ID number (barcode) as the voucher sample jar
(Step 3). Record the tag number.in the "Tag No." field on the corresponding line for the
species on the Vertebrate Collection Form. Place the tag into the mesh bag and seal.
(continued)
-------
Table 10-4. Continued.
5. Immediately place the bag into a container (° or 1 gal plastic jar) large enough to hold all
voucher specimens and half-filled with 10% formalin. Use additional jars if necessary to
avoid tight packing and bending of voucher specimens.
6. Repeat Steps 1 through 4 for all species collected.
7. Prepare two' 'FISH-JAR'' labels (each having the same ID number [barcode]) by filling in the
stream ID and the date of collection. Place one label into the sample jar. Cap tightly and seal
with plastic electrical tape.
8. Attach the second label to the outside of the sample container by covering it with a strip of
clear tape. Record the voucher sample ID number (barcode) on page 1 of the Vertebrate
Collection Form. Record general comments (perceived fishing efficiency, missed fish, gear
operation, suggestions) in blank lines of form. NOTE: If more than one jar is required, use
labels that have the same ID number printed on them and flag.
9. Place the preserved sample in a suitable container with absorbent material. Store the
container in a well-ventilated area during transport. Follow all rules and regulations
pertaining to the transport and shipment of samples containing 10% formalin.
FISH - JAR
SITE ID: ORRV_9 _£.-
DATE: % I vJ/98
229001
FISH - BAG
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229001
Figure 10-4. Completed voucher sample label and specimen bag tag for aquatic vertebrates.
serve vouchers of sculpins, lampreys and
other difficult species from throughout the
reach.
Place specimen bags together into a large
plastic sample container. Preserve voucher
specimens with a 10% formalin solution. See
Section 3 for instructions for preparing a buff-
ered formalin solution. Larger voucher speci-
mens (total length > 160 mm) should be slit on
the lower abdomen of the RIGHT side to allow
for complete fixation of internal tissues and or-
gans. If a fish is too large for ajar, photograph
and place in bag on ice. Flag on recording sheet;
freeze at lab separately from tissue. Start with a
concentrated solution of formaldehyde and di-
lute to the final volume with water. The final vol-
ume of 10% formalin in the sample container
should equal the total volume of specimens. Use
-------
additional containers if necessary and avoid tight
packing of specimen bags.
Delays in carrying out the anaesthetiza-
tion and preservation procedures, overpack-
ing a. sample container, or an inadequate vol-
ume of preservative will produce unidentifiable
specimens.
Formaldehyde (37%) and formalin (10%
formaldehyde by volume) are extremely caustic
agents and may cause severe irritation on con-
tact of vapors or solution with skin, eyes or
mucus membranes. It is a potential carcinogen.
Contact with vapors or solution should be
avoided. Wear gloves and safety glasses and
always work in a well-ventilated area. In case
of contact with skin or eyes, rinse immediately
with large quantities of water. Store stock solu-
tion in sealed containers in safety cabinet or
cooler lined with vermiculite. If possible, trans-
port outside of the passenger compartment of a
vehicle.
A set of two sample labels is completed
for each sample container as shown in Figure
10-4. Place one label inside each sample con-
tainer, and attach the second label to the out-
side of the jar with clear tape. Record the sample
ID number on the Vertebrate Collection Form
as shown in Figure 10-1. Carefully complete
the collection form at each transect. Tag num-
bers must be linked to each species, and each
bag of species. Be careful with fish names and
their spellings (computers see errors as differ-
ent species). Some museums may also require
that a separate collection card be completed and
inserted into each jar of voucher specimens.
10.3 Equipment and
Supplies
Figure 10-5isachecklistof equipmentand
supplies required to conduct protocols described
in this section. This checklist may differ from
the checklists presented in Appendix A, which
are used at a base site to ensure that all equip-
ment and supplies are brought to the stream site.
Field teams are required to use the checklist
presented in this section to ensure that equip-
ment and supplies are organized and available
to conduct the protocols efficiently.
-------
Equipment and Supplies for Aquatic Vertebrates
Qty. Item
1
3
2
1
1
1
2
1
1
2
2
1
1
2
many
1
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1
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1
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10+extras
1+ extras
1
1 set
1 4 ft. boat with frame mounted electrof ishing gear (anodes, cathodes, control box)
Oars (1 as extra)
Dip nets, long handled
Dip net, short handled
Generator and filled gas can; jrag
Fire extinguisher .
Anodes and cathodes (Spare)
Livewell cooler
Measuring board and ruler
Pesola scales for weighing primary tissue samples
Buckets (5 gallon)
Seat cooler with ice
Air pump, hose and fitting
Personal floatation devices
Boat straps and ropes
Tool box (Leatherman, duct tape, spare oarlock, straps, electrical tape, fuses, zipties)
Aluminum foil
Dry bags
Boat repair kit
First aid kit
Taxonomic reference books and keys for fishes of the region
List of vertebrate species common names (and species codes, if required)
List of external anomaly codes
Small nylon mesh bags for holding voucher specimens (bags can also be
constructed from sections of nylon stockings or panty hose)
Small fillet knife or scalpel for preparing larger voucher specimens for preservation
1/2- or 1 -gallon screw-top plastic jars for voucher sample
10% (buffered) formalin solution
Cooler to hold formalin solution and preserved voucher sample jars
Safety glasses
Chemical-resistant gloves
Topographic map(s)
Laser rangefinder, stopwatch, camera and film, whistles
Pruning saw and sheath
Rubber Linesman gloves, clipboards, polarized glasses, ziplock bags
Carbon dioxide tablets (Alka-Seltzer® or equivalent)
Sheet of pre-printed jar labels (4) and voucher bag tags (36), all with same preprinted
sample ID number (barcode)
Scissors for cutting labels
Plastic electrical tape
Clear tape strips
Soft lead pencils for recording data and completing tags
Fine-tipped indelible markers for completing sample labels
Vertebrate Collection Form
Vertebrate Length Recording Form
Field operations manual
Laminated sheets of aquatic vertebrate procedure tables and/or quick reference guides
Figure 10-5. Equipment and Supplies for Aquatic Vertebrates.
-------
10.4 Literature Cited
Lagler, K.R. 1956. Freshwater Fishery
Biology. 2nd. Edition. William C. Brown
Co., Dubuque, Iowa.
McCormick, F.H. 1993. Fish. pp. 29-36 IN:
R.M. Hughes (ed.). Stream Indicator
Workshop. EPA/600/R-93/138. U.S. En-
vironmental Protection Agency, Corvallis,
Oregon.
ill
-------
Section 11
Fish Tissue Contaminants
James M. Lazorchak1, Frank H. McCormick1, Robert M. Hughes2, and
Spence A. Peterson3
In addition to gathering data on the
aquatic vertebrate assemblage (Section 10),
fish are retained for analysis of tissue contami-
nants. In general, the focus is on fish species
that commonly and occur throughout the re-
gion of interest, and that are sufficiently abun-
dant within a sampling reach. The fish tissue
contaminants indicator, which measures bio-
accumulation of persistent toxics, is used to
estimate regional risks of consumption to fish
predators, either wildlife or human. Various
studies that have been done on fish tissue con-
taminants have focused on different parts of
the fish: whole fish, fillets, livers. EMAP-SW
will focus on whole fish because of its em-
phasis on the ecological health of the whole
'U.S. EPA, Natioinal Exposure Research Laboratory, Eco-
logical Exposure Research Division, 26 W. Martin Luther
King Dr., Cincinnati, OH 45268.
2Dynamac International Corp., 200 SW 35th St., Corvallis,
OR 97333.
3U.S. EPA, National Health and Environmental Effects Labo-
ratory, Western Ecology Division, 200 S W 35th St., Corvallis,
OR 97333.
stream (as opposed to a focus on human health
concerns). Whole fish are a good ecological
indicator and a better indicator of risk to pis-
civorous wildlife than fillets. It is hoped to
also be able to say something about risks to
human health by analyzing whole fish. Whole
fish also present fewer logistical problems for
field crews (no gutting required in the field)
and the analytical lab (no filleting necessary).
For the fish contaminants indicator in
EMAP-SW STREAMS, an attempt was
made to collect two fish samples at as many
sites as possible. One sample, of Primary Tar-
get Species, was stream fish whose adults are
small (in Mid-Atlantic streams examples are:
dace, chub, sculpins, stqnerollers, shiners, and
darters). The second sample, where available,
of a Secondary Target Species, was a spe-
cies whose adults are of larger size (In Mid-
Atlantic streams examples are: bass, trout, sun-
fish, suckers, carp). In addition to being more
ubiquitous than the larger fish (and therefore
more likely to be present in sufficient num-
bers to composite), small fish have other ad-
-------
vantages over large fish. Most importantly, it
may be possible to get a more representative
sample of the contaminant load in that stream
section (although it would be at a lower ex-
pected level of bioaccumulation) by
compositing say, in the range of 20 to 200
small fish individuals than by compositing 3
to 5 large fish. Small fish may be a more ap-
propriate indicator for assessing ecological
risk, as they might be expected to be prey for
a larger number of fish-eating animals (the
majority of which will be piscivorous birds
and small mammals). The major advantage
that larger fish could potentially offer, whether
predators (piscivores) or bottom feeders, is a
higher level of bioaccumulation and thus
greater sensitivity to detect contaminants. The
relative bioaccumulation of contaminants by
large and small stream fish is not known, thus
the reason for having Primary and Second-
ary Target Species in this study.
In trying to answer these questions, the
field crews' efforts to apply the protocol for
sampling, handling and shipping in a consis-
tent manner are critical. The diligence of
the field crews in following the protocols is
especially important in a status and trends
study such as EMAP-SW where it is critical
to get a standard sample from each site so
that there is confidence that differences seen
over time and between sites represents varia-
tions in the ecosystems and not differences in
sampling and handling between the crews.
Suggestions from field crew members on how
the protocol can be improved are welcomed
and will be incorporated to improve them, but
protocols should be followed as written until
official changes are made.
11.1 Selecting Fish
Tissue Specimens
If possible, obtain one sample each, of
the desired weight or number (see below)
of similarly sized* individuals, from the Pri-
mary and Secondary target species lists (2
composite samples total). To judge if the
proper amount of a target species is present
in the fish catch, weight will be used for pri-
mary target species and number of individu-
als of sufficient size will be used for second-
ary target species.
I. Primary Target Species
Small adult fish
(in priority order)
Weight
l)BlacknoseDace
2) Another Dace species
3) Creek Chub or Fallfish
4) Slimy Sculpin/Mottled
Sculpin
5) Stoneroller
6) A Darter species
7) A Shiner species
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
A) Choose the highest priority target
species from the above list, that has at least
enough individuals to attain the minimum
weight (50 g). Get as much weight offish as
possible within the desired weight range (50-
400 g).
B) If less than the desired weight of any
primary target species is collected, send indi-
viduals of a small nontarget species if 50 g or
more are available.
* - The general rule-of-thumb for simi-
lar size is that the smallest individual in the
sample should be at least 75% of the length
of the largest individual. This rule applies to
both primary and secondary target species.
Crews just need to keep this criterion in mind
while selecting the final sample. Any obvi-
ously small or large individuals should not be
kept if there is a sufficient sample to return
-------
without them. If there is a conflict between
criteria, getting a sufficient sample is a higher
priority than getting similar-sized individuals.
