I-14_BenthicMacroinvertebrateMonitoringSOP_signed

Title:
Version
Approvals
CITY OF
DURHAM
City of Durham, North Carolina
Department of Public Works
Stormwater and GIS Services
Water Quality Unit
Standard Operating Procedures for Benthic
Macroinvertebrate Community Monitoring
1.0
Approved by: l� &/ a Date:
Michelle Woolfolk, Water Quality Manager
Approved by:
2/3/2022
Date: February 3, 2022
Paul Wiebke, Assistant Director for Stormwater & GIS Services
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Revision Record
Revision
Date
Updated by
Description of change
1.0
8/13/2021
Joseph Smith
Minor edits
Distribution
Current copies of this approved SOP should be provided to the following positions with the City of
Durham Stormwater and GIS Services, Water Quality Unit:
• Water Quality Manager
• Assistant Water Quality Manager
• Environmental Planning & Compliance Sr. Analyst
• Environmental Planning & Compliance Analyst, Monitoring and Assessment
• Environmental Planning & Compliance Coordinator, Monitoring and Assessment
• Environmental Planning & Compliance Specialist, Monitoring and Assessment
• Other staff assigned to the project
A copy of this approved SOP should also be provided to any individuals or businesses contracted to
perform benthic macroinvertebrate monitoring services to the City of Durham Stormwater and GIS
Services.
Abbreviations
BI
Biotic Index
DA
Drainage area
EPT
Ephemeroptera (mayflies), Plecoptera (stoneflies), and Trichoptera (caddisflies)
NCDEQ
North Carolina Department of Environmental Quality
QAPP
Quality Assurance Project Plan
QA/QC
Quality Assurance/ Quality Control
SDS
Safety Data Sheet
SOP
Standard Operating Procedures
TV
Tolerance Value(s)
USGS
United States Geological Survey
WQU
Water Quality Unit
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Table of Contents
Title: Standard Operating Procedures for Benthic Macroinvertebrate Community Monitoring ..........1
Approvals..................................................................................................................................................
1
RevisionRecord........................................................................................................................................
2
Distribution...............................................................................................................................................
2
Abbreviations...........................................................................................................................................
2
Tableof Contents..........................................................................................................................................
3
Procedures....................................................................................................................................................
5
Purposeand scope...................................................................................................................................
5
Methodsummary.....................................................................................................................................
5
Healthand safety warnings......................................................................................................................
6
General safety considerations for fieldwork........................................................................................
6
Personal protective equipment............................................................................................................
7
Chemicalsafety....................................................................................................................................
8
Cautionsand interferences......................................................................................................................
8
Personnel qualifications and responsibilities...........................................................................................9
Equipmentand supplies...........................................................................................................................
9
Fieldsampling.......................................................................................................................................9
Methods.................................................................................................................................................10
Site selection and characterization....................................................................................................10
Preparation for field sampling............................................................................................................
11
Selection of sampling method............................................................................................................11
Fieldmeasurements...........................................................................................................................12
Benthic macroinvertebrate sampling.................................................................................................
12
Field processing of samples (sample "picking").................................................................................18
Habitat assessments and Stream Survey Collection Card..................................................................18
Digitalphotographs............................................................................................................................19
Laboratory analysis and taxonomic identification.............................................................................19
Data analysis and reporting................................................................................................................
20
Data and records management..........................................................................................................
24
Quality assurance and quality control(QA/QC)..........................................................................................
24
References..................................................................................................................................................
26
Appendix 1. Habitat Assessment Field Form..........................................................................................
27
Appendix 2. Collection Card Field Form.................................................................................................
32
Appendix 3. Taxa list for seasonal correction of Full Scale samples......................................................33
Appendix 4. NCDEQ Taxa List and Tolerance Values (TVs).....................................................................34
Appendix 5. Resources for Identification of Freshwater Macroinvertebrates.......................................44
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Tables
Table 1 Descriptions of required samples for Full Scale and Qual4 sampling methodologies ...................13
Table 2 Wentworth scale for classification of stream substrate based on size..........................................19
Table 3 Relative abundance codes for laboratory analysis of benthic macroinvertebrate samples .......... 2O
Table 4 Description of metrics calculated from benthic macroinvertebrate community data..................2O
Table 5 Biotic index (BI) thresholds for determining Bioclassification of Qua14 samples .......................... 21
Table 6 Look -up table for Scores associated with EPT S and BI.................................................................. 23
Table 7 Seasonal corrections for BI for Full Scale samples......................................................................... 23
Table 8 EPT N criteria for rounding decisions using Full Scale criteria....................................................... 23
Table 9 Assignment of final Bioclassification for Full Scale samples.......................................................... 24
Figures
Figure 1 Sampling method selection flow chart.........................................................................................12
Figure 2 Water Quality staff demonstrating the proper use of a kick net..................................................14
Figure 3 Washing down the kick net into a sieve bucket to remove the captured benthic organisms .....15
Figure 4 Water Quality staff member sampling submerged aquatic plants with a sweep net..................15
Figure 5 Water Quality staff member demonstrating a woody debris wash using a fine -mesh sampler..15
Figure 6 Rock -log wash sample being processed into a fine -mesh sampler..............................................15
Figure 7 Water Quality staff member preparing to elutriate a sand sample.............................................16
Figure 8 Water Quality staff member pouring elutriate from a sand sample into a fine -mesh sampler ...
16
Figure 9 An example of an Elliptio complanata mussel found in New Hope Creek....................................17
Figure 10 Sampler conducting visual searches for aquatic organisms.......................................................17
Figure11 Axes of sediment.........................................................................................................................
19
Figure 12 Flow chart for calculation of Bioclassifications for Full Scale samples.......................................22
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Procedures
Purpose and scope
The purpose of these Standard Operating Procedures (SOP) is to describe the methods used by the City
of Durham's Stormwater and GIS Services Division, Water Quality Unit (WQU) for:
• Site selection and characterization
• Field collection of benthic macroinvertebrate samples
• Completion of instream habitat assessments
• Taxonomic identification of samples
• Analysis of results
This type of monitoring provides information about the impacts of water quality, instream habitat, and
hydrology on instream biological communities.
While the procedures described in this document are based on widely accepted methods developed by
other agencies (see NCDEQ 2016), this SOP is only applicable to benthic macroinvertebrate monitoring
conducted as part of the WQU's water quality monitoring projects. As such, these procedures will
generally be limited to assessment of wadeable freshwater streams and rivers with visible flow within
the jurisdictional limits of Durham City and Durham County. While the terms and methodologies are
similar to those used by NC Department of Environmental Quality (NCDEQ), the WQU has different site
criteria for determining applicability of methods for certain sampling locations within the Triassic Basins
ecoregion (Griffith 2002).
Method summary
Benthic macroinvertebrates are invertebrate animals that are visible to the naked eye most often
associated with the substrate (benthos) of streams, rivers, and other water bodies. Many benthic
macro invertebrates are aquatic insects (both larval and adult), though the term also includes organisms
such as mollusks, crayfish, and oligochaetes (worms). There is great diversity in terms of the number of
benthic taxa found in eastern North American freshwaters, and consequently great diversity in life
histories and sensitivities to pollutants and other stressors. Because of these characteristics,
macroinvertebrate community surveys can serve as one type of indicator of biological integrity in
streams and rivers (NCDEQ 2016). These surveys, in conjunction with other water quality data, can be
used to assess biological responses to long-term environmental conditions that may not be adequately
addressed by short-term or discrete sampling efforts.
The primary purpose of the methods described in this document is the determination of a categorical
bioclassification of water bodies under study. The bioclassification for a stream is based on the diversity
and overall tolerance to stressors of the benthic macroinvertebrate community. There are five possible
bioclassifications (from best to worst): Excellent, Good, Good -Fair, Fair, and Poor. These bioclassification
ratings are based on tolerance values (TV) generated by the NCDEQ Biological Assessment Branch, using
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established procedures outlined in A Biotic Index for the Southeastern United States: Derivation and List
of Tolerance Values, with Criteria for Assigning Water Quality Ratings, 1993.
The methods for benthic macroinvertebrate monitoring conducted by the WQU include the following
major steps:
• Selection and characterization of monitoring sites and sampling methods
• Collection of benthic macroinvertebrates from a variety of instream habitats at a stream site
• Field processing and preservation of specimens in accordance with their relative abundance
• Laboratory identification of preserved specimens to the lowest possible taxonomic level and
documentation of the relative abundance (Rare, Common, or Abundant) of each taxon
• Calculation of a bioclassification and associated metrics using the taxonomic identifications,
relative abundances, and relative tolerance to stressors for each taxon
Health and safety warnings
General safety considerations for fieldwork
The WQU conducts sampling throughout the City of Durham and surrounding areas at times and places
where medical facilities may not be readily available. It is imperative that all employees receive
instruction and follow safety precautions when using equipment and handling hazardous materials to
prevent injury. "Safety first" must always be the rule.
All employees should receive a copy of the Department of Public Work's Safety Operating Procedures
(City of Durham 2017) and current Safety Policies (City of Durham Employee Policies S-201— S-208)
upon hire. Any injuries or incidents should be documented and recorded in accordance with City and
Department policies. The first priority should be to provide appropriate care (first aid or emergency
treatment) to the injured employee; the second priority is to notify the appropriate supervisor.
While the remainder of this section summarizes some of the more common hazards encountered during
fieldwork, all employees are expected to review and comply with all City procedures and policies.
One of the most prevalent situations encountered by field staff is walking over uneven and often wet or
muddy terrain while carrying heavy or bulky equipment. Streams in Durham are also often deeply
incised, with steep banks and soils that are clay -based and slick when wet. These conditions increase
risks for slips, trips, and falls. Many sites require working in urban areas where hazards such as roadway
traffic, domestic animals, and possibly wild animals can make access dangerous. In -stream hazards such
as glass shards, construction debris, medical waste, and miscellaneous trash are commonly present.
Staff must be vigilant to take proper precautions in order to decrease the chance of injury. Working in
streams also exposes staff to uneven, slick, or unstable substrates and potentially high flows, all of which
can challenge balance and lead to falls and even potential risk of drowning. Water depth and velocity
should always be considered during site reconnaissance and fieldwork.
