BUREAU OF CLEAN WATER
Appendix C – Biological Field Methods
C2. Benthic Macroinvertebrates
DECEMBER 2013
BENTHIC MACROINVERTEBRATES
This chapter describes field and laboratory methodology for sampling and processing
benthic samples collected using the Department’s Instream Comprehensive Evaluation
(ICE, 2011) and habitat specific stream protocols for riffle/run freestone (2012),
limestone (2009), and multi-habitat pool/glide (2007) streams. The following definitions
are applied accordingly:
Freestone streams: Streams that normally have higher gradient so as to be dominated
by riffle/run habitat consisting of various particle-sized substrates of unconsolidated
(“free” stone) sand, gravel, and cobble. Further, ‘freestone’ commonly refers to
streams with lower alkalinity and habitat conditions that do not meet the “limestone
stream” definition. Note: Some freestone streams may be ‘limestone influenced’
where they exhibit high alkalinities but still display unconsolidated freestone
substrates not characteristic of true limestone streams.
Limestone streams: Cold water streams that originate from limestone springs (or are
very strongly influenced by limestone springs) and are usually characterized with
alkalinities > 140 mg/L; a water temperature range of 40-65 0F (4-18 0C); and
drainage areas < 20 sq. miles. Limestone streams will often display low-gradient flow
conditions and are almost completely void of clearly defined freestone habitat beds.
Low-Gradient/Multihabitat streams: Lower gradient streams where cobble/gravel
substrate (riffle/run) habitat does not dominate (may sometimes be absent) but is
also characterized by snags (submerged woody debris), pools, and depositional
areas of coarse-particulate organic matter (CPOM), sand, and other fine sediments.
Riffle/Run habitat: Riffle conditions are demonstrated when the water flowing over the
substrate is shallow enough to create a broken, rough, and turbulent white water
surface. Runs are stream segments where deeper flowing water keeps the
gravel/cobble substrate completely submerged, which creates a smoother, less
turbulent, non-white water surface. Riffle/Run habitats are generally considered to
represent a stream’s most productive macrobenthic areas to sample.
A. Net Mesh Considerations
One area of concern relating to the Quality Control of statewide biological sampling is
standardization of the mesh size on various types of benthic macroinvertebrate
sampling gear. Without standardization of mesh size, standardization of overall methods
is of limited value.
Benthic macroinvertebrates have historically been defined as animals large enough to
be retained by a U.S. Standard No. 30 sieve (595 micron openings). A review of
sampling equipment that was in use and commercially available during the early
development of DEP’s water quality program indicated that the 595 micron criterion was
very seldom met. DEP hand-screens have mesh with about 800 - 1000 micron (µ)
openings. Standard Surber nets have mesh openings of 1024µ (silk) or 1050µ (nylon).
Surber nets of 728µ and 850/900µ are also available. Standard D-frame nets had
800 x 900µ openings.
It was apparent from the above discussion that the most common mesh size in use for
many years was in the 800-900µ range. Consequently, this size range was adopted and
has been DEP’s standard for many years. Multifilament nylon screen cloth with 800 900µ mesh was used for kick screens to ensure consistency. This 850/900µ mesh size
was also the standard for replacement Surber nets.
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In recent years, many state water quality programs, federal agencies (e.g. EPA, USGS),
and other water quality monitoring organizations began using net sampling devices with
500µ
µ
adopted for D-frame nets used in the Department’s PaDEP-RBP sampling protocols.
The one exception is the PaDEP-RBP protocol for limestone streams where an 800 900µ mesh D-frame net is used because of the nature of sampling true limestone
streams (described below).
Except as noted above for limestone streams, future references to the D-frame sampler
in the document assume 500µ mesh netting. The net mesh size of other screen
samplers and limestone stream employed D-frames have not changed and still are to
be 800-900µ. Because of differing net mesh size requirements of various PaDEP
protocols, the mesh size of the sampler and sampling protocol used must be noted on
field and bench identification sheets for the collected benthic sample.
B. Qualitative Methods
The type of sampling gear used is dependent on survey type and site-specific conditions. The recommended gear for sampling wadeable streams is 3 x 3 ft. flexible kickscreens and 12 inch diameter round D-frame nets. In larger streams or rivers, grab-type
samplers may be used to obtain qualitative samples. While generally thought of as
quantitative devices, Eckman, Peterson, or Petite Ponar grab samplers can also be
used to obtain qualitative data. The type of gear, dimensions, and mesh size must be
reported for all collections. When more than one gear type is used, the results must be
recorded separately.
Physical variables should be matched as closely as possible between background and
impact stations when selecting locations for placement of the sampling gear within each
station. Matching these variables helps minimize or eliminate the effects of
compounding variability.
Macrobenthos often exhibit clustered distributions, and if the sampling points are
selected in close proximity to each other, a single clustered population may be obtained
rather than a generalized measure of the overall population within the selected subhabitat. Spacing the sampling points as far apart as possible within the sub-habitat can
minimize the problem of clustered distributions.
B.1. Kick-screen. A common qualitative sampling method uses a simple hand-held
kick-screen. This device is designed to be used by two persons. However, with
experience, it may be used by one person and still provide adequate results. The kickscreen is constructed with a 1x1m piece of net material (800-900 µ mesh size) fastened
to two dowel handles (approximately 1”d. X 4’ long).