** - This weight represents the mini-
mum amount of tissue needed for laboratory
analysis. Crews should not settle for the
minimum amount (weight) if more fish are
present, but instead send as many fish as pos-
sible up to the 400 g weight goal.
II. Secondary Target Species
Collect and save a sample of secondary
target species if such a sample of desired num-
ber of individuals of desired size is available.
Collect similar sized individuals if enough
are present.
Larger adult fish
(in priority order)
Desired Desired
Size Number
1) A Bass species
2) A Trout species*
3) A Sunfish species
4) Catfish
5) White sucker
6) Hogsucker
7) Carp
120mm
120mm
120mm
120mm
120mm
120mm
120mm
5
5
5
5
5
5
5
* - Collect only those trout that appear
not to be recently stocked.
A) If fewer than the desired number
of secondary target species individuals of de-
sired size are collected, add smaller individu-
als of the same species, if available, to achieve
the desired number (5).
B) If fewer than 5 fish of any size are
available, you may send as few as 3 fish that
are at least at or near the minimum desired
size (120 mm).
C) If an acceptable secondary target spe-
cies sample (by the above criteria) is not avail-
able send only the primary target species
sample. If neither a primary or secondary spe-
cies sample that meets these criteria is avail-
able, use your best judgement in sending some
type offish sample.
11.2 Preparing
Composite Samples for
Primary and Secondary
Target Species
To determine the proper quantity for
each composite sample, weight is used for the
primary target species and the number of in-
dividuals of sufficient size is used for the sec-
ondary target species. Prepare each compos-
ite sample using similar sized individuals if
possible, but getting a sufficient sample is a
higher priority than getting similar-sized in-
dividuals.
Prepare a primary sample as described
in Table 11-1. Choose a species that has at
least enough individuals to attain the mini-
mum weight (50 g). Send as many fish as
possible up to the 400 g weight goal. If there
is no single species with enough individuals
available, prepare the sample using individu-
als of multiple species.
Prepare a secondary sample as described
in Table 11-1. Choose a species that has 5
similar-sized individuals (minimum total
length =120 mm) available. If fewer than 5
fish of any size for any secondary species are
available, prepare the composite sample us-
ing as few as 3 fish that are at least at or near
the minimum desired size.
If neither a primary nor secondary
sample is available, use your best judgement
to obtain some type offish tissue sample from
the available species collected. Use the pro-
cedure for either primary or secondary spe-
-------
Table 11-1. Procedure to Prepare Fish Tissue Samples.
Note: If neither a primary nor secondary species sample is available, use your best judgement in sending
some type of composite fish tissue sample.
Primary Sample (PI
After all voucher specimens have been prepared, choose a primary species that has enough similarly sized
individuals to weigh to 400 g (smallest to largest should not differ by more than 25% in length).
Secondary Sample (S)
: After all voucher specimens have been prepared, select a large secondary species that has at least 5
._ individuals 120 mm. Include similar sized individuals if available (smallest to largest should not differ by more
than 25% in length).
: 1. Place the fish into a bucket with two carbon dioxide tablets (e.g.," Alka Seltzer®") and a small volume
of water. After they have been anaesthetized, use clean hands to transfer them to aluminum foil.
2. Prepare a clean work surface to prepare the primary composite sample. Keep hands, work surfaces, and
wrapping materials clean and free of potential contaminants (mud, fuel, formalin, sun screen, insect
repellant, etc.)
;; 3-P. For primary samples, record the common name (from a standardized list) of the species, its species
code (if required), and the number of individuals in the .sample in the appropriate fields on line "P1" of
the Sample CollectionForm (Figure 11-1).
• 3-S. Measure the total length (TL) of each secondary individual. Record the common name (from a
standardized list) of the secondary target species, its species code (if required), and the total length for
each individual on lines S1 through S5 in the secondary sample section of the Sample Collection Form.
« 4. If the individuals included in composite samples were collected from throughout the sampling reach,
place an "X" in the "Yes" box in the sample section of the Sample Collection Form. If the individuals
;; were only collected from a limited segment of the sampling reach, place an "X" in the "No" box and
explain in the "Explain" field on the form.
5-P. Wrap all primary fish together in a single piece of aluminum foil, making sure the dull side of the
aluminum foil is in contact with the fish. Place the sample in a self-sealing plastic bag.
5-S. Wrap each fish of the secondary sample separately in aluminum foil, with the dull side of the foil in
: contact with the fish. Place ajll the wrapped individuals into a single self-sealing plastic bag.
, 6. Expel excess air and seal the bag. Wrap clear tape around the bag to seal and make a surface for each
sample label..
7-P. Prepare two Fish Tissue sample labels (each having the same sample ID number [Figure 11-2]) by filling
in the stream ID and the date of collection. Circle "PRIMARY" on each label. Record the sample ID
number (barcode) in the primary sample section of the Sample Collection Form.
, 7-S. Prepare two Fish Tissue sample labels (each having the same sample ID number [Figure 11 -2]) by filling
: in the stream ID and the date of collection. Circle" SECONDARY" on each label. Record the sample ID
number (barcode) in the secondary sample section of the Sample Collection Form.
: 8, Attach the appropriate label to the tape surface of the bag. Cover the label with a strip of clear tape.
Place the labeled bag into a second self-sealing plastic bag. Seal the bag and attach the second label
to the outside of the appropriate bag. Cover the label with a strip of clear tape.
9. Place the double-bagged sample into a cooler containing bags of ice until shipment. Keep the sample
frozen until shipment.
-------
eies, depending upon the species used and the
size range of individuals selected.
Individuals comprising the primary com-
posite sample are wrapped together in alumi-
num foil and placed into a single plastic bag.
Each individual comprising the secondary
composite sample is wrapped separately, but
all individuals are placed into a single plastic
bag. Each composite sample is labeled as
shown in Figure 11-1. Prepare two identical
labels for each composite sample. Double-bag
each sample, and place a label on each bag.
Record information about each composite
sample on page 2 of the Sample Collection
Form (Figure 11-2). Make sure the sample
ID numbers (barcodes) recorded on the col-
lection form match those on the sample la-
bels.
Tissue samples are stored in a cooler with
several bags of ice. When using ice, double
bag the ice and tape the last bag shut to pre-
vent contamination of samples by melting ice.
Store tissue samples frozen until they can be
shipped (Section 3). Tissue samples can be
stored and shipped with other samples requir-
ing freezing (periphyton chlorophyll, periphy-
ton biomass, periphyton APA, and sediment
metabolism samples).
11.3 Equipment and
Supplies
Figure 11-3 is a checklist of equipment
and supplies required to conduct protocols
described in this section. This checklist may
differ from the checklists presented in Appen-
dix A, which are used at a base site to ensure
that all equipment and supplies are brought
to and are available at the river site. Field
teams are required to use the checklist pre-
sented in this section-to ensure that equipment
and supplies are organized and available to
conduct the protocols efficiently.
FISH TISSUE
SITE ID: ORRV_ig.-7 9. 9.
DATE: JT/,51/98
/—*• N
SAMPLE:(PRIMARY)SECONDARY
229000
FISH TISSUE
SITE ID: ORRV _3_ S.-3.3. _2
DATE: ST/>5/98
SAMPLE: PRIMARY (SECONDARY
229000
Figure 11-1. Completed sample labels for fish tissue contaminants.
-------
Reviewed by (initial)
SAMPLE COLLECTION FORM - RIVERS (continued)
SITE NAME: BEAVE"R RlV£/^ DATE: % >5 1 98 VISIT D0jg]l D2 D3
SITEID: , ORRV Jl JL-J?.3_2_ TEAM ID (X): JS1 D2 D3 D4 D5
D 6 D7 D8
CHEMISTRY AND MICROBIAL WATER SAMPLE (Chem: 4-L Cubitainer and 2 Syringes, Micro: Glass Bottle)
SAMPLE ID (BARCODE) TRANSECT FLAG COMMENTS
CHEMISTRY 2/r-3°'Z K
MICROBIAL .2^3)13 K
SEDIMENT TOXICITY SAMPLES
SAMPLE ID (BARCODE) FLAG
COMMENTS
FISH TISSUE SAMPLES - PRIMARY SAMPLE (min. 50g total wgt)
SAMPLE ID (BARCODE) ->-
LINE
PI
SPECIES CODE
CC7TT Z.5ZZ.2L
2*32.2.3
COMMON NAME NUMBER OF
UNICVOWN CDTTfA i
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X) ->•
NOIVIDUALS FLAG
1
G YES JST NO
IFNO.EXPLAIN: COLLECTED AT OWE PROFILE
FISH TISSUE SAMPLES - SECONDARY SAMPLE (where available; 5 individuals)
SAMPLE ID (BARCODE) -*•
LINE
SI
S2
S3
84
Si
SPECIES CODE
PTYC-ORe 6-
PTYCORT&
PTYCOREG-
PTYCOREG-
PTYCOREG
243^2^
COMMON NAME TOTAL LE^
NORTHERN soi/Aynsw 22
NORTHERN; SQI/AWFISH 25
/\|ORTHEI?IV SQUAWFJSH 2t
A/ORTHERN SOUAWFISH 2C
NORTHERN 50L/A1A/FJSH 2
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS QOLLECTED FROM THROUGHOUT REACH? (X) ->•
JGTH (MM) FLAG
0
!0
?D
IS
^ YES 0 "°
IF NO, EXPLAIN:
LINE
COMMENT OR FLAG EXPLANATION FOR FISH TISSUE
Flagaxfe*.' K-Sample not collected: U- Suspect sample: Fl,F2,etc.« misc. Hag assigned by field crew. Explain all Hags in Comments sections.
Rev. 05/29/98 (_rvscmb.98) SAMPLE COLLECTION FORM - RIVERS - 2
Figure 11-2. Sample Collection Form, showing information recorded for fish tissue samples.
-------
Qty.
Equipment And Supplies For Fish Tissue Contaminants
Item
1
4
1 roll
1
Iroll
4
1
2 sets
Ipkg.
1
1 copy
1 set
Plastic bucket for anesthetization
Carbon dioxide tablets (Alka-Seltzer® or equivalent)
Clear tape for sealing tissue sample bags
Pesola® portable scale, precision ±5g
Aluminum foil
1 -gallon self-sealing plastic bags
Sample Collection Form
Fish tissue sample labels (each set with a different sample ID number [barcode])
Clear tape strips
Soft (#2) lead pencils to record data
Fine-point indelible markers to fill out labels
Cooler with ice (double-bagged and taped)
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for fish tissue
contaminants
Figure 11-3. Equipment and supplies checklist for fish tissue contaminants.
-------
-------
Section 12
Visual Stream Assessments
Alan T. Herlihy1
After all other samples and field data have
been collected, the field team makes a gen-
eral visual assessment of the river, and per-
forms a final check of the data forms and
samples before leaving the river site (see Sec-
tion 13). The objective of the visual river as-
sessment is to record field team observations
of catchment and stream characteristics that
are useful for data validation, future data in-
terpretation, ecological value assessment, de-
velopment of associations, and verification of
stressor data. The observations and impres-
sions of field teams are extremely valuable.
12.1 Visual Stream
Assessment
The objective of the visual river assess-
ment is to record field crew observations of
catchment/river characteristics useful for fu-
ture data interpretation, ecological value as-
sessment, development of associations, and
'Dept. of Fisheries and Wildlife, Oregon State University,
c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
verification of stressor data. Observations and
impressions of field crews are extremely valu-
able. Thus, it is important that these observa-
tions about river characteristics be recorded
for future data interpretation and validation.