Weather forecasts should be consulted prior to field sampling activities. Weather (such as heavy rain,
thunderstorms, lightning, high winds, and excessive heat or cold) can create unsafe conditions. In
certain cases, such as heavy rain or excessive heat or cold, fieldwork may be modified to minimize
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increased risks, such as falls, heat stress, or hypothermia, due to weather conditions. Staff should
immediately seek shelter or reschedule fieldwork if there are active thunderstorms with lightning or
other severe weather.
Personal protective equipment
The following PPE are required for field staff under most conditions encountered during sampling,
though they may be modified based on current weather conditions, project requirements, and site -
specific conditions.
• Appropriate footwear - chest waders, hip boots, knee boots, or steel toe work boots. The
primary purpose of protective footwear is to protect feet from abrasions and crushing injuries.
Field staff should not sample in shoes that do not cover the entire foot (such as sandals) or
shorts, neither of which offer full coverage of feet and lower legs. If monitoring activities require
wading in water bodies then waders, hip boots, or rubber knee boots should be worn. These
keep staff dry to reduce risks of cold -related stress and to reduce the potential exposure of staff
to pathogenic organisms and other contaminants that are commonly found in urban water
bodies. During the winter and spring, neoprene waders can provide additional protection from
cold temperatures and reduce risks associated with hypothermia. During warmer periods,
however, waders can increase the risk of heat -related illnesses. Staff can reduce these
additional risks by minimizing the time spent in direct sunlight, staying hydrated, and taking
frequent breaks (including removing waders) as needed.
• Long pants provide protection to the lower legs from abrasions and reduce potential for bites
from ticks, insects, and snakes.
• Long gloves/gauntlets may be helpful under certain situations, such as during cold weather
work to reduce cold -related stress, when sampling below a known discharge to minimize
exposure to pathogens, or to reduce the risk of cuts in the presence of hazards (glass shards,
etc.) within the stream.
• A hat, such as a baseball cap or wide -brimmed hat, helps to shade the head and face, which
helps reduce the potential for sunburn and heat -related illnesses.
• Polarized sunglasses reduce ambient glare and protect the eyes from reflection on the water.
• Sunblock/sunscreen reduces the risk of sunburn.
• Insect repellent is important for preventing vector -borne diseases and reactions (including
allergic reactions). Mosquitoes and ticks are very prevalent in Durham and can be active at any
time of the year, and therefore insect repellents containing DEET or other proven chemical
repellent should be worn during all field activities. Staff should check themselves for ticks at the
end of each field day and document any embedded ticks found.
• Poison ivy pre- or post -exposure treatment is necessary for staff that are prone to allergic
reactions to poison ivy or other plants. Affected employees should minimize exposure whenever
possible and take steps to remove irritating plant oils from their skin and clothes as soon as
possible after exposure to reduce the risk of allergic reactions. Soap -and -water is often effective
for removal, though may not feasible under field conditions. There are commercial products
(such as Tecnu) that are more easily used under field conditions. All staff should be mindful of
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transferring oils from their clothing to vehicle seats, etc., which can be a potential source of
exposure for other staff.
• First aid kits should be carried by all field staff during all fieldwork.
• High visibility vests or other clothing must be worn when working alongside roads, bridges, and
in the right-of-way.
• Orange safety cones should be used behind vehicles that are parked on the road or in the right-
of-way and the flashing warning lights on the City vehicle should be on to alert other traffic.
• Rain gear (jacket and pants) should be worn to keep staff dry and reduce risks of cold -related
illness.
• Additional safety equipment (flares, fire extinguisher, horn, etc.) may be required for boat or
field work and will be described in the project QAPP or study plan.
Chemical safety
City staff must follow any applicable City guidance and/or policies for activities such as chemical storage
and handling and the storage of preserved specimens. Chemical handling, storage, and disposal should
comply with requirements in associated Safety Data Sheets (SDS). When sampling unknown substances
in the field, staff should assume they are hazardous and wear appropriate PPE, such as disposable nitrile
gloves. The Stormwater Laboratory (SW Lab) is located in the Annex Building in City Hall and is equipped
with safety equipment such as eyewash stations and fully stocked first aid kits in case of emergency.
Staff should be familiar with the location of all SW Lab eye wash stations and first aid kit locations.
Laboratory sample processing and analyses (including chemical and biological samples) may be
performed by private contractors. Contractors are responsible for following all appropriate laws and
regulations as well as any internal City mandated safety procedures.
Cautions and interferences
Biological monitoring is especially sensitive to seasonality, skill of the field crew and taxonomist, and
short- and long-term weather and climatic events.
• It is critical that data collections, specimen identifications, and data analysis be performed under
the direction of an experienced biologist to ensure that results are truly representative of the
stream under study. Sampling conducted by inexperienced field staff should be overseen by
more experienced staff or the contractor to ensure that samples are correctly collected and
picked.
• The misapplication of sampling method (Qua14 or Full Scale) and associated metric calculations
can lead to erroneous bioclassifications. Sampling location and methodology should be
discussed with an experienced biologist prior to final site selection. Sampling methods are
further discussed in the Methods section.
• Sampling outside of the recommended seasons for each method will generally result in a lower
number of taxa collected, which can result in a lower bioclassification than would be found
during the designated sampling seasons. Sampling outside of recommended seasons should be
avoided. If necessary, corrections can be made based on seasonal correction factors developed
by NCDEQ. This is further discussed in the Methods section.
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• Extended periods of low stream flows, common in Triassic Basin streams, can stress instream
communities. This makes it difficult to determine if stressed biological communities are due to
natural events (such as drought) or due to anthropogenic sources (such as water quality issues).
• High flow events will often scour key instream habitat. Episodic high flows (such as after heavy
rains) will also cause many taxa to burrow or retreat into refugia, making them inaccessible
using the sampling methods described here. Instream habitats may not be feasibly sampled
during high flow events. Monitoring immediately after these types of events may result in
artificially depressed bioclassifications and habitat assessment scores.
• Deep and/or strong streamflows present safety issues, but also make key instream habitats
difficult or impossible to sample.
• Macroinvertebrates may also burrow or drift downstream in response to acute events such as
chemical spills or high sediment loads, which can also result in depressed bioclassifications.
• Calculation of bioclassifications by the WQU are based on the methods used by NCDEQ, with the
important difference of applying and reporting bioclassifications for streams in the Triassic
Basins with drainage areas <165 mi.z. NCDEQ does not rate these streams, but past studies by
the WQU have provided sufficient evidence that bioclassifications in these Triassic streams are
useful for the assessment of biological health of water bodies in conjunction with other water
quality monitoring efforts (City of Durham, 2011).
• All equipment should be thoroughly rinsed before leaving a site to minimize the potential for
carrying specimens to the next site. This also reduces the chances of transporting nuisance
species (such as invasive aquatic plants) to other areas.
Personnel qualifications and responsibilities
Field sampling and habitat assessments must be led by an experienced benthic biologist that is trained
and knowledgeable in benthic macroinvertebrate sampling methods and taxonomy. Before assisting
with sampling, all field staff must review this SOP and have an understanding of basic insect morphology
and common body forms of benthos found in Durham streams. Appendix 5 contains a basic visual key
and references to assist with basic identification of aquatic macroinvertebrates.
Oversight of field collections and laboratory identification of benthic macro invertebrate samples may be
performed by biologist(s) under contract with the City. Acceptable qualifications for the contract
biologist(s) may be described in the Request for Proposals or Request for Qualifications that is issued
when seeking a new contractor.
Equipment and supplies
Field sampling
• Kick net: 600 µm mesh nylon bolting cloth, approximately 1 m x 1 m, weighted on the bottom,
with attached wooden handles (approximately 1.3 m long)
• 1-2 sweep net(s): long -handled A- or D-frame net
• Sieve bucket: heavy duty bucket with US Standard No. 30 (600 µm) mesh bottom
• 2-3 plastic dish tubs
• 90% ethanol
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• Sample container: lidded tub container (about 1L volume) to hold ethanol, glass vials, and to
preserve larger specimens
• For each field person:
o Picking tray: shallow, white, plastic tray (such as developing trays used for photography)
o Fine point forceps
o Glass screw top vial (approximately 15-20 mL) with cap
• Sample labels and pencil for marking
• Field forms: habitat assessment form (Appendix 1) and "collection card" form (Appendix 2)
• Digital camera
• Multi -parameter water quality meter (temperature, pH, specific conductance, dissolved oxygen)
• Turbidimeter and sample vials
• GPS receiver (recreational grade)
• Aquatic macroinvertebrate field identification guide(s)
Additionally, at Full Scale sampling sites the following items are required:
• Fine -mesh sampler ("midge getter") and soaking/storage container; the sampler consists of a
piece of fine nitex mesh (300 µm) affixed between two 4-inch PVC pipe fittings, and a lidded
container large enough to hold the sampler and a sufficient volume of 90% ethanol.
• Sand sample bag; an open -top bag constructed of fine nitex mesh (300 µm).
Methods
Site selection and characterization
1. Consult the project's Quality Assurance Project Plan (QAPP) or study plan to determine criteria
for selection of representative sites that meet the objectives of the specific project. This can be
done through consulting maps, existing data, or other "desktop" screening tools.
2. Complete field reconnaissance of the proposed site(s). Sampling site(s) should meet the
following criteria:
a. Streams and rivers should be wadeable with visible stream discharge (flow) during the
growing season (March 1— September 30). Streams with very deep water, very strong
current, or high turbidity that are not safely wadeable should not be sampled using
these methods, or sampling should be scheduled during lower flow conditions. Streams
should generally have a wetted width of at least one meter.
b. Sites must have sufficient quantity and variety of suitable habitats (see Field Sampling
subsection below) to allow collection of all sample types required for the desired
sampling method.
3. Delineate the watershed boundary for each site. Possible tools to use for this include ESRI
ArcGIS, USGS' StreamStats web application, or hard copy topographic maps. Calculate the
watershed drainage area in square miles.