B.1.a. Traditional Method. Facing up stream, one person places the net in the stream
with the bottom edge of the net held firmly against the streambed. An assistant then
vigorously kicks the substrate within a 3x3 ft area immediately upstream of the net to a
depth of 3-4” (approximately 10cm). The functional depth sampled may vary due to
ease of disturbance as influenced by substrate embeddedness.
The amount of effort expended in collecting each sample should be approximately
equivalent in order to make valid comparisons. The effort, expressed as area, must be
reported for all collections.
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Collect a minimum of four screens at each site or until no new taxa are collected. Initial
sampling should be conducted in riffle areas. Collection in additional habitats to
generate a more complete taxa list can be conducted at the discretion of the
investigator. Initial analysis of the data must be limited to the riffle data for
standardization. A second analysis including other habitats may be conducted as
needed.
Data observations shall be recorded on a Flowing Waterbody Field Form (Appendix A)
created for each station sampled. Record the relative abundance of each recognizable
Family in each individual collection in the field. Relative abundance categories, with the
observed “total” ranges indicated in parenthesis, include: rare (0-3), present (3-10),
common (11-24), abundant (25-99), and (occasionally) very abundant (100+). The
investigator, at his/her discretion, may elect to enumerate certain target taxa.
The results of each collection are recorded to document conditions of the site for future
decision making.
a. A stressed or enriched community often exhibits little variability in community
structure over an area while a healthy community should have a more complex
structure. If varied taxa are found on each screen, the community is probably
complex, while the presence of only a few dominant taxa on every screen indicates
the community is a simple one.
b. Collecting intolerant taxa in a majority of screens is a good indication of an
unstressed community. However, collecting intolerant taxa in only one out of four
screens may be an indication that the intolerant taxa have only a marginal existence
at that location. A comparison of the composited taxa lists for each location may not
indicate the rarity of the intolerant taxa, but this rarity would be readily apparent if the
taxa lists for individual screens were compared.
c. Separate screen taxa lists provide information concerning the distribution of taxa. For
example, mayflies are taken in one of four screens at the background station and in
none of the four screens at the impact station. All the other taxa collected at both the
stations are tolerant forms. Based on a composited taxa list for each station, one
might conclude that the impact station is depressed due to the absence of mayflies.
However, the individual screen taxa lists would indicate that the mayflies may have a
clumped distribution and there is a possibility that the collector simply missed the
clumps at the impact station. This will be apparent to the biologist while in the field
and he/she can continue collecting until comfortable that mayflies are indeed absent
or less abundant at the impact station. Later, it can be reported, for example, that 4 of
10 screens contained mayflies at the background station while only 1 of 10 screens
contained mayflies at the impact station. This is an instance when the collector, while
still in the field, may choose to count the mayflies in each screen (especially if the
background screens had many mayflies while the impact screens only had one or
two).
d. Separate screen data can lend weight to an analysis when classification techniques
(ordination or clustering) are used. Results that cluster or score the individual
background screens differently than the individual impact screens indicates a
difference between the locations. When the classification technique scores
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background and impact screens in an apparent random manner, then it is likely that
there is no impact or that the natural variability is large and masks any impacts.
Individuals of representative taxa for a station may be composited in a single vial and
preserved for later laboratory verification or identification. Generally, the level of
taxonomic identification would follow that as listed in Section E.1.
Answers to several questions that require collector judgment and other taxa notes, can
be useful in subsequent analysis and can be stored with the taxa lists as remark fields.
Such questions include but are not limited to: What are the dominant and rare taxa? Are
there any taxa that are found to be unusually abundant?
B.1.b (SSWAP) Assessment Method. This method was used for assessments
conducted as part of the Statewide Surface Waters Assessment Program (SSWAP). It
employed the same kick screen gear, physical disturbance techniques, and relative
abundance determinations as the traditional method (B.1.a). The main difference is that
only two kicks were usually required and macroinvertebrate identifications were done
streamside to family level taxonomy with hand-held lens (10X) if necessary. Data was
recorded on field forms (Appendix A). This method is no longer used for official use
Assessment purposes but is kept here for historical reference and as a quick field
assessment tool during survey planning reconnaissance.
B.2. D-Frame. The handheld D-frame sampler consists of a bag net attached to a halfcircle (“D” shaped) frame that is 1 ft. wide. The net’s design is that of an extended,
tapering bag (routinely 500µ mesh size except for true limestone streams 800-900µ
mesh size).
B.2.a Sampling Riffle/Run Habitat. This D-frame methodology is basically the same
as with the kick-screen - except for the following points: The net is employed by one
person facing downstream and holding the net firmly on the stream bottom. One “Dframe effort” is defined as such: the investigator vigorously kicks an approximate area
of 1m2 (1X1 m) immediately upstream of the net to a depth of 10cm (or approximately
4”, as the embeddedness of the substrate will allow) for approximately one minute. All
benthic dislodgement and substrate scrubbing should be done by kicks only. Substrate
handling should be limited to only moving large rocks or debris (as needed) with no
hand washing. Since the width of the kick area is wider than the net opening, net
placement is critical in order to assure all kicked material flows toward the net. Avoiding
areas with crosscurrents, the substrate material from within the 1 m 2 area should be
kicked toward the center of the square meter area – above the net opening.