The assessment form is designed as a tem-
plate for recording pertinent field observa-
tions. It is by no means comprehensive and
any additional observations should be re-
corded in the Comments section of the form.
Complete the assessment form after all
other sampling and measurement activities
have been completed. Take into account all
observations the sampling team has made
while at the site. The assessment includes the
following components: watershed activities
and observed disturbances, reach character-
istics, waterbody character, general assess-
ment, and local anecdotal information. The
procedure for conducting the visual assess-
ment of the sampling reach is presented in
Table 12-1. Record data and observations for
each component of the assessment on the
Assessment Form as shown in Figures 12-1
and 12-2.
-------
Table 12-1. Procedure for Conducting the Final Visual Assessment of a River.
1. After all other sampling and measurement activities are completed, fill out the header section of an
Assessment Form. Use your perceptions obtained during the course of the day, while at the river or
driving/walking through the catchment to complete the remainder of the form.
2. Watershed Activities and Disturbances Observed: Rate each type of activity or disturbance listed on
the form as either "Not observed", "Low", "Medium", or "High", and record the rating on the
Assessment Form. Keep in mind that ratings will be somewhat subjective and that an extensive effort
to quantify the presence and intensity of each type of stressor is not required. General categories of
activities and types of disturbance are described below:
• Residential: The presence of any of the listed disturbances adjacent to or near the river.
• Recreational: The presence of organized public or private parks, campgrounds, beaches or other
recreation areas around the river. If there are signs of informal areas of camping, swimming or
boating around the river (e.g., swimming hole), record them as "primitive" parks, camping.
• Agriculture: The presence of cropland, pasture, orchards, poultry, and/or livestock.
• Industrial: Any industrial activity (e.g., canning, chemical, pulp), commercial activity (stores,
businesses) or logging/mining activities around the river or in the catchment. Describe in more,
detail in the comments section.
• Management: Any evidence of liming activity, water treatment, dredging or channelization, flow
control structures, etc.
Any oddities, or further elaboration should be recorded in the Comments section.
3. Reach Characteristics: For each type of riparian vegetation cover or land use category listed on the
Assessment Form, estimate the proportion of the sampling reach immediately adjacent to the river that
is affected. Place and "X" in the appropriate extent class box (Rare [<5%],Sparse [5 to 25%], Moderate
[25 to 75%], and Extensive [> 75%]) on the form.
4. Classify the overall water clarity within the sampling reach as clear, murky, or highly turbid. Place an
"X" in the appropriate box on the "Water Clarity" line of the Assessment Form. If you believe that
water clarity has been influenced by a recent storm event, also place an "X" in the "Storm Influenced"
box.
5. Water Body Character: Assign a rating of 1 (highly disturbed) to 5 (pristine) based on your general
impression of the intensity of impact from human disturbance. Place an "X" in the box next to the
; assigned rating on the Assessment Form.
' 5. Waterbody Character (cont.): Assign a rating to the river based on overall aesthetic quality, based on
your opinion of how suitable the river water is for recreation and aesthetic enjoyment today. Place and
"X" in the box next to the assigned rating on the Assessment Form.
5. Beautiful, could not be any nicer.
4. Very minor aesthetic problems; excellent for swimming, boating, enjoyment.
3. Enjoyment impaired.
2. Level of enjoyment substantially reduced.
1. Enjoyment nearly impossible.
Add any comments you feel might aid data interpretation in the Comments Section.
6. General Assessment: record comments on wildlife observed, perceived diversity of terrestrial
vegetation, and the estimated age class of forest (0 to 25 yr, 25 to 75 yr, or > 75 yr.) on the Assessment
Form.
7. Local Anecdotal Information: Record any information regarding the past or present characteristics or
condition of the river provided by local residents.
Each watershed activity or disturbance
is rated into one of four categories of abun-
dance or influence: not observed, low, me-
dium, or high. Leave the line blank for any
activity or disturbance type not observed. The
distinction between low, medium, and high
will be subjective. For example, if there are
2-3 houses on a river, the rating for "Houses"
would be low. If the river is in a suburban
housing development, rate it as high. Simi-
-------
Reviewed by (initial) A/C4-
ASSESSMENT FORUM - STREAMS/RIVERS
SITE NAME:
RlV£/?
DATE: g" IS I 98 VISIT DO JS1 D2 D3
SITE ID: ORRV
TEAM ID (X):
D2 D3 Q4 D5 D6 D7 D8
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED (INTENSITY: BLANK = NOT OBSERVED, L = LOW, M = MODERATE. H . HEAVY)
RESIDENTIAL
RECREATIONAL
AGRICULTURAL
INDUSTRIAL
STREAM MANAGEMENT
LJM H RESIDENCES
L M H PARKS. CAMPGROUNDS
L M H CROPLAND
L M H INDUSTRIALPLANTS
L M H LIMING
L M H
MAINTAINED LAWNS
PRIMITIVE PARKS. CAMPING
L M H
L M H
MINES/QUARRIES
L M H
DRINKING WATERTREATMENT
L M
CONSTRUCTION
TRASH/LITTER
LIVESTOCK USE
L M H
OIL/GAS WELLS
L M(H)
ANGLING PRESSURE
L M
L M H
SURFACE FILMS. SCUMS. OR SLICKS
L M H
L M H
POWER PLANTS
L M H
L M H
L M H
L M H
L M H
CHANNELIZATION
L M H
IRRIGATION PUMPS
L M H
EVIDENCE OF FIRE
1M H
WATER LEVEL FLUCTUATIONS
BRIDGE/CULVERTS
L M H
FISH STOCKING
REACH CHARACTERISTICS (percent of reach)
FOREST
SHRUB
GRASS
WETLAND
BARE GROUND
MACROPHYTES
n
SPARSE (5 TO 25%)
n
MODERATE (25TO 75%)
n
EXTENSIVE C>75%)
n
n
SPARSE (5 TO 25%)
MODERATE (25 TO 75 75%)
n
n
SPARSE (5TO25%)
MODERATE (25 TO 75%)
n
EXTENSIVE > 75%)
n
SPARSE (5 TO 25%)
[3 m
IODERATE(25T075%)
n
EXTENSIVE (> 75%)
\[
SPARSE (5 TO 25%)
n
MODERATE (25 TO 75%)
n
EXTENSIVE (> 75%)
SPARSE (5 TO 25%)
MODERATE (25 TO 75%)
n
EXTENSIVE (> 75%)
AGRICULTURE - ROW CROP
AGRICULTURE - GRAZING
LOGGING
DEVELOPMENT (RESIDENTIAL & URBAN)
D
SPARSE {5 TO 25%)
n
MODERATE (25TO 75%)
n
EXTENSIVE (> 75%)
n
SPARSE (5 TO 25%)
MODERATE (25 TO 75%)
n
EXTENSIVE (> 75%)
n
SPARSE (5TO 25%)
n
MODERATE (25 TO 75%)
n
EXTENSIVE (> 75%)
SPARSE (5 TO 25%)
MODERATE (25 TO 75%)
n
EXTENSIVE (> 75%)
WATER CLARITY
n
HIGHLY TURBID
a
STORM INFLUENCED
WATERBODY CHARACTER (X ONE) -
PRISTINE
HIGHLY DISTURBED
|g|4
n
UNAPPEALING
aciNtHAL A&afcbbMfcN I (wilalite, vegetation diversity, forest age class (0-25 yrs, 25-75 yrs, >75)
ROCKY CANYON LANDS, SAGE/G-RASS COVER - OCCASIONAL RlMRlAM TREE.
NO LWA LIMITED FfSH COVER/ REGULATED FLOW /WATER LEVEL.
LUUAL ANtODOTAL INFORMATION:
Rev. 05/29/98 (strvasse.98)
ASSESSMENT FORM - STREAMS/RIVER - 1
Figure 12-1. Assessment Form (page 1).
-------
Reviewed by (initial)
ASSESSMENT FORUM - STREAMS/RIVERS (continued)
SITE NAME:
DATE:
798 VISIT DO D1 D2 EH3
SITE ID: O R R V
TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
Rev. 05/29/98 (strvasse.98)
ASSESSMENT FORM - STREAMS/RIVER - 2
Figure 12-2. Assessment Form (page 2).
-------
larly, a small patch of clear cut logging on a
hill overlooking the river would be rated as
low. Logging activity right on the river shore,
however, would be rated as high.
When assessing reach characteristics,
make your best estimate as to the percent of
the sampling reach (100 or 40 channel widths)
that had each type of listed riparian zone land
use immediately adjacent to the river. Also rate
the water clarity, including whether you be-
lieve the clarity is influenced by recent storm
events (see Section 4).
Water body character is defined as "the
physical habitat integrity of the water body,
largely a function of riparian and littoral habitat
structure, volume change, trash, turbidity,
slicks, scums, color, and odor." Water body
character is assessed using two attributes, the
degree of human development, and aesthet-
ics. Rate each of these attributes on a scale of
1 to 5. For development, give the river a "5"
rating if it is pristine, with no signs of any
human development. A rating of "1" indicates
a river which is totally developed (e.g., the
entire river is lined with houses, or the ripar-
ian zone has been removed). For aesthetics,
base your decision on any faptor about the
river that bothers you (e.g., trash, algal
growth, weed abundance, overcrowding).
The general assessment component in-
cludes any observations that will help in data
interpretation in the pertinent section. Gen-
eral assessment comments can include com-
ments on wildlife observed, diversity of ter-
restrial vegetation, age class of forest, or any
other observation. Comments from locals are
often useful and should be recorded in the
"Local Anecdotal Information" section. The
back side of the form (Figure 12-2) is avail-
able for general comments.
12.2 Equipment and
Supplies
Figure 12-3 is a checklist of the supplies
required to complete the rapid habitat and vi-
sual stream assessments. This checklist may
differ from the checklists presented in Appen-
dix A, which are used at abase site to ensure
that all equipment and supplies are brought
to and are available at the river site. Field
teams are required to use the checklist preT
sented in this section to ensure that equipment
and supplies are organized and available to
conduct the protocols efficiently.
Qty.
1
6
1
1 copy
1 set
Equipment and Supplies for Visual Stream Assessments
Item
Assessment Form for visual stream assessment
Soft (#2) lead pencils '
Covered clipboard or forms holder
Field operations and methods manual ~
Laminated sheets of procedure tables and/or quick reference guides for rapid
habitat and visual assessments
Figure 12-3. Checklist of equipment and supplies required for visual assessments
-------
-------
Section 13
Final Site Activities
James M. Lazorchak1 and Daniel K. Averill2
After the boat crews have safely reached
the take-out location, team members begin fi-
nal site aptivities.,Composite samples for per-
iphyton (Section 7) and benthos (Section 9)
are processed. The incubation for sediment
metabolism (Section 8) is initiated, if not al-
ready started. Equipment and supplies are
unloaded from rafts, vehicles are shuttled,
equipment and supplies are loaded in the ve-
hicles, rafts are loaded onto the trailer, and
data forms, labels, and samples are inspected.
The team leader reviews all of the data
forms and sample labels for accuracy, com-
pleteness, and legibility. A second team mem-
ber inspects all sample containers and pack-
ages them in preparation for transport, storage,
or shipment. Refer to Section 3 for details on
preparing, delivering and/or shipping samples.
When reviewing field data forms, ensure
that all required data forms for the river have
'U.S. EPA, National Exposure Research Laboratory, Eco-
logical Exposure Research Division, 26 W. Martin Luther
King Dr., Cincinnati, OH 45268.