4. Determine the Level IV ecoregion (Griffith 2002) for the site and/or watershed using hard copy
or digital maps. Some watersheds may contain more than one ecoregion. In these cases, best
professional judgment may be needed to classify the predominant ecoregion for a site and
should take into account spatial coverage of each ecoregion within the watershed, proximity of
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each ecoregion to the sampling site, and local characteristics of the specific stream reach to be
sampled (such as flow, substrate, and soils).
5. Determine if sensitive species (e.g., threatened or endangered) are present in the area. This
information can be obtained from the NC Natural Heritage Program or from the NC Wildlife
Resources Commission. The presence of these types of species may require obtaining
permission from state or federal agencies or applying for a special collection permit.
6. Determine if the site is on public lands (e.g., state park) or other designated conservation area.
Contact the appropriate agency to determine if there are any requirements or restrictions for
collection of aquatic invertebrates.
Preparation for field sampling
1. Local weather conditions, (e.g., NC Drought Monitor or nearby US Geological Survey (USGS)
stream gages) should be monitored in the weeks and days prior to sampling to determine if
unusual conditions such as drought or heavy rain events exist that may affect site suitability for
sampling. Sampling should be rescheduled if it is determined that existing or forecasted
conditions are unsuitable.
2. Ensure that appropriate equipment will be available for use on the scheduled date(s) of
sampling and prepare necessary maps, field forms, and other materials required for field
assessments. All field staff must review this SOP, review basic insect morphology and common
body forms, and compile any necessary taxonomic references (see Appendix 5 for examples) for
use in the field.
3. All water quality field meters should be calibrated in the morning before use and the
calibrations checked at the end of the day, in accordance with WQU SOP (City of Durham, 2019).
Selection of sampling method
1. There are two basic methodologies that are currently used by the WQU: Full Scale (Standard
Qualitative) and Qual4. The selection of the appropriate sampling methodology depends on the
ecoregion, drainage area, stream size (wetted width), and diversity of habitat of the stream or
river to be assessed. Each sampling method includes a specific acceptable time period for
sampling. Method selection is presented visually as a flow chart in Figure 1. Criteria are further
described below.
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• Full Scale sites must meet the following criteria:
o Not in Triassic Basins ecoregion and drainage area
(DA) > 3 mi.2 or in Triassic Basins ecoregion and DA >
20 m i.2.
o The required diversity of habitat types are present.
o Sampleable during summer season (June 1— August
31), unless seasonal corrections will be made (see
Data analysis and reporting section).
• Qua14 sites must meet the following criteria:
o Not in Triassic Basins ecoregion and DA < 3mi.2 or in
Triassic Basins ecoregion and DA < 20 mi.2.
o Sampled during spring season (March 1— May 31).
Field measurements
Upon arrival at the site, prior to disturbing the stream bed, field
No
Site located in Triassic
Basins ecoregion?
No �Ye
Yes
Diverse
habitat
present?
Nv Yes
Qua14 Full
Scale
No
measurements should be collected. Figure 1 sampling method selection flow chart
1. All field measurements should be completed in accordance with Standard Operating Procedures
for Field Measurements and Observations (City of Durham 2020), including pre -sampling
calibration and post -sampling accuracy checks of field meters.
2. Field staff should measure the following water chemistry parameters in situ in the thalweg using
a multiparameter field meter: dissolved oxygen concentration (mg/L), dissolved oxygen
saturation (%), pH (S.U.), specific conductance (µS/cm at 25°C), and temperature (°C).
3. Turbidity (NTU) should be measured using a portable turbidimeter.
Record results on the habitat assessment field form.
Benthic macroinvertebrate sampling
1. Following the collection of field measurements, the stream reach to be sampled for benthic
macroinvertebrates should be determined. The stream should be viewed from a bridge or
similar vantage point to assess site conditions, flow status, available habitat, and a safe way to
access the stream. Sampling reaches should be located upstream of bridges and other stream
crossings to minimize the effects of localized impacts from the crossing or road (such as scour
pools) whenever possible. There should be appropriate stream flow within the sampling reach,
and it should include a variety of habitat types, such as riffles (shallow area of well -mixed, higher
velocity water), pools, undercut banks, sand deposits, and leaf packs. Stream conditions should
be assessed to determine if high flows, deep water, high turbidity, or other conditions exist that
present safety issues, habitat access issues, or concerns over recent scour. If such conditions
exist, sampling should be rescheduled.
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2. When entering and walking in the stream, avoid disturbing habitats that will likely be sampled,
such as riffles, aquatic vegetation (macrophytes), sand deposits, and leaf packs.
3. Collect the required number and types of samples. Each sampling method (Full Scale or Qua14)
requires a certain number of different sample types be collected (Table 1). When more than one
sample is required for a single sample type, field staff should target multiple locations in the
stream. For example, if two kick samples are needed, they should occur in two separate riffles,
preferably one with faster streamflow and one with slower streamflow. In cases when a certain
habitat is absent, the sample is omitted, and the sampling modification is noted in field notes.
Table 1 Descriptions of required samples for Full Scale and Qua14 sampling methodologies
Sample type
Habitat targeted
# samples for
Full Scale
# samples for
Qua14
Kick net
Riffle
2
1
Sweep net
Undercut banks, root mats, aquatic
vegetation (macrophyte) beds
3
1
Leaf pack
Decaying leaf packs (shredders)
1
1
Rock/log wash
Rocks/logs in non -riffle areas
2
None
Sand bag
Depositional areas
1
None
Visuals
All potential habitats
10 min./person
10 min./person
Sampling methods for each of the habitat types are described below.
a. Kick net samples: Riffles are areas in the stream that are shallow with faster -moving
water. The riffle(s) to be sampled will ideally have a good mix of larger substrate
(cobble, gravel) and an area of convergent flow. For Full Scale samples, the two samples
should be collected from areas of differing current speed (one in faster current, one in
slower current) if possible. In very small streams, or in sandy areas lacking riffles, kicks
can also be taken from root masses, snags, or other areas that create conditions similar
to a rocky riffle.
To begin a kick net sample, one field staff member stands at the bottom of the riffle
section to be sampled, facing upstream, with the net positioned upright and on the
stream bed where the current converges (Figure 2). The weighted bottom should be
adjusted to ensure no gaps exist between the bottom of the net and stream substrate,
which may require moving larger rocks/debris. Angle the net backward in the direction
of the streamflow to maximize the wetted surface area of the net while ensuring that
water does not flow over the top of the net. A second field staff member (the "kicker")
then stands approximately 1-1.5m upstream of the net and physically disturbs the
stream substrate with their feet and hands, scrubbing rocks and vegetation, as they
work their way towards the net. Disturbed and dislodged benthic organisms will flow
downstream and be caught in the net.
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Figure 2 Water Quality staff demonstrating the proper use of a kick net
Once the entire area upstream of the net has been disturbed, the kicker carefully lifts
the weighted bottom of the net and place it in a sieve bucket (Figure 3), ensuring that
specimens and detritus on the net are not lost or washed off. Then one person will hold
the net upright in the bucket while the other uses a plastic dish tub to collect stream
water to rinse specimens and debris off the net and into the bucket. Continue rinsing
until most specimens and debris have been washed into the bucket. Visually check the
net and remove any attached specimens with forceps. Large leaves and other debris in
the sample in the bucket can be elutriated by holding the bucket partially underwater,
with large debris shaken within the bucket to dislodge any specimens. Then the debris
should be discarded. This will make sample processing ("picking") easier and faster.
Once the sample has been rinsed into the sieve bucket and larger debris removed,
transfer the sample to a washtub for field processing. The kick net should be rinsed
thoroughly between sampling sites to prevent contamination between sites.
b. Sweep net sample: Sweep net samples are taken by using a long -handled D- or A -frame
net to physically disrupt an area and then vigorously sweep through the disturbed area.
Sweeps are usually taken from bank areas, including undercut banks and root mats, as
well as from beds of aquatic vegetation (Figure 4). Sweeps should generally begin
downstream and work upstream to reduce stirring up sediments which hinders the
sampler's view. Sweep nets should be thoroughly rinsed between sites to reduce
between -site contamination.
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Figure 3 Washing down the kick net into a sieve bucket to Figure 4 Water Quality staff member sampling submerged
remove the captured benthic organisms aquatic plants with a sweep net
c. Rock/log wash: Medium to large size rocks, sticks, and logs are rinsed in a large plastic
dish tub partially filled with water. Rocks are selected which have visible growths of
periphyton, Podostemum, or moss. A variety of woody material, from moderate -sized
branches to larger logs, should also be washed (Figure 5). A single composite sample can
be made from several (usually 5-10) rocks and/or logs. Habitat types sampled to create
this composite should represent their relative frequency within the stream. Any large
particulate material (leaves, etc.) in the plastic tub should be elutriated and discarded.
The material remaining in the tub is then poured through the fine mesh sampler
("midge -getter") and the water drained out completely (Figure 6). In case of organic
muck and mud clogging the sampler, tap the bottom of the mesh to help it drain. The
midge -getter should be well rinsed between sites.
Figure 5 Water Quality staff member demonstrating a
woody debris wash using a fine -mesh sampler
Figure 6 Rock -log wash sample being processed into a fine-
mesh sampler
The sample in the midge -getter is placed into a large lidded container that is half-filled
with 90% ethanol to kill and preserve collected organisms. The sample is allowed to sit
for several minutes, pulled out of the alcohol, and then backwashed into a picking tray
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for processing. This method of field preservation requires only a small amount of
alcohol, and it may be re -used several times.
d. Sand bag sample: Sandy areas are sampled with a large bag constructed of the same
fine mesh (300 µm) nitex netting material as the fine -mesh sampler ("midge -getter").
The bag can be employed similarly to a kick net, with the opening placed on the
streambed facing upstream, held in place with one hand and one foot, and the substrate
upstream of the bag manually disturbed with the free hand. The intent is not to shove
sediment into the bag, but rather to collect the suspended particles in the water column
when the substrate is disturbed. The material collected is then emptied into a large dish
tub half-filled with water. The "pour -and -preserve" elutriation technique is then used in
conjunction with a fine -mesh sampler ("midge -getter"): the sand and water mixture is
stirred by hand and the supernatant is poured off into the midge -getter (Figure 7, Figure
8). Once all the water in the dish tub has been decanted, the sample in the midge -getter
is placed in alcohol and processed similarly to the rock/log wash.