B.2.b Sampling Low-gradient Habitat. Geomorphological processes in low-gradient
streams create a variety of habitats (Multihabitat conditions) that cannot be sampled like
riffle/run habitats. Therefore, with the exception of still collecting stationary D-frame
kicks (described in B.2.a above) from any riffle/run habitat that may be present, for the
other habitat types encountered, the D-frame net is used to sweep or ‘jab’ through a
given area of substrate. A Multihabitat sample consists of a compilation of 10 D-frame
collections - jabs and riffle/run kicks - as needed: distributed proportionally from the
available habitat types within each 100 meter sample reach. Each jab consists of a 1meter-long sweep of a 0.3-meter wide area, using a D-frame dip net (500 micron mesh).
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One jab equals disturbing and capturing organisms from an area of ~0.23 m 2 (12” x
30”).
C. Semi-Quantitative Methods (PaDEP-RBP Sampling Protocols):
In Plafkin (1989), EPA presented field-sampling methods designed to assess impacts
normally associated with pollution impacts, cause/effect issues, and other water quality
degradation problems in a relatively rapid manner. These are referred to as Rapid
Bioassessment Protocols (RBPs). Barbour et. al. (1999) made revisions to these RBPs
and expanded them to include Multihabitat methodology. The PaDEP-RBP methods are
bioassessment techniques involving systematic field collection and subsequent lab
analysis to allow detection of benthic community differences between reference (or
control) waters and waters under evaluation. The PaDEP-RBPs are modifications of the
EPA RBP III (Plafkin, 1989; Barbour, et al 1999 revisions) and Multihabitat methodology
(Barbour, et al 1999) designed to be compatible with Pennsylvania's historical database.
Modifications include: 1) the use of a D-frame net as the collection gear for all PaDEPRBP sampled stream and habitat types, 2) different laboratory sorting procedures, 3)
elimination of the CPOM (coarse particulate organic matter) sampling (except for the
Multihabitat protocol), 4) number of multihabitat jabs; and 5) metrics substitutions.
Unlike the EPA’s RBP III methodology, no field sub-sample sorting is done. Only larger
rocks, detritus, and other debris are rinsed and removed while in the field before the
sample is preserved. Further, because the debris and detritus fraction of the samples
may at times be extremely voluminous, rinsing and discarding as much of the materials
as practical in the field prior to jar preservation and transport is strongly encouraged as
it will facilitate easier and more efficient lab processing. While EPA’s RBP methods
were designed to compare impacted waters to reference conditions (cause/effect
approach), the PaDEP-RBP modifications were designed for un-impacted waters, as
well as impacted waters.
C.1. Sample Collection. The purpose of the standardized PaDEP-RBP collection procedure is to obtain representative macroinvertebrate fauna samples from comparable
stations.
For most of the PaDEP-RBP sampling protocols, the riffle/run habitat is targeted for
sampling as it is routinely the most productive habitat. Exceptions to this rule are
discussed in the Limestone Stream and Multihabitat protocols. Within each station
reach, the habitat is sampled in a downstream-to-upstream direction using the D-frame
net method described above. For all sampling methods, when compositing materials
collected from multiple D-frame kicks, care must be taken to minimize “wear and tear”
on the collected organisms when compositing the materials. It is recommended that the
benthic material be placed in a bucket filled with water to facilitate gentle stirring and
mixing. The number of D-frame efforts is dependent on the type of survey conducted as
described below:
C.1.a. Wadeable Freestone Stream Surveys. Wadeable freestone surveys
include but not are limited to; Antidegradation, Aquatic Life Use, Existing Use,
Instream Comprehensive Evaluation (ICE) and Cause and Effect surveys. For these
surveys, it is necessary to characterize macroinvertebrate fauna communities from
an area larger than a single riffle. Therefore, an Antidegradation, Aquatic Life Use,
Existing Use, ICE, or Cause and Effect survey station is defined as a stream reach
of approximately 100 meters in length. At each station, six “D-frame efforts” are
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collected. Make an effort to spread the samples out over the entire reach. Choose
the best riffle habitat areas and be certain to include areas of different depths (fast
and slow) and substrate types that are typical of the riffle.
C.1.b. Limestone Stream Surveys. For limestone stream surveys, two paired Dframe efforts are collected from each station - one from an area of fast current
velocity and one from an area of slower current velocity within the same riffle.
Limestone streams have low gradient often making it difficult to locate well
developed riffles. If there are no riffles in the sample area, a run or the best rock
substrate available is sampled. The resulting “D-frame efforts” (two) are composited
into one sample jar (or more as necessary). Because limestone samples are very
abundant in organic material, the sample is preserved in 95 % ethanol and returned
to the lab for processing.
C.1.c. Low-gradient Multihabitat Surveys. Aquatic macroinvertebrate samples
are collected using a multihabitat sample collection method modified from that
described in Barbour et al (1999). Organisms are collected from a maximum of five
different habitat types within the sample reach. Table 1 describes the five habitat
types and explains the different sampling techniques. Sample collection consists of
10 D-frame jabs: 2 from each of five habitat types or distributed proportionally from
the available habitat types within each 100 meter sample reach. Each jab consists of
a 30-inch-long sweep of a 0.3-meter wide area, using a D-frame dip net (500 micron
mesh).