2Dynamac International Corp., 200 SW 35th St., Corvallis,
OR 97333.
been completed. Confirm that the site identi-
fication code, the year, the visit number, and
the date of the visit are correct on all forms.
On each form, verify that all information has
been recorded accurately, the recorded infor-
mation is legible, and any flags are explained
in the comments section. Ensure that written
comments are legible and use no "shorthand"
or abbreviations. Make sure the header infor-
mation is completed on all pages of each form.
After reviewing each form, initial the upper
right corner of each page of the form. A check
by a team member that has not filled out the
sheets for a particular section might be the best
person to review the field data forms before
leaving the site.
When inspecting samples, ensure that
each sample is labeled, all labels are com-
pletely filled in and legible, and each label is
covered with clear plastic tape. Compare
sample label information with the information
recorded on the corresponding field data forms
(e.g.,"the Sample Collection Form) to ensure
accuracy.
Keep equipment and supplies organized
so they can be inventoried using the equip-
-------
ment and supply checklists presented in Ap-
pendix A. Clean up the take-out site and trans-
port all waste material out of the area.
If samples are to be shipped by FEDEX,
check the 1 800 number for the nearest Motel
or location that samples can be left for pickup.
In most eastern states, FEDEX will pick up
coolers at most motels and ship the following
morning. This can help reduce travel time to
the next sample site. FEDEX also will ship
on Saturdays for Sunday or Monday pickups
as of the year 2000. Check their 1 800 num-
ber for locations that will accept Saturday
drop off and Sunday delivery.
iiiiiiiiM
-------
Appendix A
Equipment and Supply Checklists
Field Data Forms and Sample Labels A-2
Office Supplies and Tools A-3
Personal Equipment and Supplies A-4
Chemicals A-5
Packing and Shipping Supplies... , A-5
Site Verification and Sampling Reach Layout,.- ; A-6
Water Chemistry and Microbiology .,A-6
Physical Habitat .............; ..„. ........ .... A-7
Periphyton „ A-7
Sediment Metabolism A-8
Benthic Macroinvertebrates A-8
Aquatic Vertebrates and Fish Tissue Contaminants...., A-9
-------
Field Data Forms and Sample Labels
Number
per site
Item
1 -r extras
1 1 + extras
0 4- extras
1 + extras
1
1
3
1
4
5
2
4
1
2 sets
2 copies
2 sets
Verification Form
Sample Collection Form
7ield Measurement Form
Channel/Riparian Transect Form
Thalweg Profile Form
PHAB Commentes Form
Vertebrate Collection Form
Vergebrate Length Recording Form
Assessment Form for visual river assessment
Field Sample Shipment'Tracking Form
Water chemistry labels (same ID number)
Microbial label
Periphyton labels (same ID number)
Sediment metabolism labels (different ID numbers)
Composite Benthic sample labels, with preprinted ID numbers (barcodes)
Composite Benthic sample labels without preprinted ID numbers
Sheet of preprinted aquatic vertebrate jar labels (4) and voucher bag tags
(36), all with same preprinted sample ID number (barcode)
Fish tissue sample labels (2 labels per set; each set with a different sampl
ID number [barcode])
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides
-------
Office Supplies And Tools
Number
per site
Item
1
1
1
1
1 ea.
1
1
4
12
6
Ipr
1
1
Dossier of access and general information for scheduled river site
Topographic map with "X-site," reach boundaries, and launch points
marked
Site information sheet with map coordinates, bank to be sampled, elevation
of X-site, and other general information
Portable file folder used to organize field and administrative forms
Map wheel, calculator, metric ruler, shoulder bag, and field notebook
Sampling itinerary notebook
Safety log and/or personal safety information for each team member
Covered clipboards or forms holders
Soft (#2) lead pencils
Fine-tip indelible markers
Scissors for cutting labels
Pocket knife or multipurpose tool
Toolbox with basic tools needed to maintain/repair sampling gear (other
than electrofishins equipment)
-------
Personal Equipment and Supplies
Number
per site
Item
2
6
2
2
1
3
1 pr/person
1 per person
2 pair
1
1 per person
1
2
2
1
2
2
14 ft. inflatable rafts with custom frames
Oars
Extra oar locks
Raft pump - AIR
Raft pump -ELECTRIC '
Raft patch kits
Felt-soled wading boots + neoprene booties.
PFD's with pockets
Polarized sunglasses and leather gloves
First aid kit, eye wash unit, sunscreen, whistle, antibacterial hand wash
Rain gear
Water purifier
Throw bags and dry bags
Pruning saws
5 gal. water jug
Pulaski's and shovels
Booster cables
Tie-down straps, ropes, bungee cords
.
-------
Chemicals
Number
per site
Item
Ipr
2pr
1
1
2 gal
1
2 gal
Igal
2 jars
Safety glasses
Chemical-resistant gloves
Laboratory apron, resistant to ethanol and formalin
Cooler or large tote for transporting ethanol and samples
95% ethanol
Cooler or large tote for transporting formaldehyde/formalin
10% (buffered) formalin solution
Gasoline for electrofishing generator in approved container
Methyl Ethyl Ketone (MEK) for patching rafts
Number
per site
Packing and Shipping Supples
Item
2 bags
1 box
1 box
Iroll
2pkg
4 rolls
3
1
1
2
6
Ice
1-gal heavy-duty self-sealingn (e.g., with a zipper-type closure) plastic bags
30-gal plastic garbage bags
Clear tape for sealing tissue sample bags and shipping containers
Clear tape strips for covering labels
Plastic electrical tape
Insulated shipping containers for samples (heavy plastic coolers)
Plastic container with snap-on lide
Cooler with bags of ice to store frozen samples
Containers suitable to transport and/or ship samples preserved in formalin or
ethanol (coolers)
Shipping airbills and adhesive plastic sleeves
-------
Site Verification and Sampling Reach Layout
Number
per site
Item
1
1
1
1
1
1
1 ea
GPS receiver and operating manual
Extra batteries for GPS receiver
Laser rangefinder (400 yard range) and clear
waterproof bag (dry bag)
50 m fiberglass measuring tape with reel
Dossier of site and access information
Waterproof camera and film
Topographic map and gazetteer
Map wheel, calculator, metric ruler
Number
per site
Water Chemistry and Microbiology
Item
1
1
1
1
1
1
1
1
1
2
1
1
2
Dissolved oxygen/Temperature meter with probe, manual, & storage case
DO repair kit containing additional membranesand probe filling solution
Conductivity meter with probe, operating manual, and padded storage case
Extra batteries for dissolved oxygen and conductivity meters
500-mL plastic bottle of conductivity QCCS labeled "Rinse" (in plastic bag)
500-mL plastic bottle of conductivity QCCS labeled "Test" (in plastic bag)
500-mL plastic bottle of deionized water to store conductivity probe
Field thermometer
500 mL plastic beaker with handle (in clean plastic bag)
4-L cubitainer •
60-mL plastic syringes
200 mL square glass microbial bottle
Plastic container with snap-on lid to hold filled syringes
Syringe valves
-
-------
Physical Habitat
Number
per site
Item
1
1
1
1
Iroll
1
1
Surveyor's telescoping rod (round profile, metric scale, 7.5 m extended) for depth
measurements and substrate estimation
Clinometer (or Abney level) with percent and degree scales
Convex spherical canopy densiometer, modified with taped "V"
Bearing compass (Backpacking type)
Colored surveyor's plastic flagging
Meter stick for bank angle measurements
SONAR depth sounder - narrow beam (16 degrees)
Number
per site
Periphyton
Item
1
1
1
1
1
2
1
4
1 box
1 pair
1
1
1
2
1
1
Large funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle Toothbrush with handle bent at 90° angle
1-L wash bottle for stream water
1-L wash bottle containing deionized water
500-mL plastic bottles for composite samples
60 mL plastic syringe with a 3/8" hole bored into the end
50-mL screw-top centrifuge tubes (or similar sample vials)
Glass-fiber filters for chlorophyll and biomass samples
Forceps for filter handling
25-mL or 50-mL graduated cylinder
Filtration unit, including filter funnel, cap, filter holder, and receiving chamber
Hand-operated vacuum pump with length of flexible plastic tubing
Aluminum foil squares (3" x 6")
Small syringe or bulb pipette for dispensing formalin
Collapsible bucket
-------
Sediment Metabolism
Number
per site
Item
1
1
1
1 set
1
5
1
1
1
Small scoop sampler for sediments
Wide-mouthed plastic jar labeled "COMPOSITE SEDIMENT SAMPLE". If
sediment is only being collected for metabolism samples, a 250-mL jar is
sufficient.
YS1 Model 95 Dissolved Oxygen meter
Spare batteries for DO meter
Small plastic spoon or spatula to transfer sediment from the composite sample
container to respiration tubes
50-mL, screw-top, centrifuge tubes
50-mL screw-cap centrifuge tube labeled "BLANK"
Small cooler used as incubation chamber
l,OOQ-mL plastic beaker to holding centrifuge tubes during incubation
Number
per site
Benthic Macroinvertebrates
Item
1
2
1
2pr
1
1
1
10
2
1
Ipkg
Modified kick net (closed bag with 595 pm mesh) and 4-ft handle (Wildco #425-
J50-595)
Spare net(s) for the kick net sampler or extra sampler
Drift nets, 595 jam mesh, closed end
Sieve bucket, 595 (jm mesh openings
Watchmakers' forceps
Wash bottle, 1-L capacity.
Small spatula, spoon, or scoop to transfer sample
Funnel, with large bore spout
Sample jars, plastic with screw caps, 500 mL and 1 L capacity, suitable for use
wfth ethanol
Buckets, plastic, eight to ten quart capacity
Stopwatch
Kimwipes in small self-sealing plastic bag
-------
Aquatic Vertebrates and Fish Tissue Contaminants
Number
per site
Item
1 set
1
2
1
2
1
1
Ipr
2
2
2
1
1 set
1
1
15-20
1
2 ea
1
2 gal
4
1 roll
Electrofishing equipment - cathode and anode droppers, other frame mounted
electrical boxes and connections
Electrofishing control box with an connectors
Anodes and cathodes (SPARE)
Generator and filled gas can + rag
Dip nets, Long handled
Dip net, Short handled
Live weH cooler
Heavy-duty rubber gloves for electrofishing
Fish measuring board and rulers
Portable scafe, precision ±5g to weigh tissue samples
Buckets (5 gallon)
Tools for electrofishing assembly
Fire extinguisher
Taxonomic reference books and keys for fishes and amphibians of the region
List of vertebrate species codes and common names
List of external anomaly codes
Small nylon mesh bags for holding voucher specimens (bags can also be
constructed from sections of nylon stockings or panty hose)
Small fillet knife or scalpel for preparing larger voucher speciments for
preservation
1, 2 or 4 L screw-top plastic jars for voucher samples
Plastic bucket for anesthetization
10% (buffered) formalin solution
carbon dioxide tablets (Alka-Seftzer® or equivalent)
Aluminum foil
-------
-------
Appendix B
Quick Reference Guides
The following pages are tabular summaries of different field activities and procedures
described in this manual. These were developed by the principal investigators for each eco-
logical indicator to provide a field team with a quick way to access information about each
procedure. They are intended to be laminated and taken to the river site after the crew has
been formally trained in the detailed procedures as presented in the manual. They are ar-
ranged here in the general sequence of their use in the field.