Figure 7 Water Quality staff member preparing to elutriate Figure 8 Water Quality staff member pouring elutriate from
a sand sample a sand sample into a fine -mesh sampler
e. Leaf pack: Leaf -packs and sticks are collected from multiple locations within the stream
and collected in the sieve bucket until it is approximately half full. The best leaf packs
consist of older leaves (not freshly fallen) that have begun to decay, but still retain some
structure. Piles of leaves in pool areas should not be collected. Leaf pack and small log
samples are particularly useful in large sandy rivers. Generally, three to four leaf packs
are collected from rocks or snags in fast current areas. Once the sieve bucket is
approximately half full, the more intact leaves are elutriated and discarded. The
remaining material is then transferred to a plastic dish tub to be held for field processing
("picking")
f. Visual search: Visual searches are intended to collect taxa from additional habitats that
are not well -represented by the other sampling methods used. Each field person should
complete approximately ten minutes of active searching. Visual inspection of large rocks
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and logs (the larger, the better) often adds to the species list. Large rocks and logs are
preferred microhabitat because of their stability during high flow events. Field staff
should look in a number of different areas, not just riffles. Rocks and logs in pools often
yield additional species, as this habitat is not well sampled by other methods. Using a
transect approach across the stream channel allows sampling of a variety of stream
habitats and flow regimes.
Experienced biologists may use visuals to target particular species that would be
expected to be at the site but have not been found during the standard sampling
methods. The tops and side of rocks are a specialized microhabitat for some caddisflies.
Decaying logs should be picked apart to look for midges (family Chironomidae) and
many taxa can be found under loose bark. Large woody debris are also a good place to
find Elmid beetles and snails. Rocks near the shore (in negligible current) will harbor
taxa such as Stenacron spp. and Pycnopsyche spp., and leaves near the shore may be the
primary habitat for certain snails. Certain caddisflies (Nyctiophylax spp. and related
genera) select crevices in rocks or logs, often along the edge, and cover them over with
silk strands. Microcaddisflies make small (2-4 mm) cases that can be found attached to
rocks and logs, usually on the top or along an edge.
A mussel search should be conducted if dead shells are evident along the shore, though
only live specimens should be added to the site's taxa list. Mussels can be found by
careful visual inspection of the stream bottom (Figure 9). During periods of receding
water levels, many species will move to deeper water, leaving a visible "track". The
bases of aquatic weeds (especially water willow) may contain many mussel species and
must be searched by hand. Mussels should be identified in the field and returned alive
to the stream. If sampling in an area with known populations of endangered or
threatened mussels, any live mussels should be photographed or sketched and returned
to the stream.
Figure 9 An example of an Elliptio complanata mussel
found in New Hope Creek.
Figure 10 Sampler conducting visual searches for aquatic
organisms
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Field processing of samples (sample "picking")
1. Place a small amount of sample in a shallow plastic tray with a small amount of stream water.
Evenly distribute the sample across the bottom of the tray.
2. Remove invertebrates from the sample using fine -tip forceps and place in 90% alcohol in small
screw-top vials (for smaller specimens) and one or more larger lidded wide -mouth jars or
containers (for large specimens). Not all invertebrates are necessarily collected. However, it is
often difficult for inexperienced field staff to identify unique taxa under field conditions so the
benthic biologist should provide guidance to less experienced staff during the picking process.
3. Certain taxa should not be removed from the area or preserved. Rather individuals are to be
field -identified by the benthic biologist and released after their presence is recorded. The
benthic biologist should provide guidance on which taxa should be handled this way, but this
generally this includes some taxa of crayfish and mussels.
4. Once picking of a tray is completed by less experienced staff, the tray should be reviewed by the
benthic biologist to ensure that no taxa were overlooked, with feedback provided to the less
experienced staff member.
S. Once picked and reviewed (if necessary), the remaining liquid and detritus are discarded. Repeat
steps 1-4 until all samples have been processed.
6. Unless otherwise specified in the study plan, specimens from all habitats and sample types can
be composited during picking and preservation, i.e., specimens from kick samples do not need
to be segregated in separate vials from specimens from sweep samples
7. When picking is complete, a label should be added to each specimen container that includes the
site number, date/time of sample, and initials of field staff. Use a pencil to prepare the label,
since most inks will dissolve in ethanol.
Habitat assessments and Stream Survey Collection Card
1. The form used for habitat assessments is provided in Appendix 1. This is the same form used by
the NCDEQ Biological Assessment Branch for Mountain/Piedmont sites (NCDEQ, 2016). The
website containing this form should be checked at least annually for updates to the form.
2. Guidance on completing the habitat assessment is provided on the form itself. However, the
assessor needs some background and/or training to understand the stream features of interest
and experience with the range of possible conditions associated with Piedmont streams. It is
recommended that the benthic biologist or another staff member with experience and training
in the method complete the habitat assessment. The form contains a page of general site
information followed by eight major categories of habitat assessments, each with associated
numerical score(s): extent of channel modifications; diversity of instream habitat; stream
substrate composition; pool variety; extent of riffle habitat; bank stability and riparian
vegetation; light penetration (riparian canopy cover); and riparian vegetative buffer width. The
assessments are primarily visual observations or estimates and procedures are provided on the
form itself. The maximum possible score is 100, which is associated with very high -quality
habitat.
The Stream Survey Collection Card is provided in Appendix 2 and is similar to the Collection Card
used by NCDEQ. The following information is recorded: site information (site ID, stream, river
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basin, COD watershed, ecoregion, drainage area), sampling information (personnel, date/time,
number of each type of sample collected, recent precipitation), habitat types present, evidence
of pollutants, substrate composition in pools and riffles, and substrate characterization (%
boulder, cobble, gravel, sand, silt, other). Substrate sizes are based on the Wentworth scale with
size measured along the intermediate axis of the particle (Table 2, Figure 11). The percentages
of each category of substrate within the reach should be visually estimated separately by two
different field staff and those estimates used to develop a consensus estimate for the reach.
They should total 100%.
Table 2 Wentworth scale for
classification of stream substrate
based on size.
Substrate
Size
Boulder
> 10 in.
Cobble
2.5 —10 in.
Gravel
0.08 — 2.5 in.
Sand
0.062 — 0.08 in.
Silt
< 0.062 in.
(A) Long axis
(B) Intermediate axis
q r.
(C) Short axis
The intermediate axis is the
8 pebble's diameter
C
Figure 11 Axes of sediment.
Digital photographs
1. Digital photographs should be taken at each field site to document conditions at the time of
sampling. At a minimum, one photograph should be taken facing upstream from the
downstream end of the main riffle sampled, and one photograph should be taken facing
downstream from the upstream end of this same riffle. Any notable conditions should be
documented with additional digital photographs.
Laboratory analysis and taxonomic identification
1. Analysis and taxonomic identifications of samples collected in the field are conducted by the
lead biologist in a laboratory due to the highly specialized skills required. This work may be
performed by a contract biologist. It is expected that the lead biologist or contract biologist will
follow the procedures and guidance outlined in the current NCDEQ SOP and taxonomic guides.
Current copies of the documents are available from the Biological Assessment Branch web page.
2. In general, the lead biologist or contract biologist will:
a. Identify all specimens to the lowest possible taxonomic level (usually genus or species),
which may require slide mounting and use of a compound microscope for certain taxa
(e.g., Oligochaeta, Chironomidae, and some Ephemeroptera structures)
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b. Record actual number of specimens for each taxon
c. Record relative abundance code for each taxon (see Table 3)
d. Composite ethanol -preserved specimens
e. Label all specimens (ethanol -preserved and slides)
Table 3 Relative abundance codes for laboratory analysis
of benthic macroinvertebrate samples.
Relative
Relative
abundance
Description
abundance
code
Rare
1
1-2 specimens
Common
3
3-9 specimens
Abundant
10
>_10 specimens
Data analysis and reporting
Taxonomic identifications and enumerations are used to calculate a number of metrics and an overall
bioclassification for each site. Roughly defined, the bioclassification is the weighted average of tolerance
values for all sampled species at a site. A summary of these metrics is provided in Table 4. Steps for
calculating these metrics are provided in this section.
Table 4 Description of metrics calculated from benthic macroinvertebrate community data.
Metric
Metric aliases
Description
S
Total taxa
Number of unique taxa (all Orders)
Total taxa richness
EPT S
Total EPT
Number of unique taxa in the Ephemeroptera, Plecoptera, and
EPT taxa richness
Trichoptera Orders only
N
Total organisms
Total number of individual specimens
Total abundance
EPT N
EPT abundance
Number of individual specimens in the Ephemeroptera,
Plecoptera, and Trichoptera Orders only. Used for "rounding
decisions" (if needed) for Full Scale samples.
BI
Biotic index
Numerical metric calculated using the relative abundance and
tolerance value for all taxa in a particular sample. Lower values
indicate less stressed communities.
EPT Score
EPT Diversity
Score derived from EPT S and reference table. Used to
determine Bioclassification for Full Scale samples
BI Score
Biotic Rating
Score derived from BI and reference table. Used to determine
Bioclassification for Full Scale samples
Bioclassification
Water Quality Rating
Qualitative rating based on other metrics. Exact calculation
method dependent on: ecoregion, drainage area, sampling
method, and season.
1. For each site, compile a table of taxa found and the relative abundance for each.
2. Determine the TV for each taxon and record in the digital workbook for tracking tolerance
values and calculating overall biotic index (BI) scores. TVs indicate the relative sensitivity of a
specific taxon to stressors (including water quality). Lower values (closer to 0) indicate less
tolerant taxa (i.e., more sensitive to stressors) and higher values indicate more tolerant taxa
(i.e., less sensitive to stressors). TVs are derived by benthic biologists and there is not necessarily
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agreement on the appropriate TV for all taxa. Because of this variability, it is important to
document the source of the TV. The City of Durham uses the current taxa list and associated TVs
that is published by the NCDEQ, Biological Assessment Branch as part of their SOP (NCDEQ
2016). This list was last updated in April 2010. The version that is current at the time of writing is
provided in Appendix 4, but the NCDEQ Biological Assessment Branch's web site should be
checked regularly (at least annually) to determine if an updated list has been issued.