Table 1. Stream Habitat Types and Sampling Techniques for Multihabitat Surveys
Habitat Type
Description
Sample Technique
Cobble/Gravel
Substrate
Stream bottom areas consisting of mixed
gravel and larger substrate particles;
Cobble/gravel substrates are typically
located in relatively fast-flowing,
“erosional” areas of the stream channel
Macroinvertebrates are collected by placing the net on
the substrate near the downstream end of an area of
gravel or larger substrate particles and simultaneously
pushing down on the net while pulling it in an upstream
direction with adequate force to dislodge substrate
materials and the aquatic macroinvertebrate fauna
associated with these materials; Large stones and
organic matter contained in the net are discarded after
they are carefully inspected for the presence of attached
organisms which are removed and retained with the
remainder of the sample; One jab consists of passing the
net over approximately 30 inches of substrate.
Snag (includes
beaver dams)
Snag habitat consists of submerged
sticks, branches, and other woody debris
that appears to have been submerged
long enough to be adequately colonized
by aquatic macroinvertebrates; Preferred
snags for sampling include small to
medium-sized sticks and branches
(preferably < ~4 inches in diameter) that
have accumulated a substantial amount of
organic matter (twigs, leaves, uprooted
aquatic macrophytes, etc.) that is
colonized by aquatic macroinvertebrates.
When possible, the net is to be placed immediately
downstream of the snag, in either the water column or on
the stream bottom, in an area where water is flowing
through the snag at a moderate velocity; The snag is
then kicked in a manner such that aquatic
macroinvertebrates and organic matter are dislodged
from the snag and carried by the current into the net; If
the snag cannot be kicked, then it is sampled by jabbing
the net into a downstream area of the snag and moving it
in an upstream direction with enough force to dislodge
and capture aquatic macro-invertebrates that have
colonized the snag; One jab equals disturbing and
2
capturing organisms from an area of ~0.23 m (12” x
30”).
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Habitat Type
Description
Sample Technique
Coarse
Particulate
Organic Matter
(CPOM)
Coarse particulate organic matter (CPOM)
consists of a mix of plant parts (leaves,
bark, twigs, seeds, etc.) that have
accumulated on the stream bottom in
“depositional” areas of the stream
channel; In situations where there is
substantial variability in the composition of
CPOM deposits within a given sample
reach (e.g., deposits consisting primarily
of white pine needles and other deposits
consisting primarily of hardwood tree
leaves), a variety of CPOM deposits are
sampled; However, leaf packs in highervelocity (“erosional”) areas of the channel
are not included in CPOM samples.
CPOM deposits are sampled by lightly passing the net
along a 30-inch long path through the accumulated
organic material so as to collect the material and its
associated aquatic macroinvertebrate fauna; When
CPOM deposits are extensive, only the upper portion of
the accumulated organic matter is collected to ensure
that the collected material is from the aerobic zone.
Submerged
Aquatic
Vegetation
(SAV)
Submerged aquatic vegetation (SAV)
habitat consists of rooted aquatic
macrophytes.
SAV is sampled by drawing the net in an upstream
direction along a 30-inch long path through the
vegetation; Efforts should be made to avoid collecting
stream bottom sediments and organisms when sampling
SAV areas.
Sand/Fine
Sediment
Sand/fine sediment habitat includes
stream bottom areas that are composed
primarily of sand, silt, and/or clay.
Sand/fine sediment areas are sampled by bumping or
tapping the net along the surface of the substrate while
slowly drawing the net in an upstream direction along a
30-inch long path of stream bottom; Efforts should be
made to minimize the amount of debris collected in the
net by penetrating only the upper-most layer of sand/silt
deposits; Excess sand and silt are removed from the
sample by repeatedly dipping the net into the water
column and lifting it out of the stream to remove fine
sediment from the sample.
The resulting number of “D-frame efforts” as required per survey type are composited
into one sample jar or as few as necessary. Care must be taken to minimize “wear and
tear” on the collected organisms when compositing the materials. It is recommended
that the benthic material be placed in a bucket and filled with water to facilitate gentle
stirring and mixing and reiterated that as much of the rocks, debris, and detritus be
rinsed and discarded as practical in the field prior to jar preservation and transport.
The sample is preserved in ethanol and returned to the lab for processing.
C.2. Sample Processing. Samples collected with a D-frame net are generally
considered to be qualitative. However, the preserved samples can be processed in a
manner that yields data that is “semi-quantitative” - data that was collected by qualitative
methods but gives information that is almost statistically as strong as that collected by
quantitative methods.
The following procedures are adapted from EPA 1999 RBP methodology and used to
process qualitative D-frame samples so that the resulting data can be analyzed using
benthic macroinvertebrate biometric indices (or “metrics”). Equipment needed for the
benthic sample processing are:

2 large laboratory pans gridded into 28 squares* (more gridded pans may be
necessary depending on the size of the sample) DEP Central Office staff use
Rubbermaid 2 Gallon White Dur-x® Container 18" x 12" x 3 1/2" from US.