Quick Reference Guide For Initial Site Activities B-2
Quick Reference Guide For Water Chemistry And Microbiology B-4
Quick Reference Guide For Physical Habitat Characterization B-7
Quick Reference Guide For Periphyton B-13
Quick Reference Guide For Sediment Metabolism B-15
Quick Reference Guide For Benthic Macroinvertebrates B-16
Quick Reference Guide For Aquatic Vertebrates B-20
Quick Reference Guide For Fish Tissue Contaminants B-23
-------
1.
2.
3.
4.
5.
6.
7.
8.
Quick Reference Guide for Initial Site Activities
Find the river location in the field corresponding to the "X" on 7.5" topo map (X-site). Crews should use
all available means to insure that they are at the correct site, as marked on the map, including: 1:24,000
USGS map orienteering, topographic landmarks, county road maps, local contacts, boat launches, and
global positioning system (GPS) confirmation of site latitude and longitude.
Classify the site. AT THE X-SITE. as:
NON-TARGET
TARGET
INACCESSIBLE
No Stream Channel
Impounded River
Marsh/Wetland
Regular Wadeable Stream
Regular - Partial Boatable and Wadeable Combination
Regular Boatable
Intermittent Stream
Dry Channel
Altered Channel (channel different form map representation)
Physical Barriers (Physically unable to reach the X-site)
No Permission
Record class on Site Verification form, do not sample Non-target or inaccessible sites. Take samples from
Target sites as discussed in field operations and methods manual.
At the launch site, unload the rafts and all equipment, supplies, and sample containers. Shuttle the
vehicles.
Using a laser rangefinder, measure the river width in several places, specifically the X-site and the two
boat launches. Record the width on the site verification form. Lay out a sample reach with a length of 100
times the river width by rolling a map wheel on the topographic map and marking the reach boundaries.
Do a reconnaissance of the sample reach while shuttling vehicles, obtaining widths, and evaluating
launch sites. Extensive shallows, large log jams, absence of launch sites or vehicle access, and hazardous
Whitewater may all preclude rafting.
Determine the float distance, if any, from the put-in to the first transect (Transect "A"), and from the last
transect (Transect "K") to the take-out.
Using a laser rangefinder at the most upriver transect (Transect "A"), measure 10 channel widths
downriver to the next transect (JTransect "B"). This distance is a profile.
Sample odd numbered site ID'S along the left shore (facing downriver); sample even numbered sites
along the right shore.
NOTE: There are some conditions that may require adjusting the reach about the X-site (i.e., the X-site
no longer is located at the midpoint of the reach) to accommodate river access or to avoid river
hazards or obstacles. If the beginning or end of the reach cannot be sampled due to obstacles or hazards,
make up for the loss of reach length by moving ("sliding") the other end of the reach an equivalent
distance away from the X site. Similarly, access points may necessitate sliding the reach. Do not "slide"
the reach so that the X-site falls outside of the reach boundaries. Also, do not "slide" a reach to avoid man-
made obstacles such as bridges, rip rap, or channelization.
-------
Quick Reference Guide for Water Chemistry and Microbiology
/. Equipment to Carry in Field for Water Chemistry and Microbiology
Rinse/Test bottles of QCCS in self-sealing plastic bag
D.O./Temperature/Conductivity Meter
FieldForms
One 500-mL plastic beaker with handle, in clean self-sealing plastic bag
One cubitainer in clean self-sealing plastic bag (barcode label attached)
Two 60-mL syringes in a plastic container (each one with a bar code label attached)
One 200-mL sterile square glass microbial bottle (barcode label attached)
Two syringe valves in the plastic container
Plastic cooler and several bags of ice
Opaque garbage bag
Electricians-tape
II. Extra Equipment to Carry in Vehicle
Back-up labels, forms, cubitainers, syringes, syringe valves, and microbial bottles
III. Daily Activities after Sampling
1. Check that cubitainer lid is on tight, has a flush seal, and is taped. Also tape the microbial cap.
2. Prepare the sample for shipping (label and seal cooler, replace ice as close as possible to shipping time)
OR direct delivery to the laboratory.
3. Call Overnight shipping company to arrange pick-up of cooler.
4. Rm'se the sampling beaker with deionized water three times.
5. Make sure field meters are clean and are stored with moist electrodes.
6. Label the next days sample containers (cubitainer, syringes, and microbial bottle), pack cubitainer
and sample beakers in clean self-sealing plastic bag, and pack two syringes, syringe valves, and a
microbial bottle in a plastic container with a snap-on lid.
(continued)
-------
Quick Reference Guide for Water Chemistry and Microbiology (continued)
Summary of Site Procedure for Water Chemistry and Microbiology
/. Collect Water Sample
A. Make sure cubitainers and syringes are labeled and have the same barcode ID.
B. Make sure the microbial bottle is labeled with a barcode ID.
C. Cubitainer, syringe, and microbial samples are taken only from the middle of the flowing river at
the last sample transect (Transect" K ").
D. Rinse the 500-mL sample beaker three times with river water from the mid-channel.
E. Rinse Cubitainer three times with 25-50 mL of river water, using the sample beaker. Rinse cubitainer
lid with river water.
F. Fill cubitainer with river water using the 500 mL sample beaker. Expel any trapped air and cap the
cubitainer. Make sure that the lid is seated correctly and that the seal is tight.
DO NOT EXPAND CUBITAINER BY BLOWING IN IT.
G. Rinse each of the two, 60-mL syringes three times with 10-20 mL of river water.
H. Fill each of the syringes with river water from mid-river by slowly pulling out the plunger. If any air
gets into the syringe, discard the sample and draw another.
L Invert the syringe (tip up) and cap the syringe with a syringe valve. Open the valve, tap the syringe
to move any air bubbles to the tip, and expel any air and a few mL of water. Make sure there is 50-60
mL of river sample in the syringe. Close the valve and place the syringes in their transport container.
J. Keep the microbial bottle closed until filled. Do not contaminate inner surface of cap or bottle. Fill
the bottle without rinsing.
K. Take sample from upriver side of boat by holding bottle near base and plunge neck downward below
water's surface. Turn bottle until neck points slightly upward and mouth is directed toward the
current.
L. After sample is collected, leave ample air in the microbial bottle (~ 2.5 cm) and tape the cap tight.
M. Place the cubitainer, syringes, and microbial bottle on ice in a cooler to keep cool (keep dark as well)
until shipment.
//. InSituMeasurements
A. Conductivity
1. Turn on and check the zero and red line (if applicable) of the conductivity meter.
2. Measure and record the conductivity of the QCC solution. Rinse the probe in the "Rinse" bottle
of QCC solution before immersing in the "Test" bottle of QCC solution.
3. Measure and record river conductivity in mid-river AT EACH TRANSECT.
B. Dissolved Oxygen/Temperature
1. Calibrate the DO meter following meter instructions.
2. Measure the DO in mid-river at the middle of the flowing river of the last sample transect
(Transect "K"). If water velocity is slow, jiggle the DO probe as you take the reading. Measure
temperature mid-river ATEACHTRANSECT.
-------
Quick Reference Guide for Physical Habitat Characterization
Field Summary: P-hab Layout And Workflow
1. Habitat Sampling Layout:
A. Thalweg Profile: At 10 equally spaced intervals between each of 11 channel cross-sections (100
along entire reach):
* Classify habitat type, record presence of backwater and off-channel habitats. (10 between cross-
sections, 100 total)
* Determine dominant substrate visually or using sounding rod. (10 between cross-sections, 100
total)
At 20 equally spaced intervals between each of 11 channel cross-sections (200 along entire
reach):
* Tally mid-channel snags (20 between cross-sections, 200 total).
* Measure thalweg (maximum) depth using Sonar or rod (20 between cross-sections, 200 total)
B. Littoral/Riparian Cross-Sections: @ 11 stops ("transects") at equal intervals along reach length
2. Work Flow: In a single mid-river float down a 100 channel-width reach
v At the upriver start point (Transect "A") and along the designated shoreline: Move boat in a "loop"
within a 10 x 20 m littoral plot, measuring 5 littoral depths and probing substrate. Also estimate
dominant and subdominant littoral substrate within the "loop." After the "loop," estimate areal fish
cover within and tally LWD within or partially within the 10 x 20 m plot. Record densiometer
measurements at the bank (up, down, left, right), and choose bank angle class, and estimate bankfull
• height, width and channel incision (for BOTH banks). Estimate and record distance to riparian
vegetation on the chosen bank. Estimate visually riparian vegetation cover for the 10 x 20 m plot on
BOTH sides of channel (plot starts at bankfull, continues back 10m from bankfull). For the largest
riparian tree, estimate Dbh, height, species, distance from river edge. Visually tally human
disturbances in the same plot as riparian vegetation. No bearing or slope at first cross section.
• Proceed downriver between Transects "A" and "B", making 20 thalweg depth measurements and
substrate snag probes; also classify habitat types. Estimate thalweg distance intervals by tracking boat
lengths or channel-widths. One person measures thalweg depths and the other records those
measurements. At the 20th thalweg measurement location (close to Transect "B"), backsite a compass
bearing in mid-channel, then distance and % slope back to your visual "mark" on the bank at the
previous transect ("A").
• When you complete 20 thalweg intervals and reach one of 11 cross sections, stop at the chosen shore
and take out a new Channel/Riparian Transect Form for Transect "B". Repeat all the Channel/Riparian
measurements at this new location.
• Repeat the cycle of thalweg and cross section measurements until you reach transect 11 ("K") at the
downriver end.
(continued)
-------
Quick Reference Guide for Physical Habitat Characterization
Field Summary: Components of P-Hab Protocol
Thalweg Depth Profile, Mid-Channel Snags, Hab. Type, Off-channel, Substrate:
• At 20 approximately equal spaced intervals between each of 11 channel cross-sections (200 along
entire reach) while floating mid-channel:
- Measure max. depth ("Thalweg") at each increment
- Tally mid-channel snags
• At 10 approximately equal spaced intervals between each of 11 channel cross-sections (100 along
entire reach) while floating mid-channel:
- Classify habitat type and off-channel habitats
- Determine dominant substrate
Channel and Riparian Cross-Sections:
• Measurements: Wetted width, mid-channel bar width, gradient (clinometer or Abney level), sinuosity
(compass backsite), riparian canopy cover (densiometer).
• Visual Estimates: Bankfull width, bankfull height, incision height, bank angle, shoreline substrate,
large woody debris, areal cover class and type of riparian vegetation in Canopy, Mid-Layer and
Ground Cover; areal cover class of fish cover features, aquatic macrophytes, and filamentous algae;
presence and proximity of human disturbances.
(continued)
-------
Quick Reference Guide for Physical Habitat Characterization (continued)
Field Summary: Rip. Veg., Human Disturb., In-Channel Cover:
Observations upriver 10 meters and downriver 10 meters from each of the 11 cross-section transects.
For riparian vegetation and human disturbances, include the visible area from the river back a distance of
10m (30 ft) shoreward from both the left and right banks. If the wetted channel is split by a mid-channel
bar, the bank and riparian measurements shall be for each side of the channel, not the bar.
Three vegetation layers: "
CANOPY LAYER (>5 mhigh)
UNDERSTORY (0.5 to 5 m high)
GROUND COVER layer (<0.5 m high)
Canopy and UnderstOry Vegetation Types:
(Deciduous, Coniferous, Broadleaf Evergreen, Mixed, or None) in each of the two taller layers
(Canopy and Understory). "Mixed" if more than 10% of the areal coverage made up of the alternate
type.