3. Calculate the bioclassification for Qua14 samples. These calculations are based on the "Small
Streams" criteria developed by NCDEQ and detailed in their SOP (NCDEQ 2016).
a. For each taxon, the relative abundance value assigned (1, 3, or 10) is multiplied by the
current tolerance value (TV). These weighted TVs from all taxa are then summed and
divided by the sum of all relative abundance values to get an "average" tolerance,
referred to as the Biotic Index (BI). The BI calculation can also be described
mathematically as:
n 1 n 1
Biotic index (BI) _ I TV; x Ni I - (Y Ni I
(Equation 1) Where:
TV; = Tolerance value for taxon i
Ni = Relative abundance code for taxon i
n = Total number of unique taxa
b. Use Table 5 and the calculated BI to determine the final Bioclassification for each site.
Table 5 Biotic index (BI) thresholds for determining Bioclassification of
Oual4 samples
BI
Bioclassification
< 4.31
Excellent
4.31— 5.18
Good
5.19 — 5.85
Good -Fair
5.86-6.91
Fair
> 6.91
Poor
4. Calculate the Bioclassification for Full Scale samples. This is based on the methods used for Full
Scale samples by NCDEQ and detailed in their SOP (NCDEQ 2016). An overview of the process is
shown in Figure 12.
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Site sampled between
Jun. 1— Sep. 30?
No Yes
Removewinter I I Calculate
stoneflies from II .II EPTSand get
taxa list EPT S Score
Site sampled between Calculate
Jun. 1 — Sep. 30? BI and get
BI Score
No Yes
Add seasonal Average EPT S Score
correction to BI and BI Score.
Score Round result
Assign bioclassification:
5 = Excellent
4 = Good
3 =Good -Fair
2 = Fair
1 = Poor
Figure 12 Flow chart for calculation of Bioclassifications for Full Scale samples.
a. If the sample was collected between June 1— September 30, no corrections for winter
stoneflies are needed. If the sample was collected between October 1— May 31, then
winter stoneflies (Order Plecoptera) must be removed from the taxa list before
continuing with calculations. The list of winter stonefly taxa that have been previously
found in Durham County is provided in Appendix 3. A full statewide list of winter
stonefly taxa can be found in the NCDEQ Biological Assessment Branch SOP (NCDEQ
2016). The NCDEQ Biological Assessment Branch's web site should be checked regularly
(at least annually) to determine if an updated list has been issued.
b. Calculate EPT taxa richness (EPT S), excluding winter stoneflies (if applicable), by
counting the total number of unique taxa in the orders Ephemeroptera, Plecoptera, and
Trichoptera. Find the EPT S in the left column of Table 6 and record the associated Score
found in the middle column. This is the EPT Score.
c. Calculate BI for the sample (excluding winter stoneflies, if applicable), using Equation 1
(see Qua14 section above). Refer to Table 7 to determine the seasonal correction to be
added to the BI.
d. Find the seasonally corrected BI in the right column of Table 6 and record the
corresponding Score in the middle column. This is the BI Score.
e. Calculate the average of the EPT S Score and the BI Score. If the average is 1.5, 2.5, 3.5,
or 4.5, then use the EPT N (number individual EPT specimens in the sample) and Table 8
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to determine the direction of rounding. For all other average values, round to the
nearest integer.
f. Use the rounded average and Table 9 to assign the final Bioclassification.
Table 6 Look -up table for Scores associated with EPT S and BI.
EPT S
Score
BI value
> 33
5.0
< 5.14
32-33
4.6
5.14 - 5.18
30-31
4.4
5.19 - 5.23
26-29
4.0
5.24 - 5.73
24-25
3.6
5.74 - 5.78
22-23
3.4
5.79 - 5.83
18-21
3.0
5.84 - 6.43
16-17
2.6
6.44 - 6.48
14-15
2.4
6.49 - 6.53
10-13
2.0
6.54 - 7.43
8-9
1.6
7.44 - 7.48
6-7
1.4
7.49 - 7.53
0-5
1.0
> 7.53
Table 7 Seasonal corrections for BI for Full Scale samples.
Sample date
Season
BI adjustment
June 1- September 30
Summer
None
October 1- November 30
Fall
+0.1
December 1- February 28
Winter
+0.1
March 1- May 31
Spring
+0.2
Table 8 EPT N criteria for rounding decisions using Full Scale criteria.
Average of
BI Score and EPT S Score
EPT N
Round to...
4.5
>_ 135
5
4.5
<_ 134
4
3.5
>_ 125
4
3.5
<_ 124
3
2.5
>_ 85
3
2.5
<_ 84
2
1.5
>_ 45
2
1.5
<_ 44
1
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Table 9 Assignment of final Bioclassification for Full Scale samples.
Rounded average of
BI Score and
EPT S Score
Final bioclassification
5
Excellent
4
Good
3
Good -Fair
2
Fair
1
Poor
Data and records management
Currently, results are summarized in Excel workbooks. The following information should be recorded for
each sample collected:
• Site information (site number, stream, location, City of Durham watershed, drainage
area, river basin, ecoregion)
• Date of sampling
• Names of field staff
• Name of biologist that performed specimen identifications and analysis
• Sampling methodology used
• Habitat subscores and total score
• Qualitative assessments recorded on the Habitat Assessment and "Collection Card" field
forms
• All metrics described in Table 4.
• Taxa list, relative abundance, and tolerance values used for analysis
Excel files and digital site photos are stored in the appropriate location in the Water Quality server
folder.
The preserved specimens from each sample (either in ethanol or mounted on microscope slides) will be
archived by WQU in case questions arise in the future over specific identifications. Hard copy habitat
assessment and Stream Sample Collection Card forms will be scanned and stored on the internal WQU
server and original hard copies stored in the WQU offices. All samples and paper copies of forms will be
stored for a minimum of ten years. Digital copies will be retained indefinitely. Older samples may be
discarded in accordance with the SDS. Qualified data will be also be stored in the Benthic Sampling
Access Database for future use. This database is capable of generating numerous reports in support of
the water quality program and will be maintained by a member of the WQU staff.
Quality assurance and quality control (QA/QC)
QA/QC procedures for benthic macroinvertebrate sampling, identification, and analysis are the
responsibility of the lead biologist or contract biologist whose services are secured to conduct sampling
described in the methods section. If a contractor is being used, during the competitive contracting
process, potential contractors are screened to determine if they comply with NCDEQ procedures,
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guidance, and requirements. It should be confirmed that the contractor conducts work described in this
SOP using NCDEQ's most current taxa list and associated tolerance values.
New or inexperienced field staff must be supervised by experienced personnel (either contract staff or
other City staff) during field activities. Additional oversight of inexperienced staff includes close
supervision during sampling, a quality control examination of their picking trays before discarding, and
mentoring during visual samples.
Data and results received from contractors will be reviewed by WQU staff for completeness and to
ensure that taxa appear to have been identified to the appropriate taxonomic level, appropriate TVs
were used, and appropriate calculations (including seasonal corrections) were applied.
QA/QC procedures for other activities associated with benthic monitoring (such as field water quality
measurements) should comply with the appropriate City of Durham Stormwater and GIS Services SOP.
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References
City of Durham. 2011. Benthic Monitoring Split Sampling Study (Technical Report). Department of Public
Works, Stormwater & GIS Services, Water Quality Section. Durham, North Carolina.
City of Durham. 2017. Safety Operating Procedures. Department of Public Works. Durham, North
Carolina.
City of Durham. 2019. Standard Operating Procedures for Field Meter Calibration and Maintenance.
Department of Public Works, Stormwater & GIS Services, Water Quality Unit. Durham, North Carolina.
City of Durham. 2020. Standard Operating Procedures for Field Measurements and Observations.
Department of Public Works, Stormwater & GIS Services, Water Quality Unit. Durham, North Carolina.
Griffith, G.E., Omernik, J.M., Comstock, J.A., Schafale, M.P., McNab, W.H., Lenat, D.R., MacPherson, T.F.,
Glover, J.B., and Shelburne, V.B., 2002, Ecoregions of North Carolina and South Carolina, (color poster
with map, descriptive text, summary tables, and photographs). Reston, Virginia, U.S. Geological Survey
(map scale 1:1,500,000).
Lenat, D.R. 1993. A Biotic Index for the Southeastern United States: Derivation and List of Tolerance
Values, with Criteria for Assigning Water Quality Ratings. Journal of the North American Benthological
Society 12:279-290
Merritt, R.W, Cummins, K.W., Berg, M.B. 2008. An Introduction to the Aquatic Insects of North America
(Fourth Edition). Kendall Hunt Publishing Company. Dubuque, IA.
NC Department of Natural and Cultural Resources. North Carolina Natural Heritage Data Explorer.
Natural Heritage Program. https://ncnhde.natureserve.org. Accessed March 6, 2017.
NC Department of Environmental Quality (NCDEQ). 2016. Standard Operating Procedures for the
Collection and Analysis of Benthic Macroinvertebrates. Division of Water Resources. Raleigh, North
Carolina.
Voshell, J.R. Guide to Common Freshwater Invertebrates of North America. 2002. McDonald &
Woodward Publishing Company. Granville, Ohio.
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Appendix 1. Habitat Assessment Field Form
The Water Quality Unit used the Habitat Assessment form developed by the NC Department of Environmental Quality's
Biological Assessment Branch, which is available from their website. The current version is provided on the following pages.
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11/13 Revision 8
Habitat Assessment Field Data Sheet
Mountain/ Piedmont Streams
Biological Assessment Branch, DWR
OTALSCORE
Directions for use: The observer is to survey a minimum of 100 meters with 200 meters preferred of stream, preferably in an
upstream direction starting above the bridge pool and the road right-of-way. The segment which is assessed should represent average
stream conditions. To perform a proper habitat evaluation the observer needs to get into the stream. To complete the form, select the
description which best fits the observed habitats and then circle the score. If the observed habitat falls in between two descriptions, select
an intermediate score. A final habitat score is determined by adding the results from the different metrics.