Plastic Corp. (catalog #6575)
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


http://www.usplastic.com/catalog/product.asp?catalog%5Fname=usplastic&cat
egory%5Fname=20369&product%5Fid=16540
slips of paper or other uniform objects (numbered from 1 to 28) for drawing
random numbers, and
forceps (or any tools that can be used to pick floating benthic organisms),
Grid cutters made from tubular material that approximates an inside area of 4
in2*.
* EPA’s (1989) gridding techniques suggested using “5 cm x 5 cm” (2”x2”) grids.
Existing equipment consisted of 14”x8”x2” pans which were conducive to dividing into
2”x2” grids and thus, contained 28 squares. The 4 in 2 grid cutters conform to these
pan dimensions. While pan size is not critical, the number of grids (28) must be
maintained if any basic density comparisons wish to be made between samples.
Grid cutters (or similar subsampling devices) used with different sized pans should
conform to the pans’ grid dimensions.
The procedures described below begin with the premise that the collected samples
have been properly composited according to the type of survey. For Antidegradation,
Aquatic Life Use, Existing Use, ICE and Cause-Effect surveys, a station sample
represents a composition of six D-frame efforts (collected from fast and slow riffle areas
in a 100 meter reach). For Limestone surveys, a station sample is a composition of two
D-frame efforts. For Multihabitat surveys, a station sample is a composite of 10 D-frame
jabs.
Following the steps listed below; process each composited D-frame sample to render a
sub-sample size targeted for the specific survey type. The targeted sub-sample size for
the various PaDEP-RBP sampling protocols are presented below:
Table 2. Targeted Sub-Sample Sizes for PaDEP-RBP Sampling Protocols
Sampling Protocol
Standard (Andtidegradation,
Aquatic Life Use, Existing
Use, Cause/Effect,
Multihabitat)
Limestone Stream
Sub-Sample Size Requirements of
Identifiable Organisms^ 20%
Range
Target
#
Preferred
Required 20%
200
160 - 240
190 - 210
300
240 - 360
280 - 320
^ “Identifiable Organisms” – organisms identified to the finest required taxon
resolution per Section E.1. This also excludes pupae, larval bodies missing too
many critical structures to render confident IDs, extremely small instar larvae,
empty shells or cases, and non-benthic taxa.
#
PaDEP-RBP IBI and metric scoring thresholds are based on the respective subsample target + 20% size range of identifiable organisms. However, at times, it
has been found that the lower end of the targeted range may not render enough
identifiable organisms. The sample processor may have sorted just over the lower
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-20% of the target range only to find out later during the ID step that some
individuals could not be identified to genus and had to revert to their family
taxonomic level. If a taxonomic Family count is represented by several genera
counts and a count of organisms identified to the Family level, either the Family
total count or the genus counts can be used in the IBI and metrics analysis. For
example, all the genera counts must be bumped up to be included in the Family
count or only the genera counts can be used and the Family counts left out.
Either of these scenarios results in weaker scores or, in the case where excluding
Family counts drops the sub-sample total below the acceptable lower range limits,
invalidate the IBI or metrics analysis. In order to assure IBIs or metrics analyses
are usable, sub-sampling to the narrower range is preferred and recommended to
minimize the probability of these invalidating sub-sampling scenarios occurring.
Therefore, it is prudent to exceed the 200 organism target to ensure adequate
individuals that can be identified.
C.2.a Processing Riffle/Run Habitat Samples collected by PaDEP-RBP
Antidegradation, Aquatic Life Use, Existing Use, ICE and Cause/Effect
Protocols.
1) The composited sample is placed in a 28-square gridded pan (Pan1). It is
recommended to remove fine materials and residual preservative prior to subsampling by rinsing the sample through a sieve (Standard USGS or sieve bucket)
that matches the mesh size (or smaller) of the net gear used to collect the sample.
2) The sample is gently stirred to disperse the contents evenly throughout Pan1 as
thoroughly as possible. (In order to ease mixing and to minimize “wear-and-tear” on
delicate organisms, water may be added to the pan to the depth of the sample
material before stirring)
3) Randomly select a grid using the 28 random number set and, using the grid cutters,
remove the debris and organisms entirely from within the grid cutter (centered over
the selected grid and “cut” into the debris) and place removed materials in a second
gridded pan (Pan2).
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Figure 1. Pan1 with 4 grid cutters in randomly selected grids.
i.
Float and pick, count, and sub-total all identifiable organisms (as noted above in
Table 2) from each cut grid placed in Pan2. Repeat until at least 4 grids have
been sub-sampled from Pan1. If, after 4 Pan1 grids have been sorted, the subtotal is less than the respective preferred sub-sample size (Table 2), then
continue to remove and sort grids one at a time until the preferred sub-sample
size is obtained from Pan2. If the benthic organism yield from the 4 Pan1 grids
exceeds the targeted sub-sample size, then proceed to Step ii.
ii. With all of the 240+ (or 360+) identifiable organisms remaining in Pan2, randomly
select one grid and “back count” (removing) all the organisms from that grid.