Areal Cover Classes for Vegetation and In-Channel Cover:
0: (absent — zero cover)
1: (sparse - cover <10%)
2: (moderate - cover 10-40%)
3: (heavy - cover 40-75%)
4: (very heavy — cover >75%).
Tallying Human Disturbances:
B: PRESENT within the defined 20 m river segment and located in the river or on the wetted or
bankfull river
C: CLOSE - Present within the 10 x 20 m riparian plot area, but above bankfull level
P: PRESENT, but observed outside the riparian plot area
0: NOT PRESENT within or adjacent to the 20 m river segment or riparian plot
-------
Quick Reference Guide for Physical Habitat Characterization (continued)
Field Summaries: Substrate And Woody Debris Size Classes
Observe bottom substrates within a 10m swath along the 20m of channel margin that is centered on each
transect location. Determine and record the dominant and subdominant substrate size class at 5 system-
atically spaced locations estimated by eye within this 10m x 20m plot and 1m back from the waterline.
Substrate Size Classes:
RS Bedrock (Smooth)
RR Bedrock (Rough)
HP Hardpan
BL Boulders
CB Cobbles
GC Gravel(Coarse)
GF Gravel (Fine)
SA Sand
FN Fines
WD Wood
OT Other
>4000 mm
>4000 mm
>4000 mm
>250 to 4000 mm
64 to 250 mm
16 to 64 mm
2 to 16 mm
0.06 to 2 mm
<0.06 mm
Regardless of Size
Regardless of Size
smooth surface rock or hardpan (bigger than a car)
Rough surface rock (bigger than a car)
(consists of firm, consolidated fines)
(basketball to car size)
(tennis ball to basketball size)
(marble to tennis ball size)
(ladybug to marble size)
(smaller than ladybug size, but visible as particles -
gritty between fingers).
Silt-Clay-Muck (not gritty between fingers)
Wood or other organic material
Metal, Tires, Car bodies, asphalt, concrete, etc.
(Describe in comments if you enter "OT").
Large Woody Debris Size Classes:
LWD Definition:
Diameter (small end) > 30 cm (>1 ft.)
Length > 5 m (> 15 ft) — count only part with diam > 30 cm.
Two Tallys:
(1) LWD at least partially in the baseflow channel (wetted).
(2) LWD presently dry but contained within the bankfull (active) channel, and LWD
spanning above the active channel.
Size Categories for Tally (12 potential combinations):
Diameter (large end):
Length:
0.3 to <0.6 m
0.6 to <0.8 m
0.8 to <1.Om
>1.0m
(1 to 2 ft.)
(2 to 2.6 ft)
(2.6 to 3.3ft)
(> 3.3ft)
5-<15m
15 - <30 m
>30m
(16-49 ft)
(49 - 98 ft)
(>98ft)
(continued)
-------
Quick Reference Guide for Physical Habitat Characterization (continued)
Field Summary: Habitat Classification At Channel Unit Scale
Class (Code)
Pools (PO):
Plunge Pool
Trench Pool
Lateral Scour Pool
Backwater Pool
Dam Pool
Glide (GL)
Riffle (RI)
Rapid (RA)
Cascade (CA)
Falls (FA)
Dry Channel (DR)
Off-Channel Areas
Channel Unit Habitat Classes"
Description
Still water, low velocity, smooth, glassy surface, usually deep compared to other parts
of the channel:
Pool at base of plunging cascade or falls.
Pool-like trench in the center of the stream
Pool scoured along a bank.
Pool separated from main flow off the side of the channel.
Pool formed by impoundment above dam or constriction.
Water moving slowly, with a smooth, unbroken surface. Low turbulence.
Water moving, with small ripples, waves and eddies - waves not breaking, surface
tension not broken. Sound: "babbling", "gurgling".
Water movement rapid and turbulent, surface with intermittent Whitewater with
breaking waves. Sound: continuous rushing, but not as loud as cascade.
Water movement rapid and very turbulent over steep channel bottom. Most of the
water surface is broken in short, irregular plunges, mostly Whitewater. Sound: roaring.
Free falling water over a vertical or near vertical drop into plunge, water turbulent and
white over high falls. Sound: from splash to roar:
No water in the channel
Side-channels, sloughs, backwaters, and alcoves that are separated from the main
channel.
"Note that in order for a channel habitat unit to be distinguished, it must be at least as wide or long as the
channel is wide.
Field Summary: P-hab Problem Areas
Mid-channel Bars: dry at baseflow, inundated at bankfull flow.
Measure wetted width across and over mid-channel bars, but record bar width in the column
provided on the Channel/Riparian Transect Form.
Islands: as high as the surrounding flood plain; dry even at bankfull flow.
Measure only the width of the main channel between island and shore
Both bars and islands cause the river to split into side channels. When a bar or island is encountered along the
thalweg profile, choose to navigate and survey the channel that carries the most flow.
Side channels (off-channeD:
When present, check the "Off-channel" column on the Thalweg Profile Form. Begin checking at
the point of divergence continuing until convergence. In the case of a slough or alcove, "off-
channel" checkmarks should continue from the point of divergence downriver to where it is no
longer evident.
Dry and Intermittent rivers:
Record zeros for depth and wetted width in places where no water is in the channel. Record habitat
type as dry channel (DR).
-------
Quick Reference Guide For Periphyton
Field Equipment
1. Large funnel (15-20 cm diameter).
2. Scrape area delimiter (3.8 cm diameter pipe, 3 cm tall).
3. Stiff-bristle toothbrush with handle bent at 90° angle.
4. Wash bottle.
5. Collection bottle to catch removed periphyton.
6. 60 mL syringes with 3/8" hole bored into the end.
7. 50 mL centrifuge tubes or similar sample vials.
8. Formalin.
9. Glass-fiber filters (0.45m average pore size) for chlorophyll a and biomass (AFDM).
10. Forceps for filter handling.
11. Millipore®-type filtration apparatus with plastic or stainless steel filter base, and Nalgene® funnel and
suction flask.
12. Nalgene® hand-operated vacuum pump (need one additional pump as a backup).
13. Aluminum foil.
14. Ice chest.
Field Protocols
1. Periphyton samples are collected from the designated shoreline at each transect location.
2. Collect a sample of substrate (rock or wood) that is small enough (< 15 cm diameter) and can be easily
removed from the river. Place the substrate in a plastic funnel which drains into a 500-mL plastic bottle
with volume graduations marked on it.
3. Use the area delimiter to define a 12-cm2 area on the upper surface of the substrate. Dislodge attached
periphyton from the substrate within the delimiter into the funnel by brushing with a stiff-bristled
toothbrush for 30 seconds. Take care to ensure that the upper surface of the substrate is the surface that is
being scrubbed, and that the entire surface within the delimiter is scrubbed.
4. Fill a wash bottle with river water. Using a minimal volume of water from this bottle, wash the dislodged
periphyton from the funnel into the 500-mL bottle.
5. If no coarse sediment (cobbles or larger) are present, collect soft sediments by vacuuming the upper 1 cm
of sediments confined within the 12-cm2 sampling ring into a 60-mL syringe.
6 Place the sample collected at each sampling site into the single 560-mL bottle to produce the composite
index sample.
7. After samples have been collected from all 11 transects, thoroughly mix the 500-mL bottle regardless of
substrate type.
Record total volume of composited sample before proceeding to the next step!
Four subsamples will be taken from each composite sample. These are:
a. Identification/Enumeration
1) Withdraw 50 mL of mixed sample and place in a labeled sample vial (50-mL centrifuge tubes
work well). Cover label with clear tape.
2) Preserve sample with 2 mL of 10% formalin. Gloves should be worn.
3) Tightly cap tube and tape with electrical tape.
b. Chlorophyll a
1) Withdraw 25 mL of mixed sample and filter onto a glass-fiber filter (0.45 um pore size) using a
hand-operated vacuum pump. (Note: for soft-sediment samples, allow grit to settle before
withdrawing sample).
2) Fold filter so that the sample on the filter surface is folded together, wrap in aluminum foil, and
affix the tracking label to the outside, and seal with clear tape.
3) Freeze filter as soon as possible by placing it in a freezer.
4) Store frozen for laboratory analysis.
(continued)
8.
9.
-------
Quick Reference Guide For Periphyton (continued)
c. Ash Free Dry Mass (AFDM)
1) Withdraw 25 mL of mixed sample and filter onto a glass-fiber filter (0.45 um pore size) using a
hand-operated vacuum pump. (Note: for soft-sediment samples, allow grit to settle before
withdrawing sample).
2) Fold filter so that the sample on the filter surface is folded together, wrap in aluminum foil, and
affix the tracking label to the outside, and seal with clear tape.
3) Freeze filter as soon as possible by placing it in a freezer.
4) Store frozen for laboratory for analysis.
d. Alkaline/Acid Phosphatase
1) Withdraw 50 mL of mixed sample and place in a labeled sample vial (50-mL centrifuge tubes
work well). Cover label with clear tape. - -
2) Tightly cap tube and tape with electrical tape.
3) Freeze sample as soon as possible by placing it on dry ice.
4) Store frozen for laboratory analysis.
-------
Quick Reference Guide for Sediment Metabolism
Field Equipment
1. Ice chest for floating centrifuge tubes during incubation
2. 1000 mL Nalgene© beaker for holding centrifuge tubes during incubation.
3. Small scoop sampler for sediments.
4. 50-mL, screw-top, centrifuge tubes.
5. Digital dissolved oxygen meter (e.g. YSI95).
6. Spare batteries for D.O. meter.
7. Permanent markers for labeling tubes.
8. Sample labels and field data sheets.
9. Ice chest with ice for sample freezing.
Field Protocols
Dissolved Oxygen Meter Calibration (for YSI model 95)
1. Calibrate meter using the water-saturated atmosphere chamber described in the meter's operations
manual. Allow at least 15 minutes for the probe to equilibrate before attempting to calibrate.
Sediment Collection and Experimental Set-up
1. Collect and combine fine-grained, surface sediments (top 2 cm) from all depositional areas at each
transect (Transects A-K) along the designated shoreline of the river reach.
2. Fill ice chest 2/3 full with river water and record temperature and dissolved oxygen (D.O.).
3. Thoroughly mix composite sediment sample.
4. Place 10 mL of sediment in each -of 5 labeled, 50 mL screw-top centrifuge -tubes.
5. Fill each tube to the top (no head space) with stream water from the ice chest and seal.
6. Fill one additional tube with stream water only to serve as a. blank.
7. Incubate tubes in closed ice chest for 2 hours.
8. Measure D.O. in each tube, including the blank.
9. Decant overlying water and save sediment.
10. Tightly seal tubes and freeze as soon as possible.
11. Store frozen for laboratory analysis.
-------
Quick Reference Guide For Benthic Macroinvertebrates
Table I. Base Protocols for Collecting Macroinvertebrates
1. Set drift net assembly(s) near the put-in or take-out location.
2. Shore kick net samples are collected at each of the transect locations along the designated shoreline.
Drift nets collect samples during the sampling day while the crew floats the river.
3. If riffle or run, use the kick net protocol in Table II. If pool, use the kick net protocol in Table HI or hand
pick for 60 seconds if kick net cannot be used.
4. Go to next downriver transect and repeat. Combine all riffle and pool samples into one bucket. Check net
after each sample for clinging organisms and transfer to bucket.