Stream
Date CC#
Location/road:
Observer(s) Type of Study: ❑ Fish
Latitude Longitude
Water Quality: Temperature °C DO
Basin
(Road Name )County,
basin
❑Benthos ❑ Basinwide ❑Special Study (Describe)
Ecoregion: ❑ MT ❑ P ❑ Slate Belt ❑ Triassic Basin
mg/l Conductivity (corr.) µS/cm pH
Physical Characterization: Visible land use refers to immediate area that you can see from sampling location - include what
you estimate driving thru the watershed in watershed land use.
Visible Land Use: %Forest %Residential %Active Pasture % Active Crops
%Fallow Fields % Commercial %Industrial %Other - Describe:
Watershed land use : ❑Forest ❑Agriculture ❑Urban ❑ Animal operations upstream
Width: (meters) Stream Channel (at top of bank) Stream Depth: (m) Avg Max
❑ Width variable ❑ Large river >25m wide
Bank Height (from deepest part of riffle to top of bank -first flat surface you stand on): (m)
Bank Angle: ° or ❑ NA (Vertical is 90°, horizontal is 00. Angles > 90' indicate slope is towards mid -channel, < 90'
indicate slope is away from channel. NA if bank is too low for bank angle to matter.)
❑ Channelized Ditch
❑Deeply incised -steep, straight banks ❑Both banks undercutat bend ❑Channel filled in with sediment
❑ Recent overbank deposits ❑Bar development ❑Buried structures ❑Exposed bedrock
❑ Excessive periphyton growth ❑ Heavy filamentous algae growth ❑Green tinge ❑ Sewage smell
Manmade Stabilization: ON ❑Y: ❑Rip -rap, cement, gabions ❑ Sediment/grade-control structure ❑Berm/levee
Flow conditions: ❑High ❑Normal ❑Low
Turbidity: ❑Clear ❑ Slightly Turbid ❑Turbid ❑Tannic ❑Milky ❑Colored (from dyes)
Good potential for Wetlands Restoration Project?? ❑ YES ONO Details
Channel Flow Status
Useful especially under abnormal or low flow conditions.
A. Water reaches base of both lower banks, minimal channel substrate exposed ............................ ❑
B. Water fills >75% of available channel, or <25% of channel substrate is exposed ........................ ❑
C. Water fills 25-75% of available channel, many logs/snags exposed ............................................. ❑
D. Root mats out of water................................................................................................................... ❑
E. Very little water in channel, mostly present as standing pools ..................................................... ❑
Weather Conditions: Photos: ON ❑Y ❑ Digital 035mm
Remarks:
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1. Channel Modification Score
A. channel natural, frequent bends................................................................................. 5
B. channel natural, infrequent bends (channelization could be old) ........................................................ 4
C. some channelization present............................................................................................................... 3
D. more extensive channelization, > 40% of stream disrupted................................................................ 2
E. no bends, completely channelized or riprapped or gabioned, etc ....................................................... 0
❑ Evidence of dredging ❑Evidence of desnagging=no large woody debris in stream ❑Banks of uniform shape/height
Remarks Subtotal
II. Instream Habitat: Consider the percentage of the reach that is favorable for benthos colonization or fish cover. If >70% of the reach
is rocks, 1 type is present, circle the score of 17. Definition: leafpacks consist of older leaves that are packed together and have begun
to decay (not piles of leaves in pool areas). Mark as Rare, Common, or Abundant.
Rocks Macrophytes Sticks and leafpacks Snags and logs Undercut banks or root mats
AMOUNT OF REACH FAVORABLE FOR COLONIZATION OR COVER
>70% 40-70% 20-40% <20%
Score Score Score Score
4 or 5 types present ................. 20 16 12 8
3 types present ......................... 19 15 11 7
2 types present ......................... 18 14 10 6
1 type present ........................... 17 13 9 5
No types present .............................0
❑ No woody vegetation in riparian zone Remarks Subtotal
III. Bottom Substrate (silt, sand, detritus, gravel, cobble, boulder) Look at entire reach for substrate scoring, but only look at riffle
for embeddedness, and use rocks from all parts of riffle -look for "mud line" or difficulty extracting rocks.
A. substrate with good mix of gravel, cobble and boulders Score
1. embeddedness <20% (very little sand, usually only behind largeboulders)......................... 15
2. embeddedness 20-40%.......................................................................................................... 12
3. embeddedness 40-80%.......................................................................................................... 8
4. embeddedness>80%............................................................................................................. 3
B. substrate gravel and cobble
1. embeddedness<20%............................................................................................................ 14
2. embeddedness 20-40%......................................................................................................... 11
3. embeddedness 40-80%........................................................................................................ 6
4. embeddedness>80%............................................................................................................ 2
C. substrate mostly gravel
1. embeddedness<50%............................................................................................................ 8
2. embeddedness>50%............................................................................................................ 4
D. substrate homogeneous
1. substrate nearly all bedrock...................................................................................................
2. substrate nearly all sand........................................................................................................
3. substrate nearly all detritus....................................................................................................
4. substrate nearly all silt/ clay...................................................................................................
IV. Pool Variety Pools are areas of deeper than average maximum depths with little or no surface turbulence. Water velocities
associated with pools are always slow. Pools may take the form of "pocket water", small pools behind boulders or obstructions, in large
high gradient streams, or side eddies.
A. Pools present Score
1. Pools Frequent (>30% of 200m area surveyed)
a. variety of pool sizes............................................................................................................... 10
b. pools about the same size (indicates pools filling in)............................................................ 8
2. Pools Infrequent (<30% of the 200m area surveyed)
a. variety of pool sizes............................................................................................................... 6
b. pools about the same size...................................................................................................... 4
B. Pools absent............................................................................................................................................ 0
Subtotal
❑ Pool bottom boulder-cobble=hard ❑ Bottom sandy -sink as you walk ❑ Silt bottom ❑ Some pools over wader depth
Remarks
Page Total
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 29 of 50
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V. Riffle Habitats
Definition: Riffle is area of reaeration-can be debris dam, or narrow channel area. Riffles Frequent Riffles Infrequent
Score Score
A. well defined riffle and run, riffle as wide as stream and extends 2X width of stream.... 16 12
B. riffle as wide as stream but riffle length is not 2X stream width .................................... 14 7
C. riffle not as wide as stream and riffle length is not 2X stream width ............................. 10 3
D. riffles absent.....................................................................................................................0
Channel Slope: ❑Typical for area ❑Steep=fast flow ❑Low=like acoastal stream Subtotal
VI. Bank Stability and Vegetation
A. Erosion
1. No, or very little, erosion present............................................7
2. Erosion mostly at outside of meanders...................................6
3. Less than 50% of banks eroding.............................................3
4. Massive erosion......................................................................0
Erosion Score
B. Bank Vegetation
1. Mostly mature trees (>12" DBH) present ..............................7
2. Mostly small trees (<12" DBH) present, large trees rare .......5
3. No trees on bank, can have some shrubs and grasses.............3
4. Mostly grasses or mosses on bank.........................................2
5. Little or no bank vegetation, bare soil everywhere ................
0 Vegetation Score
Remarks Subtotal,
VII. Light Penetration Canopy is defined as tree or vegetative cover directly above the stream's surface. Canopy
would block out sunlight when the sun is directly overhead. Note shading from mountains, but not use to score this metric.
Score
A. Stream with good canopy with some breaks for light penetration ............................................. 10
B. Stream with full canopy - breaks for light penetration absent ..................................................... 8
C. Stream with partial canopy - sunlight and shading are essentially equal .................................... 7
D. Stream with minimal canopy - full sun in all but a few areas ....................................................... 2
E. No canopy and no shading............................................................................................................. 0
Remarks Subtotal
VIII. Riparian Vegetative Zone Width
Definition: Riparian zone for this form is area of natural vegetation adjacent to stream (can go beyond floodplain). Definition: A
break in the riparian zone is any place on the stream banks which allows sediment or pollutants to directly enter the stream, such
as paths down to stream, storm drains, uprooted trees, otter slides, etc.
FACE UPSTREAM Lft. Bank Rt. Bank
Dominant vegetation: ❑ Trees ❑ Shrubs ❑ Grasses ❑ Weeds/old field ❑Exotics (kudzu, etc) Score Score
A. Riparian zone intact (no breaks)
1. width > 18 meters..................................................................................... 5 5
2. width 12-18 meters................................................................................... 4 4
3. width 6-12 meters..................................................................................... 3 3
4. width < 6 meters...................................................................................... 2 2
B. Riparian zone not intact (breaks)
1. breaks rare
a. width > 18 meters.........................................................................
4
4
b. width 12-18 meters.......................................................................
3
3
c. width 6-12 meters.......................................................................
2
2
d. width < 6 meters.........................................................................
1
1
2. breaks common
a. width > 18 meters.........................................................................
3
3
b. width 12-18 meters......................................................................
2
2
c. width 6-12 meters.......................................................................
1
1
d. width < 6 meters.........................................................................
0
0
Remarks
Subtotal
Page Total
❑ Disclaimer -form filled out, but score doesn't match subjective opinion -atypical stream.
TOTAL SCORE
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 30 of 50
8/13/2021

Supplement for Habitat Assessment Field Data Sheet
Diagram to determine bank angle:
'Ll
900 450 1350
Typical Stream Cross-section
Normal High Water
Upper Bank
— Lower
Bank
Stream Width This side is 45' bank angle.
Site Sketch:
Other comments:
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 31 of 50
8/13/2021

Appendix 2. Collection Card Field Form
The current Benthos/Stream Survey Data Sheet (also known as the Collection Card Field Form) is
provided on the following page.
Benthic Macroinvertebrate Monitoring Collection Card
City of Durham, Water Quality Unit
Site ID: Stream: Location:
River basin: CPF / NEU
Outer Piedmont
Date:
Drainage area (sq. miles):
Other
Watershed:
Ecoregion: Triassic Basins / Slate Belt / N.