Repeat one grid at a time until the bug count remaining in Pan2 satisfies the
targeted 200 or 300 +
% rule (190-210 or 280-320 preferred as noted in Table
2).
iii. Because limestone samples are often characterized by dense invertebrate
concentrations, it is important not to exceed the upper boundary range (360+) of
the limestone targeted 300 subsample size. If it appears that the number of
benthic organisms from the last grid will cause the sub-sample to exceed its
target size by more than 20% (>360 organisms), count this last grid and place in
a separate clean gridded pan (Pan3) with enough water to facilitate gentle stirring
and even distribution. Randomly select grids from Pan3 and remove individuals
until the count of organisms remaining in Pan3 allows the subsample count to fall
below the 360 upper limit.
4) If not identified immediately, the sub-sample should be preserved and properly
labeled for future identification.
5) The benthic material remaining (Pan1) after the target sub-sample has been picked
can be returned to its original sample jar and preserved. They shall be retained in
accordance with QA retention times as specified for this respective survey type.
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6) Any grid chosen must be picked in its entirety (If picking the last grid causes the subsample to exceed its target size by more than 20% (>240 or >360 organisms), follow
back counting step C.2.a.3.iii above).
7) Record the final grid counts selected for each gridding phase (Pan1, Pan2, and
Pan2 “back counting” as necessary) on the lab bench ID sheet for the sample.
C.2.b Processing Samples collected by the PaDEP-RBP Multihabitat Protocol.
Multihabitat samples will usually require multiple sample jars because of the complexity
and degree of detritus and inorganic materials that characterize these samples.
Because of this, multihabitat samples need to be processed in a different manner than
those samples collected by other protocols.
Initial Processing of Multihabitat Macroinvertebrate Samples
1) Similarly as in C.2.a Step 1 above, multihabitat samples should be rinsed in
a standard USGS No. 35 sieve (or sieve bucket) to remove fine materials
and residual preservative prior to sub-sampling. Because of the nature of
multi-habitat samples (i.e. greater volumes of finer silts, organic, and
particulate material), rinsing while retaining the macroinvertebrates may
pose extra challenges. Using larger containers (such as five-gallon buckets)
and rinsing smaller aliquots of material may be necessary. All multihabitat
sample cleansing steps should use gentle and careful rinsing and agitation
to minimize organism damage.
2) Once the total sample has been properly rinsed, transfer the contents of the
sieve into a clean, white, 3.5” deep rectangular gridded pan (Pan1)
(measuring 14” long x 8” wide on the bottom of the pan) marked off into 28
four-square inch (2” x 2”) grids and follow Steps 2-7 as listed above in C.2.a
to target a 200 sub-sample (190-210 preferred) as indicated in Table 2.
C.2.c Processing larger, excessive amounts of D-frame sample debris and
multihabitat samples
Hopefully, the collector will rarely have very large amounts of D-frame materials to
process. The reduction of large materials by careful removal, inspection, and rinsing in
a bucket or using a sieve prior to field preservation or at the lab is encouraged.
However, if the amount of material composited in the field jars exceeds the functional
sorting capacity of Pan1, then follow this guidance:
a) Evenly distribute the material between as many pans as necessary.
b) From each pan (Pan1a, Pan1b, etc.), remove debris and organisms from 4 random
grids and place in Pan2 as described in Step C.2.a.3.i above.
c) Once the required 4 grids from each Pan1(a, b . . . etc.) have been placed in Pan2,
evenly and gently redistribute the materials as in Step C.2.a.3.ii.
d) Then, resume processing, again as described in Step C.2.a.3.iii, selecting a grid
from Pan2 and placing the materials into a gridded Pan3.
e) Process this material and repeat as described in Step C.2.a.i until the preferred subsample size is obtained from Pan3.
11
f) If, after processing 4 grids, the +20% upper limit (240+ or 360+) is exceeded, follow
“back counting” method in Step C.2.c.ii or iii (for limestone samples)
g) Once the targeted sub-sample is reached, continue with Step C.2.a.4-7.
D. Quantitative Methods
The type of sampling gear used is dependent on survey type and site-specific
conditions. The recommended gear includes Surber-type samplers (with 800 - 900
micron mesh), artificial substrate (multi-plate) samplers (meeting specifications in
Section D.2), and grab-sample devices. The type of gear, dimensions, and mesh size
must be reported for all collections. When more than one gear type is used, the results
must be recorded separately.
In order to limit the complications arising from compounding variability, follow the
guidelines discussed in Section II when selecting background stations or reference
water bodies and impact stations. Physical variables should be matched as closely as
possible between the background and impact stations when selecting locations for
placement of the sampling gear within each station. This helps minimize or eliminate the
effects of compounding variability. An additional consideration is locating the samplers
in areas where current, depth, and substrate are optimal for gear efficiency. It should be
clearly noted on field forms when sampling gear is used under sub-optimal conditions
(i.e. slow current, bedrock etc.).
Macrobenthos often exhibit clustered distributions and if the sample points are selected
within close proximity, only a single clustered population sample may be obtained rather
than a generalized measure of the overall population within the selected sub-habitat.
Spacing the sampling gear as far apart as possible within the sub-habitat can minimize
the problem of clustered distributions.