5. After a sample is collected from each of the transects and all kick net samples are combined into one
bucket^ obtain a composite sample as described in Table V.
6. Drift net(s) procedures are described in Table IV. Processing is described in Table V.
7. Preserve and label each sample as described in Table VI.
/
Table II. Procedures for Riffles and Glides using Kick Net Sampler
1. Attach four foot pole to the sampler.
2. Position sampler quickly and securely on river bottom with net opening upriver.
3. Hold the sampler in position on the substrate while checking for snails and clams in an area of about 0.5
m2 in front of the net; kick the substrate vigorously for about 20 seconds in front of the net.
4. Inspect and rub off with the hands any organisms clinging to the rocks, especially those covered with
algae or other debris. /
5. Remove the net from the water with a quick upriver motion to wash the organisms to the bottom of net.
6. Rinse' net contents into a bucket containing one or two gallons of water by inverting the net in the water.
7. Inspect the net for clinging organisms. With forceps remove any organisms found and place them into the
bucket.
8. Large objects (rocks, sticks, leaves, etc.) in the bucket should be carefully inspected for organisms before
discarding.
9. After all transects are sampled and all samples are combined in ONE bucket (riffle/glide + pool), obtain
a composite sample as described in Table V.
Table III. Procedures for Pools using the Modified Kick Net Sampler
1. Attach four-foot pole to the sampler.
2. Inspect about ° square meter of bottom for any heavy organisms, such as mussels and snails, which have
to be hand picked and placed in the net.
3. While disturbing about 0.5 m2 of substrate by kicking, collect a 20-second sample by dragging the net
repeatedly through the area being disturbed. Keep moving the net all the time so that the organisms
trapped in the net will not escape.
4. After 20 seconds remove the net from the water with a quick upriver motion to wash the organisms to the
bottom of the net.
5. Rinse net contents into a small bucket of water (about one or two gallons) by inverting the net in the
water.
6. Inspect the net for clinging organisms. With forceps remove any organisms found and place them in the
bucket.
7. Large objects in the bucket should be carefully inspected for organisms which are washed into the bucket
before discarding.
8. After all transects are sampled and all samples are combined in ONE bucket (pool + riffle/glide), obtain
a composite sample as described in Table V.
Table IV. Collection Procedures for Drift Nets
1.
Do not use drift nets for large rivers with currents less than 0.05 m/s.
(continued)
-------
Quick Reference Guide For Benthic Macroinvertebrates (continued)
2. Install the net at the downriver end of the reach (Transect K). The take-out location is 1st choice,
otherwise the put-in location - whichever is closer to the reach.
3. Set the nets in the main flow of the river (avoid backwaters, eddies, river margins) at depths of about 25
cm from the bottom substrate and 10 cm below the water's surface.
4. Anchor the net assembly using anchors and cables. Record START TIME.
5. Measure the current velocity at the entrance of the net, using a neutrally buoyant object as follows:
a. Measure out a straight segment of the river reach just upstream of the drift net location in which
an object can float relatively freely and passes through within about 10 to 30 seconds.
b. Select an object that is neutrally buoyant, like a small rubber ball or an orange; it must float, but
very low in the water. The object should be small enough that it does not "run aground" or drag
bottom.
c. Time the passage of the object through the defined river segment 3 times. Record the length of
the segment and each transit time in the Comments section of the Sample Collection Form.
6. After floating the river, retrieve the net assembly from the water, taking care not to disturb the bottom
upriver of the net. Record the END TIME.
7. Determine the current velocity again as described above, calculate the average from the 6 measurements,
and record on the form.
8. Concentrate the material in each net in one corner by swishing up and down in the river. Wash the
material into a bucket half filled with water (NOT the shore sample bucket). Remove as much as possible
from the nets.
9. The contents from both nets are combined into a single bucket. After this, pour the sample over a sieving
bucket (same bucket used in the kick net samples).
10. Large objects in the bucket should be carefully inspected for organisms which are washed into the bucket
before discarding.
11. After both nets are combined into one bucket, obtain a composite sample as described in Table V.
Table V. Procedures for Obtaining the Composite Sample
1. Pour the contents of the composite bucket through a U.S. Standard 30 sieve. Examine the bucket while
rinsing it well to be sure all organisms are washed from the bucket onto the sieve.
2. Wash contents of the sieve to one side by gently agitating in water and wash into jar using as little water
from the squirt bottle as possible. Carefully examine the sieve for any remaining organisms and place
them in the appropriate jar labeled as either "shore" or "drift" sample.
Table VI. Sample Preserving and Labeling
1. Fill in special pre-numbered barcoded label and place on jar. All additional jars used for a sample must
be labeled with same number. Enter this number which will be used for tracking purposes in the
computer.
2. Preserve samples in ethanol as follows:
a. If jar is more than 1/4 full of water, pour off enough to bring it to less than 1/4 full using proper
sieve to retain organisms.
b. Fill jar nearly full with 95% ethanol so that the concentration of ethanol is 70%. If there is a
small amount of water in the sample, it may not be necessary to fill the jar entirely full to reach
a 70% concentration.
c. Transfer any organisms on the sieve back into the jar with forceps.
3. Check to be sure that the pre-numbered stick-on barcoded label is the on jar. Cover the entire label with
clear, waterproof tape.
4. Seal the caps with electrical tape.
5. Place samples in cooler or other secure container for transport.
6. Secure all equipment in the vehicle.
-------
Quick Reference Guide for Aquatic Vertebrates
Field Protocols For Fish Collection
1. Site Selection
a. Determine river bank to be sampled. Stay along this shore the entire day, unless river aspect is
unchanging and the selected side is not representative of both.
b. Float downriver along the designated shoreline, stopping at each transect (A to K).
c. In case of emergency, determine location of means of easy egress from river.
2. Electrofishing
a. Check all electrical connections and potential conductors. Place cathodes and anodes in the
water. Fill livewell with river water.
b. Start generator, switch to pulsed DC, a frequency of SOpps, low range and 40%. These are the
initial settings. Set timer arid depress pedal switch to begin fishing.
c. With switch depressed and floating downriver near shore, maneuver the raft or anode to cover a
swath 3-4 meters wide, at an oar's length from shore, near cover, and at depths less than 3 meters
wherever possible.
d. Deposit fish in the livewell as soon as possible; do not hold them in the electrical field.
e. Continue fishing until the next transect.
f. Process fish when stopped at each transect. Record total time spent collecting and shocking time
on data sheets.
g. Identify and release any threatened and endangered species.
h. Identify and measure (TL) sport fish and very large specimens, record external anomalies, and
release unharmed.
i. Identify other specimens. Determine number of individuals in species, measure largest and
smallest individuals, and voucher as described in Voucher Protocol.
j. Large, questionable species should be placed on ice and then frozen.
k. Retain a subsample of target species for Fish Tissue Contaminants analysis.
(continued)
-------
Quick Reference Guide for Aquatic Vertebrates (continued)
Categories
Absent
Blisters
Blackening*
Extensive Black spot disease
Cysts
Copepod
Deformities
Eroded fins
Eroded gills
Fungus
Fin anomalies
Grubs
Hemorrhaging
Ich
Lesions
Lice
Mucus
None
Other
Scale anomalies
Shortened operculum
Tumors
Leeches
Exophthalmia
Anomaly Categories and Codes
Code Definition
AB
BL
BK
BS
CY
CO
DE
EF
EG
FU
FA
WG
HM
1C
LE
LI
MU
NO
or
SA
SO
TU
WL
EX
Absent eye, fin, tail.
In mouth, just under skin.
Tail or whole body with darkened pigmentation.
Small black cysts (dots) all over the fins and body.
Fluid-filled swellings; maybe small dots or large.
A parasitic infection characterized by a worm like
copepod embedded in the flesh of the fish; body
extends out and leaves a sore/discoloration at base,
may be in mouth gills, fins, or anywhere on body.
Skeletal anomalies of the head, spine, and body shape;
amphibians may have extra tails, limbs, toes.
Appear as reductions or substantial fraying of fin
surface area.
Gill filaments eroded from tip.
May appear as filamentous or "fuzzy" growth on the
fins, eyes, or body.
Abnormal thickenings or irregularities of rays
White or yellow worms embedded in muscle or fins.
Red spots on mouth, body, fins, fin bases, eyes, and
gills.
White spots on the fins, skin or gills.
Open sores or exposed tissue; raised, granular or warty
outgrowths.
Scale-like, mobile arthropod.
Thick and excessive on skin or gill, as long cast from
vent.
No anomalies present.
Anomalies or parasites not specified (Please comment).
Missing patches, abnormal thickings, granular skin
Leaves a portion of the gill chamber uncovered
Areas of irregular cell growth which are firm and
cannot be easily broken open when pinched. (Masses
caused by parasites can usually be opened easily.)
Annelid worms which have anterior and posterior
suckers. They may attach anywhere on the body.
Bulging of the eye.
(continued)
-------
Quick Reference Guide for Aquatic Vertebrates (continued)
Guidelines and Procedures for Preparing Fish Voucher Specimens
Category 1. Large easily identified species OR adults may be difficult to identify OR the species is
uncommon in that region. Preserve 1-2 small (<150 mm total length) adult individuals per site
plus 2-5 juveniles. If only large adults are collected, reserve smallest individual until voucher
procedure is complete and preserve ONLY if space is available. Photograph if considered too
large for the jar.
Category 2. Small to moderate-sized fish OR difficult to identify species. Preserve up to 20 adults and
juveniles. If less than 20 individuals are collected, voucher all of them.
Category 3. Species of "special concern." These are state or federally listed species. Photograph and release.
If specimens have died, include in voucher collection, note on data sheet and notify appropriate
state official as soon as possible.
a. After all individuals of a species have been processed, place the voucher sub sample in
a bucket with carbon dioxide tablets and a small amount of water. Individuals > 160
mm should be slit on the lower abdomen of the RIGHT side.
When specimens are dead, transfer to a small nylon bag containing a waterproof label
with tag#.Place in "Voucher" jar in 10% formalin. BE SURE THAT JAR IS LABELED
INSIDE AND OUT WITH A VOUCHER LABEL (site ID, barcode, and date).
DO NOT over pack the sample jars with specimens OR use less formalin than is needed.
If a fish will not fit in a jar, freeze the specimen.
Continue until all species are processed. Seal voucher jar with electrical or clear tape.
Check that the jar is correctly labeled. Enter BARCODE'ID in appropriate place on
field data sheet.
Transport to storage depot at end of week. Store in a cool, dark, ventilated space.
b.
c.
d.
e.
-------
Quick Reference Guide For Fish Tissue Contaminants
Selecting And Processing Fish Tissue Specimens
NOTE: If neither a primary nor secondary species sample is available, use your best judgement in sending
some type of composite fish tissue sample.
Primary Sample (P)
After all voucher specimens have been prepared, choose a cottid, cyprinid, or salmonid that has enough
similarly sized individuals to weigh to 400 g.
Secondary Sample ("SI
After all voucher specimens have been prepared, select a large piscivore or omnivore species that has at least
5 individuals 120 mm. Include similar sized individuals if available.
1. Place the fish into abucket with two carbon dioxide tablets (e.g., "Alka Seltzer®") and a small volume
of water. After they have been anaesthetized, use clean hands to transfer them to aluminum foil.
2. Prepare a clean work surface to prepare the primary composite sample. Keep hands, work surfaces, and
wrapping materials clean and free of potential contaminants (mud, fuel, formalin, sun screen, insect
repellant, etc.)