Time (start/end): / Field staff:
Sampling method: Qua14 / Full Scale /
Instream Habitats: (Check all that are present)
# of Samples:
Pools ❑ Backwaters
❑
Kicks:
Riffles ❑ Detritus
❑
Sweeps:
Snags ❑ Aquatic Weeds
❑
Leaf Packs:
Undercut Banks ❑ Podostemum
❑
Visuals:
Root Mats ❑ Other
❑
Pollutant indicators: (Check all that apply and describe)
Substrate
❑ Odor:
Boulder %
Cobble
❑ Sheen:
Gravel
❑ Floatables:
Sand
❑ Sediment in riffles, depth (in.):
Silt/clay %
Other
Field Observations:
Rock/Log:
Sand:
Other:
Obs 1 Obs 2 Final
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 32 of 50
8/13/2021

Appendix 3. Taxa list for seasonal correction of Full Scale samples
The table below provides the list of winter stonefly taxa previously found in Durham City and County
that should be used for seasonal corrections of Full Scale samples collected in Fall, Winter, or Spring
(October 1— May 31). The list is an edited version of Table 4 in the 2016 NCDEQ Benthic
Macroinvertebrate monitoring SOP and only includes taxa that have been historically recorded by the
City of Durham's previous benthic monitoring. Refer to NCDEQ, 2016 for the full list.
Family
Species
Nemouridae
Amphinemura spp.
Prostoia s
Taeniopterygidae
Strophopteryx spp
Taeniopteryx burksi
Taenio to x SPP
Perlodidae
Clioperla clio
Cultus decisus complex
Cultus spp
Isoperla holochlora-dark form
Iso erla nr holochlora,
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 33 of 50
8/13/2021

Appendix 4. NCDEQ Taxa List and Tolerance Values (TVs)
The following taxa list and corresponding tolerance values (TVs) was obtained from the NCDEQ SOP for
the Collection and Analysis of Benthic Macroinvertebrates, version 5.0 (February 2016). The NCDEQ
website or Biological Assessment Branch staff should be consulted regularly to determine if an updated
list is available. Note that this is a statewide list; most of these taxa are not found in Durham streams.
Order
Latin Name
Tolerance Value
Ephemeroptera
AMELETIDAE
Arneletus lineatus
2.4
BAETIDAE
Acentrella alachua
3.0
centrella nodineae
1.9
centrella parvula
4.8
centrella spp
2.5
centrella turbida gr
2.0
cer enna pygmaea
3.7
Baetis lavistri a
6.8
Baetis intercalaris
5.0
Baetis pluto
3.4
Baetis tricaudatus
1.5
Callibaetis spp
9.2
Centro tilum spp
3.8
Cloeon spp
7.3
Di hetorha eni
1.1
Heterocloeon amplum
3.4
Heterocloeon curiosum
2.1
Heterocloeon spp
3.7
Iswaeon anoka
4.4
Labiobaetis ephippiatus
3.5
Labiobaetis rondalis
4.6
Labiobaetis propinquus
5.8
Paracloeodes spp
8.0
Plauditus cestus
4.6
Plauditus dubius gr
2.2
Procloeon spp
1.9
BAETISCIDAE
Baetisca berneri
1.4
Baetisca carolina
4.2
Baetisca spp
3.2
CAENIDAE
Brach cercus spp
2.1
Caenis spp
6.8
EPHEMERELLIDAE
Attenella attenuata
1.1
Dannella simplex
3.4
Drunella alle heniensis
0.3
Drunella conestee
0.0
Drunella cornutella
0.0
Drunella lata
0.0
Drunella tuberculata
0.0
Drunella walkeri
0.6
Drunella wa ah
0.0
E hemerella catawba
0.0
E hemerella catawba/dorothea
4.0
E hemerella dorothea
3.3
E hemerella his ida
0.1
E hemerella invaria gr
2.6
E hemerella rossi gr
10.0
E hemerella rotunda
11.8
E hemerella spp
12.1
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 34 of 50
8/13/2021

Order
Latin Name
Tolerance Value
Eur to hella bicolor
4.8
Eur to hella doris
7.0
Eur to hello funeralis
2.5
Eur to hella spp
4.0
Eur to hella tem oralis gr
4.8
Eur to hella verisimilis
3.9
Penelomax se tentrionalis
2.1
erratella carolina
0.0
erratella serrata
1.4
erratella serratoides
1.7
Telo ano sis de iciens
2.6
EPHEMERIDAE
Ephemera blanda
2.4
Ephemera guttalata
0.0
Ephemera spp
2.0
Hexa enia spp
4.4
HEPTAGENIIDAE
Cinygmula subae ualis
0.0
E eorus dis ar
1.0
E eorus pleuralis
1.5
Epeorusspp
1.6
E eorus vitreus
1.2
He to enia mar inalis gr
2.2
He to enia pulla
2.2
He to enia spp
1.9
Leucrocuta a hrodite
2.9
Leucrocuta spp
2.0
Maccoffertium carlsoni
2.1
Maccoffertium exi uum
3.8
Maccoffertium ithaca
3.0
Maccaffertium lenati
2.5
Maccoffertium medio unctatum
4.2
Maccoffertium meririvulanum
0.5
Maccoffertium mexicanum
4.7
Maccoffertium modestum
5.7
Maccoffertium pudicum
2.1
Maccoffertium terminatum
4.4
Maccoffertium vicarium
1.5
Rhithro ena exilis
0.0
Rhithro ena spp
0.0
Rhithro ena uhari
0.0
tenacron carolina
1.3
tenacron inter unctatum
6.4
tenacron pallidurn
2.8
tenonema emoratum
6.9
ISONYCHIIDAE
Ison chia spp
3.6
LEPTOHYPHIDAE
Tricor thodes spp
5.0
LEPTOPHLEBIIDAE
Habro hlebia vibrans
0.3
Le to hlebia spp
6.0
Parale to hlebia spp
1.2
NEOEPHEMERIDAE
Neoe hemera purpurea
1.5
POLYMITARCYIDAE
E horon leukon
1.5
POTAMANTHIDAE
ntho otamus distinctus
1.6
ntho otamus spp
1.5
SIPHLONURIDAE
Siphlonuruss
6.0
Pleco tera
CAPNIIDAE
Allocapnias
3.3
CHLOROPERLIDAE
Alloperla spp
1.0
Ha to erla brevis
1.4
uwallia mar inata
12.6
weltsa 5pp
10.2
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 35 of 50
8/13/2021

Order
Latin Name
Tolerance Value
LEUCTRIDAE
Leuctra spp
1.5
NEMOURIDAE
Arnphinernura spp
3.8
Prostoia spp
5.2
PELTOPERLIDAE
Talla erla spp
1.3
PERLIDAE
Acroneuria abnormis
2.1
croneuria arenosa
2.4
croneuria carolinensis
1.2
croneuria evoluta
1.7
croneuria 1 corias
2.1
netina spp
1.1
Beloneuria spp
0.0
Ecco tura xanthenes
4.7
Neo erla spp
2.1
Para netina umosa
3.6
Para netina ichusa/media
0.2
Para netina immar inata
1.1
Para netina kansensis
1.9
Perlesta spp
2.9
Perlinella dr mo
1.3
PERLODIDAE
Clio erla clio
5.2
Cultus decisus complex
1.5
Di to erla du licata
2.8
Helo icus subvarians
1.2
Iso erla davisi/nr transmarina
4.8
Iso erla holochlora-darkformlcf powhatan
1.2
Iso erla holochlora-li ht form
0.7
Iso erla kirchneri complex
2.5
Iso erla nr holochlora
0.0
Iso erla orata
0.0
Iso erla poffiln s -Collins Cr
5.2
Iso erla similis/ seudosimilis gr
0.8
Iso erla spp
3.2
Malirekus hastatus
1.0
Remenus spp
0.9
PTERONARCYIDAE
Pteronarc s biloba
0.0
Pteronarc s dorsata
2.4
Pteronarc s proteus
0.4
Pteronarc s spp
1.8
AENIOPTERYGIDAE
Strophopteryxs
3.3
Taenio ter x burksi
6.6
Taenio ter x spp
6.0
richo tera
APATANIIDAE
atania spp
0.6
BRACHYCENTRIDAE
Brach centrus appalachia
1.0
Brach centruslateralis
1.9
Brach centrus ni rosoma
3.1
Brach centrus numerosus
1.7
Brach centrus s inae
0.0
Brach centrus spp
2.2
Micrasema bennetti
0.0
Micrasema charonis
1.0
Micrasema rickeri
0.0
Micrasema wata a
2.2
CALAMOCERATIDAE
Anisocentropus pyraloides
1.3
Hetero lectron americanum
2.0
DIPSEUDOPSIDAE
Ph loc ntropusspp
4.8
GLOSSOSOMATIDAE
0.0
_Agapetusspp
Glossosoma spp
11.4
Proto tila spp
12.3
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 36 of 50
8/13/2021

Order
Latin Name
Tolerance Value
GOERIDAE
Goera calcarata
1.0
Goera spp
0.7
HELICOPSYCHIDAE
Helico s the borealis
0.0
HYDROPSYCHIDAE
rcto s the irrorata
0.0
Cheumato s the spp
6.6
Di lectrona modesta
2.3
H dro s the C. alhedra
0.0
H dro s the C. bronta
2.3
H dro s the C. macleodi
0.7
H dro s the C. morosa
2.3
H dro s the C. slossonae
0.0
H dro s the C. s arna
2.5
H dro s the H. betteni
7.9
H dro s the H. decalda
3.2
H dro s the H. demora
2.6
H dro s the H. incommoda
4.6
H dro s the H. holerata
3.7
H dro s the H. rossi
4.8
H dro s the H. scalaris
2.6
H dro s the H. venularis
5.1