D.1. Surber-type Samplers. Surber-type samplers are defined as samplers that
delineate or confine an area of the stream bottom (usually 1 ft 2), which is to be sampled
by disturbing the enclosed substrate. The dislodged materials (organisms, detritus, and
other debris) are swept into an attached, tapering net. These samplers include the
Surber Sampler, Portable Benthic Invertebrate Sampler (PBIS), and Hess Sampler. The
sampling procedure for all these samplers and other similar devices are as follows:
a. The substrate will be completely disturbed within the confines of the sampler
frame to a depth of 3-4” (approximately 10cm). Larger rocks should be gently, but
thoroughly “scrubbed” while being held in the net mouth.
b. Collect a minimum of three quantitative samples at each site. Do not composite
samples, but place each collection in a separate container.
c. These quantitative samples may be processed in the manner described in
Section C.2), except that once all organisms have been floated and picked from
the debris, it is not necessary to pick a sub-sample from a gridded pan.
Evaluations using organisms collected by these quantitative methods rely on the
total number of individuals from the entire sample.
d. The organisms may be identified to the taxonomic level deemed necessary by
the collector and problem being investigated. Samples may be split to expedite
processing. Procedures for sub-sampling are defined in Elliott (1977) and in EPA
12
(1990). In addition, a plankton splitter may be employed to sub-sample large
numbers of chironomid larvae or other abundant taxa.
e. Complete a coded field form to ensure that the associated physical data is
recorded.
D.2. Multi-plate Samplers. Multi-plate artificial substrate samplers (Figure 1, page 17)
may be used where water depth and/or substrate prevents the use of other sampling
techniques. A description of the sampler used including the type of artificial substrate,
the individual component dimensions, and total surface area must be reported with each
collection. In continuing investigations, the samplers must be uniform from year to year.
Physical variables should be matched as closely as possible when using the samplers
on an annual basis at the same location and between stations for cause/effect surveys.
Matching these variables helps minimize or eliminate the compounding of variables. An
additional consideration is locating the samplers in areas where the current, depth, and
substrate are optimal for gear efficiency while minimizing problems of theft and
disturbance.
Multi-plate samples for WQN stations must conform to the following procedures:
a. Place a minimum of two samplers at each site to help assure the retrieval of at
least one.
b. Leave the samplers in place for a minimum of 6 weeks. The amount of time the
samplers are in place must be reported with each collection.
c. Samplers should be enclosed in a net (of equal or smaller mesh size) or plastic
bag before retrieval to prevent loss of organisms. When the use of nets or bags
is not possible, retrieve the sampler with a smooth but rapid motion to minimize
loss of organisms. The retrieved sampler should be immediately placed in a tray,
scraped, and all the scrapings preserved. Do not composite the samples.
Preserve each separately.
d. The taxa should be identified to genus whenever possible. Samples may be split
to expedite processing. Procedures for sub-sampling are defined in Elliott (1977)
or EPA (1973).
e. Complete a coded field form to insure that the associated physical data is
recorded.
D.3. Grab Samplers. Where standard shallow-water sampling methodology is not
feasible, grab-sampling devices may be necessary. These include Ekman, Peterson, or
Ponar-style grab samplers. They are designed for use in deeper waters or in areas that
have soft, unconsolidated substrate. These samplers are somewhat cumbersome and
labor intensive to use. They are heavy, by design, so that they can be dropped from a
boat and penetrate the substrate. They often need a boom and pulley retrieval system.
For specific discussions on the advantages, disadvantages, and sampling methodology,
refer to EPA 1990.
13
E. Identification
E.1. Taxonomic Level. The level of identification under magnification for most aquatic
macroinvertebrates will be to genus based on the recommended references listed in the
Taxonomy Reference List on the 2013 Assessment Methods webpage. Some
individuals collected will be immature and not exhibit the characteristics necessary for
confident identification. Therefore, with the exception of the Ephemeroptera, Plecoptera
and Trichoptera (EPT) taxa, the lowest level of taxonomy attainable will be sufficient.
EPT taxa must be identified to the Genus level. The following taxonomic groups,
however, may be identified to a higher taxonomic level as follows:
Snails (Gastropoda) - Family
Clams, mussels (Bivalvia) - Family
Flatworms (Turbellaria)
identifiable planarids - genus
or Family Planaridae
others - Phylum Turbellaria
Segmented worms (Annelida)
aquatic earthworms & tubificids - Class Oligochaeta
leeches - Class Hirudinea
Moss animacules - Phylum Bryozoa
Proboscis worms – Phylum Nemertea
Roundworms - Phylum Nematoda
Water mites- “Hydracarina” (an artificial taxonomic grouping of several mite
superfamilies)
Midges (Chironomidae) – Family
Taxonomic references can be found in the Taxonomic Reference List included with the
2013 Assessment Methods.
E.2. Verifications. For Quality Assurance purposes, certain laboratory invertebrate
processing procedures should be checked routinely. Normally, a colleague may perform
these spot checks. These include the floating/picking steps, taxonomic identifications,
and total taxa list scans:
a. Sorting. After the floating and picking has been completed for samples that
require this treatment (Pa-RBP, Surber-type, multi-plate, and grab samples), the
residue should be briefly scanned before discarding to assure that the sample
has been sufficiently “picked”. This should be done for 10% of the samples (or at
least one sample) per survey.
b. Identification. For samples not involving litigation or enforcement issues,
laboratory bench ID sheets for all samples should be reviewed. Any unusual
taxa or taxa that are not typical to the type of stream or water quality condition
that was surveyed, should be checked. For samples involving legal issues,
representative specimens of each taxon may need to be verified by independent
expert taxonomists. For each staff performing identifications, a minimum of 10%
of the samples identified should be quality assured by another taxonomist.