3-P. For primary samples, record the common name (from a standardized list) of the species, its species code
(if required), and the number of individuals in the sample in the appropriate fields on line "PI" of the
Sample Collection Form (Figure 11-1).
3-S. Measure the total length (TL) of each secondary individual. Record the common name (from a
standardized list) of the secondary target species, its species ctide (if required), and the total length for
each individual on lines SI through S5 in the secondary sample section of the Sample Collection
Form.
4. If the individuals included in composite samples were collected from throughout the sampling reach,
pjace an "X" in the "Yes" box in the sample section of the Sample Collection Form. If the individuals
were only collected from a limited segment of the sampling reach, place an "X" in the "No" box and
explain in the "Explain" field on the form.
5-P. Wrap all primary fish together in a single piece of aluminum foil, making sure the dull side of the
aluminum foil is in contact with the fish. Place the sample in a self-sealing plastic bag.
5-S. Wrap each fish of the secondary sample separately in aluminum foil, with the dull side of the foil in
contact with the fish. Place all the wrapped individuals into a single self-sealing plastic bag.
6 Expel excess air and seal the bag. Wrap clear tape around the bag to seal and make a surface for each
sample label. . .
7-P. Prepare two Fish Tissue sample labels (each having the same sample ID number [Figure 11-2]) by
filling in the stream ID and the date of collection. Circle' PRIMARY'' on each label. Record the sample
ID number (barcode) in the primary sample section of the Sample Collection Form.
7-S. Prepare two Fish Tissue sample labels (each having the same sample ID number [Figure 11-2]) by
filling in the stream ID and the date of collection. Circle "SECONDARY" on each label. Record the
sample ID number (barcode) in the secondary sample section of the Sample Collection Form.
8. Attach the appropriate label to the tape surface of the bag. Cover the label with a strip of clear tape.-
Place the labeled bag into a second self-sealing plastic bag. Seal the bag and attach the second label to
the outside of the appropriate bag. Cover the, label with a strip of clear tape.,
J. Place the double-bagged sample into a cooler containing bags of ice until shipment. Keep the sample
frozen until shipment.
-------
Appendix C
Species Codes for Aquatic Vertebrates
The following table contains the unique
8-character species code, the scientific name,
and the common name assigned to each
aquatic vertebrate species expected to be col-
lected by EMAP sampling protocols in the
Mid-Atlantic and Western regions. Gener-
ally, the species code is composed of the first
four letters of the genus plus the first four let-
ters of the species name. Modifications to this
coding scheme were made in cases where two
species could be assigned the same code.
Species entries are arranged first by family
(alphabetically), then by the assigned species
code.
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Code
Aquatic Vertebrates of the Western United States.
Latin name Common name (vouchering category)
LAMPAY Lampetra ayresi
LAMPLE Lampetra lethophaga
LAMPRI Lampetra richardsoni
LAMPTR Lampetra tridentata
LAMPSI Lampetra similis
LAMPZZ
COITAL Cottus aleuticus
COTTAS Cottus asper
COTTBA Cottus bairdi
COTTBE Cottus beldingi
COTTCF Cottus confusus
COTTGU Cottus gulosus
COITKL Cottus klamathensis
COTTMA Cottus marginatus
COTTPE Cottus perplexus
COTTPI Cottus pitensis
COTTPR Cottus princeps
COTTRH Cottus rhotheus
COTTTE Cottus tenuis
LEPTAR Leptocottus armatus
COTTZZ
ACIPME Acipenser medirostris
ACBPTR Acipenser transmontanus
ALOSSA Alosa sapidissima
CLUPPA Clupea pallasi
ONCOGO Oncorhynchus gorbuscha
ONCOKE Oncorhynchus keta
ONCOKI Oncorhynchus kisutch
ONCONE Oncorhynchus nerka
river lamprey 2
pit-klamath brook lamprey 2
western brook lamprey 2
pacific lamprey 2; goose lake lamprey 3
klamath river lamprey 2
unknown lamprey
coastrange sculpin 2
prickly sculpin 2
mottled sculpin 2; malheur motted sculpin 3
paiute sculpin 2
shorthead sculpin 2
riffle sculpin 2
marbled sculpin 2
margined sculpin 3
reticulate,sculpin 2
pit sculpin 3
klamath lake sculpin 2
torrent sculpin 2
slender sculpin 3
pacific staghorn sculpin 2
unknown Cottid
green sturgeon 1
white sturgeon 1
American shad 1
pacific herring 2
pink salmon 3
chum salmon 3
coho salmon 3
sockeye salmon 3
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Code
Latin name
Common name (vouchering category)
ONCOTS Oncorhynchus tshawytscha
ONCOAG Oncorhynchus aguabonita
ONCOCL Oncorhynchus clarki
ONCOMY Oncorhynchus mykiss
PROSWI Prosopium williamsoni
SALMSA Salmo salar
SALMTR Salmo trutta
SALVCO Salvelinus confluentus
SALVFO Salvelinus fontinalis
SALVNA Salvelinus namaycush
SALM2Z
HYPOPR Hypomesus pretiosus
SPIRTH Spirinchus thaleichthys
THALPA Thaleichthys pacificus
ACROAL Acrocheilus alutaceus
CARAAU Carassius auratus
CTENID Ctenopharyngodon idella
CYPRCA Cyprinus carpio
GILAAL Gila alvordensis ,
GILABI Gila bicolor
GDLABO Gila boraxobius
GELACO Gila coerulea
LAVISY Lavinia symmetricus
MYLOCA Mylocheilus caurinus
NOTECR Notemigonus crysoleucas
PIMEPR Pimephales promelas
PTYCOR Ptychocheilus oregonensis
PTYCUM Ptychocheilus umpquae
RHINCA Rhinichthys cataractae
RHINEV Rhinichthys evermanni
chinook salmon 3
golden trout 1
cutthroat trout; Umpqua 3, Lahontan 3
rainbow trout 1
mountain whitefish 1
atlantic salmon 1
brown trout 1
• bull trout 3
brook trout 1
lake trout 1
unknown salmonid
surf smelt 2
longfin smelt 2
eulachon 2
chiselmouth 1
goldfish 1
grass carp 1
common carp 1
alvord chub 3
tui chub 1; Callow, Hutton, Goose Lake, Oregon
Lakes, Sheldon, Summer Basin and Warner 3
borax lake chub 3
blue chub 2
California roach 3
peamouth 1
golden shiner 2
fathead minnow 2
northern squawfish 1
umpqua squawfish 1
longnose dace 2
umpqua dace 2
IMF :|ig!|iagil»i!i!iii!liilHl|ii! JiiiluiiiiiilliiiiJiipllljijIlliilllljiEiii':
-------
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Latin name
Common name (vouchering category)
RHINFA Rhinichthys falcatus
RHINOS Rhinichthys osculus
RICHBA Richardsonius balteatus
RICHEG Richardsonius egregius
TINCTI Tinea tinea
OREGCR Oregonichthys crameri
OREGKA Oregonichthys kalawatseti
CYPRZZ
CATOCB Catostomus columbianus
CATOMA Catostomus macrocheilus
CATOOC Catostomus occidentalis
CATOPL catostomus platyrhynchus
CATORI Catostomus rimiculus
CATOSY Catostomus snyderi
CATOTA Catostomus tahoensis
CATOWA Catostomus warnerensis
CHASER Chasmistes brevirostris
DELTLU Deltistes luxatus
CATOZZ
MISGAN Misgurnus anguillicaudatus
AMEICA Ameiurus catus
AMEIME Ameiurasmelas
AMEINA Ameiurus natalis
AMEINE Ameiuras nebulosus
ICTAPU Ictalurus punctatus
NOTUGY Noturus gyrinus
PYLOOL Pylodictis olivaris
PERCTR Percopsis transmountana
LOTALO Lota lota
MICRPR Microgadus proximus
leopard dace 2
speckled dace 2; foskett speckled dace 3; millicoma
dace 3
redside shiner 2
lahontan redside 3
tench 1
Oregon chub 3
umpqua chub 3
unknown cyprinid
bridgelip sucker 1
largescale sucker 1
sacramento sucker 1; Goose Lake sucker 3
mountain sucker 2
klamath smallscale sucker 1; Jenny Creek sucker 3
klamath largescale sucker 3
tahoe sucker 3
warner sucker 3
shortnose sucker 3
lost river sucker 3
unknown catostomid
oriental weatherfish 2
white catfish 1
black bullhead 1
yellow bullhead 1
brown bullhead 1
channel catfish 1
tadpole madtom 2
flathead catfish 1
sand roller 2
burbot 1
pacific tomcod 2
-------
Code
Latin name
Common name (vouchering category)
FUNDDI
LUCAPA
GAMBAF
ATHEAF
GASTAC
MOROSA
ARCHIN
LEPOCY
LEPOGI
LEOPGU
LEPOMA
LEPOMI
MCRDO
MICRSA
POMOAN
POMONI
CENTRZZ
PERCFL
STEZVI
CYMAAG
PHOLOR
PLATST
PSEURE
ASCATR
RANAAU
RANABO
RANACA
RANACT
RANAPI
RANAPR
Fundulus diaphanus
Lucania parva
Gambusia affinis
Atherinops affinis
Gasterosteus aculeatus
Morone saxatilis
Archoplites interruptus
Lepomis cyanellus
Lepomis gibbosus
Lepomis gulosus
Lepomis macrochirus
Lepomis microlophus
Micropterus dolomieui
Micropteras salmoides
Pomoxis annularis
Pomoxis nigromaculatus
Percaflavescens
Stizostedion vitrqum
Cymatogaster aggregata
Pholis omata
Platichthys stellatus
Pseudacris regilla
Ascaphus traei
Rana aurora
Rana boylii
Rana cascadae
Rana catesbiana
Rana pipiens
Rana pretiosa
banded killifish 2
rainwater killifish 2
western mosquitofish 2
topsmelt 2
threespine stickleback 2
striped bass 1
sacramento perch
green sunfish 1
pumpkinseed 1
warmouth 1
bluegill 1
redear sunfish 1
smallmouth bass 1
largemouth bass 1
white crappie 1
black crappie 1
unknown centrarchid
yellow perch 1
walleye 1
shiner perch 1
saddleback gunnel 2
starry flounder 1
pacific tree frog
tailed frog
red-legged frog
foothill yellow-legged frog
cascade frog
bullfrog
leopard frog
spotted frog
'-4TJ,
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-------
Code
Latin name
Common name (vouchering category)
BUFOBO Bufo boreas
BUFOWO Bufo woodhousii
AMBYGR Ambystoma gracile
AMBYMA Ambystoma macrodactylum
AMBYIT Ambystoma tigrinum
DICACO Dicamptodon copei
DICATE Dicamptodon tenebrosus
RHYACA Rhyacotrition cascadae
RHYAKE Rhyacotriton kezeri
RHYAVA Rhyacotriton variegatus
TARIGR Taricha granulosa
AMPHZZ Rana sp. -
western toad
woodhouse's toad
northwestern salamander
longtoed salamander
tiger salamander
cope's giant salamander
pacific giant salamander
cascade torrent salamander
Columbia torrent salamander
southern torrent salamander
rough-skinned newt
unknown amphibian
^
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United States
Environmental Protection Agency/ORD
National Exposure Research Laboratory
Research Triangle Park, NC 27711
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detach or copy, and return to the address in the upper
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Penalty for Private Use
$300
EPA/620/R-00/007
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