E.3. Sample Retention.
For Quality Assurance purposes, identified benthic
macroinvertebrate samples should be preserved and retained for later verifications.
Based on the nature and purpose of the survey, retention times would vary:
14
a. Cause/effect surveys: until all legal issues have been resolved.
b. Monitoring surveys:
1) WQN - 2 years
2) Reference WQN - 5 years
c. Stream Redesignation and Use-attainability surveys: until any proposed stream
classification changes become final (approximately 2 years).
d. Enforcement/compliance surveys: until all legal issues, including appeals and
related litigation have been resolved.
15
References
Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling. 1999. Rapid
Bioassessment Protocols for Use in Streams and Wadeable Rivers: Periphyton,
Benthic Macroinvertebrates and Fish. Second Edition. EPA/841-B-99-002. U.S.
EPA, Office of Water, Washington, D.C.
Elliott, J. M.; 1977. Statistical Analysis of Samples of Benthic Invertebrates. Freshwater
Biological Association, Publication No. 25.
Environmental Protection Agency. 1973. Biological Field and Laboratory Methods. Office
of Research and Development. EPA; Cincinnati, OH. EPA-670/4-73-001.
______. 1990.
Macroinvertebrate Field and Laboratory Methods for Evaluating the
Biological Integrity of Surface Waters, Office of Research and Development
publication: EPA/600/4-90/030, Nov. 1990.
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
macroinvertebrates and fish. EPA/440/4-89-001. U.S. Environmental Protection
Agency, Office of Water, Washington, D.C.
16
Figure 1. Multi-plate sampler diagram
17
Appendix A
Flowing Waterbody Field Form
18
3800-FM-WSFR0086
Rev. 12/2008
COMMONWEALTH OF PENNSYLVANIA
DEPARTMENT OF ENVIRONMENTAL PROTECTION
BUREAU OF WATER STANDARDS AND FACILITY REGULATION
FLOWING WATERBODY FIELD DATA FORM
(Information and comments for fields boxed in double lines are required database entries. Other fields are optional for personal use.)
Date-Time-Initials*
-
Example
20040212-0312-XYZ
Date
Watershed Code
(HUC)
Time
Stream Code
Ch. 93 Use
Initials
Surveyed by:
Secondary Station ID
SWP Watershed
*Date as YYYYMMDD, time as military time, and your initials uniquely identify the stream reach.
Survey Type
(1) Basin Survey, (2) Cause / Effect, (3) Fish Tissue, (4) Instream Comprehensive Evaluation [ICE], (5) Point-of-First-Use, (6) SERA, (7)
Antidegradation [Special Protection], (8) Toxics, (10) Use Attainability, (11) WQN, (12) Limestone, (13) Low-gradient [Multihabitat]
Location
County:
Municipality:
Topo Quad:
Location Description:
Land Use
Residential:
Abd. Mining:
%
%
Commercial:
Old Fields:
%
%
Industrial:
Forest:
%
%
Cropland:
Other:
%
%
Pasture:
%
Land Use Comments:
Canopy cover: open
partly shaded mostly shaded fully shaded
Water Quality
Collectorsequence #
0
Temp ( C)
Field Meter Readings:
DO
Cond.
(mg/L)
pH
(umhos)
Alkalinity
mg/l
Bottle Notes (N-normal, MNF-metals nonfiltered, MF-metals filtered, B-bac’t, Others:
indicate)
1.
2.
3.
Water Appearance/Odor Comments: (^see bottom of back for common descriptors)
Findings
Not
Impaired
Impaired
Is impact
Reevaluate
Impaired:
biology?
habitat?
localized?
designated use?
Decision comments. Describe the rationale for your “Not Impaired” or “Impaired” decision; reach locations for use
designation reevaluations; special condition comments; etc.:
IBI Score:
Total Habitat Score:
- 19 -
3800-FM-WSFR0086
Rev. 12/2008
Macroinvertebrate sampling
Sampling method: Std. kick screen:
D-frame:
Surber:
Other:
method?:
Comments/Abundance Notes:
Habitat Impairment Thresholds
Metric Score
#3 Riff/Run: embeddedness or #3 Glide/Pool: substrate character + #6 Sediment Deposition = 24 or less
(20 or less for warm water, low gradient streams)
#9 Condition of Banks + #10 Bank Vegetation = 24 or less (20 or less for warm water, low gradient
streams
Total habitat score 140 or less for forested, cold water, high gradient streams (120 or less for warm
water, low gradient streams)
Habitat Comments:
Special Condition
Use this block to describe conditions that justify attainment/impairment of stations with IBI score <63 and >53.
^Common descriptors: Water Odors - none normal sewage petroleum chemical other; Water Surface Oils - none slick sheen globs flecks;
Turbidity - clear slight turbid opaque; NPS Pollution - no evidence some potential obvious; Sediment Odors - none normal sewage petroleum
chemical anaerobic; Sediment Oils - absent slight moderate profuse; Deposits – none sludge sawdust paper fiber sand relict shells other. Are
the undersides of stones deeply embedded black?
- 20 -