Macrostemum spp
3.4
Para s the cardis
0.0
HYDROPTILIDAE
H dro tila spp
6.5
Leucotrichia pictipes
4.6
LEPIDOSTOMATIDAE
Le idostorna spp
1.0
LEPTOCERIDAE
Ceraclea anc lus
2.8
Ceraclea maculata
6.2
Ceraclea spp
2.2
Ceraclea transversa
2.8
M stacides se ulchralis
2.6
Necto s the candida
6.5
Necto s the ex uisita
4.3
Necto s the pavida
3.9
Oecetis georgia
3.6
Oecetis nocturna
5.0
Oecetis persimilis
4.6
Oecetis Scala gr
2.7
Oecetis 5pp
5.1
etodes spp
0.0
Triaenodes i nitus
4.8
Triaenodes in'ustus
2.7
Triaenodes pernalhelo
3.8
Triaenodes spp
4.1
LIMNEPHILIDAE
H dato h laxar us
2.4
Irono uia punctatissima
6.7
P cno s the gentilis
1.8
P cno s the guttifer
2.2
P cno s the le ida gr
3.9
P cno s the scabri ennis
2.5
P cno s the spp
2.5
MOLANNIDAE
Molanna blenda
1.6
Molanna tryphena
2.4
ODONTOCERIDAE
Psilotreta spp
0.5
PHILOPOTAMIDAE
Chimarra spp
3.3
Dolo hilodes spp
1.0
Wormaldia spp
12.4
PHRYGANEIDAE
10ligostomis pardalis
16.2
Ptilostomis 5pp
15.9
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 37 of 50
8/13/2021

Order
Latin Name
Tolerance Value
POLYCENTROPODIDAE
C rnellus raternus
6.8
Neurecli sis spp
4.0
N ctio h lax celta
0.7
N ctio h lax moestus
3.8
N ctio h lax ne ho hilus
0.6
N ctio h lax spp
0.8
Pol centro us sensu lato spp
3.1
PSYCHOMYIIDAE
Lype diversa
3.9
Ps chom is lavida
3.0
Ps chom is nomada
2.0
RHYACOPHILIDAE
Rh aco hila acutiloba
0.0
Rh aco hila appalachialnigrita
0.0
Rh aco hila atrata
0.0
Rh aco hila carolina
0.4
Rh aco hila fenestra/ledra
4.6
Rh aco hila ormosa
0.1
Rh aco hila uscula
1.6
Rh aco hila torva
1.5
SERICOSTOMATIDAE
Fatti is ele
0.0
UENOIDAE
Neo h laxconsimilis
0.3
Neo h lax uscus
0.0
Neo h lax mitchelli
0.0
Neo h lax oli ius
2.4
Neo h lax ornatus
1.3
Neo h lax spp
1.6
Odonata
AESHNIDAE
Basiaeschna 'anato
7.1
Bo eria graflana
3.8
Bo eria vinosa
5.8
Nasiaeschna pentacantha
6.6
CALOPTERYGIDAE
Calo ter x spp
7.5
Hetaerina spp
4.9
COENAGRIONIDAE
r is spp
8.3
Enalla ma spp
8.5
Ischnura spp
9.5
CORDULEGASTRIDAE
Cordule aster spp
5.7
CORDULIIDAE
E itheca princeps
7.3
E itheca spp
8.0
Helocordulia spp
5.8
Neurocordulia obsoleta
5.3
Neurocordulia spp
5.3
Neurocordulia vir iniensis
1.1
omatochlora spp
8.9
GOMPHIDAE
Dromo om hus spp
5.6
Gom hus s inice s
6.1
Gom hus spp
5.9
Hagenius brevist lus
4.4
Lanthus parvulus
0.6
Lanthus spp
1.6
Lanthus vernalis
0.8
0 hio om hus spp
5.9
Pro om hus spp
8.2
t to om hus albist lus/si mast lus
5.0
LIBELLULIDAE
Libellula spp
9.4
Pach di lax lon i ennis
9.6
Perithemis spp
9.4
Plathemis 1 dia
19.8
MACROMIIDAE
IMacrornia spp
16.2
Hemi tera
BELOSTOMATIDAE
lBelostoma spp
19.5
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 38 of 50
8/13/2021

Order
Latin Name
Tolerance Value
CORIXIDAE
Sigaras
8.7
NEPIDAE
Ranatra spp
6.3
Me alo tera
CORYDALIDAE
Cor dalus cornutus
5.2
Ni ronia fasciatus
6.1
Ni ronia serricoml.s
4.6
SIALI DAE
sialis spp
7.0
Coleo tera
DRYOPIDAE
Helichus basalis
0.5
Helichus litho hilus
3.0
Helichus spp
4.1
DYTISCIDAE
Co totornus spp
8.5
H dro orus spp
7.0
Lacco hilus spp
9.8
Lio oreus spp
4.0
Neo orus mellitus
3.9
Neo orus spp
5.0
tictotarsus griseostriatus
4.9
ELMIDAE
nc ron x varie atus
6.8
Dubira hia spp
5.5
Dubira hia vittata
5.0
Macron chus glabratus
4.7
Microc lloe us pusillus
3.3
0 tioservus ovalis
2.1
0 tioservus spp
2.1
Oulimnius latiusculus
1.9
Promoresia ele ans
2.1
Promoresia spp
3.1
Promoresia tardella
0.0
tenelmis crenata
7.8
tenelmis spp
5.6
GYRINIDAE
Dineutus spp
5.0
6 rinus spp
5.8
HALIPLIDAE
Peltod tes spp
8.4
HYDROPHILIDAE
eerosus spp
8.8
Enochrusspp
8.5
Laccobius spp
6.5
ercho sis tessellatus
4.4
Tro isternus spp
9.3
PSEPHENIDAE
Ecto ria nervosa
4.3
Pse henus herricki
2.3
PTILODACTYLIDAE
Anchytarsus bicolor
2.4
Di tera
BLEPHARICERIDAE
ele haricera spp
0.0
CERATOPOGONIDAE
tricho 0 on spp
6.1
Culicoides spp
8.6
Pal om is complex
5.7
CHIRONOMIDAE
Ablabesmyia mallochi
7.4
blabesm is rham he gr
6.8
erillia lavi rons
3.9
erillia spp
5.7
erundiniella eumor ha
2.0
Cardiocladius spp
6.2
Chironomus spp
9.3
Cladotan tarsus cf daviesi
2.8
Cladotan tarsus sp e
4.7
Cladotan tarsus spp
4.0
Clinotanypus spp
7.8
Cor noneura spp
5.7
Cricoto us annulator complex
8.4
Cricoto us bicinctus
8.7
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 39 of 50
8/13/2021

Order
Latin Name
Tolerance Value
Cricoto us u ax
5.6
Cricoto us in uscatus gr
9.1
Cricoto us vierriensis gr
5.4
Cryptochironomus blarina gr
8.5
Cryptochironomusfulvus
6.7
Cryptochironomus spp
6.4
Cryptotendi es spp
6.2
Demicr tochironomus spp
2.2
Diamesa spp
6.6
Dicrotendi es umidus
8.8
Dicrotendi es modestus
9.4
Dicrotendi es neomodestus
7.9
Dicrotendi es nervosus
9.5
Dicrotendi es sim Boni
9.8
Dicrotendi es spp
7.2
Di locladius cultri er
8.0
Eukiefferiella brehmi gr
2.5
Eukiefferiella brevicalcar gr
2.9
Eukiefferiella clari ennis gr
6.2
Eukiefferiella devonica gr
3.4
Eukiefferiella gracei gr
4.4
Eukiefferiella pseudomontana gr
1.3
Glyptotendipes spp
8.6
Heleniella spp
0.0
H drobaenus spp
9.2
Kribiodorum perpulchrum
4.0
Labrundinia pilosella
6.2
Labrundinia spp
6.2
Larsia spp
6.5
Lo escladius spp
1.2
Micro sectra spp
2.4
Microtendi es pedellus gr
3.9
Microtendi es r dalensis gr
1.1
Microtendi es spp
4.6
Nanocladius downesi
2.4
Nanocladius spp
7.4
Natarsia spp
9.6
Nilotanypus fimbriatus
4.9
Nilotan us spp
4.1
Nilothauma spp
5.1
Odontomesa ulva
4.9
Orthocladius clarkei gr
5.6
Orthocladius dorenus
5.8
Orthocladius dubitatus
9.0
Orthocladius li nicola
5.4
Orthocladius lutei es/thienemanni
6.3
Orthocladius ni ritus
3.8
Orthocladius obumbratus gr
8.1
Orthocladius robacki
6.4
Orthocladius spp
4.4
Pa astia ortho onia
1.5
Parachaetocladius abnobaeus
0.7
Parachironomus spp
8.0
Paraclado elma spp
6.3
Paraclado elma undine
4.5
Parakiefferiella sp A
8.5
jParakiefferiella spp
14.8
Paralauterborniella ni rohalteralis
14.9
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 40 of 50
8/13/2021

Order
Latin Name
Tolerance Value
Paramerina spp
4.1
Parametriocnemus spp
3.9
Paratan tarsus spp
8.0
Paratendi es spp
5.6
Pentaneuraincons icua
5.0
Phaeno sectra obediens gr
6.6
Phaeno sectra punctipes gr
7.1
Polypedilum avice s
3.6
Polypedilum fallaxlsp A
6.5
Polypedilum flavurn
5.7
Polypedilum halterale gr
7.4
Polypedilum illinoense gr
8.7
Polypedilum loetum
2.2
Polypedilum scalaenum gr
8.5
Potthastia cf gaedii
2.4
Potthastia Ion imana
8.4
Procladius spp
8.8
Prodiamesa olivacea
8.8
Psectrotanypus d ari
10.0
Pseudochironomus spp
4.9
Rheocricoto us robacki
7.9
Rheocricoto us spp
4.7
Rheocricoto us tuberculatus
4.7
Rheo elo is spp
0.3
Rheosmittia spp
6.8
Rheotan tarsus spp
6.5
Robackia clavi er
1.9
Robackia demei'erei
4.3
aetheria t lus
7.3
tem ellinella spp
5.6
tenochironomus spp
6.3
tictochironomus spp
5.4
ublettea coffmani
1.4
m otthastia spp
4.5
northocladius spp
4.2
Tan tarsus sp 2
6.9
Tan tarsus sp 3
7.3
Tan tarsus sp 4
4.7
Tan tarsus sp 6
7.8
Tan tarsus sp A
6.9
Tan tarsus sp C
6.1
Tan tarsus sp L
4.7
Tan tarsus sp M
3.2
Tan tarsus sp P
4.8
Tan tarsus sp U
6.4
Tan tarsus spp
6.6
Thienemanniella spp
6.4
Thienemanniella xena
8.0
Thienemannim is gr
8.4
Tribelos jucundum
5.7
Tribelos spp
6.4
Tvetenia bavarica gr
3.6
Tvetenia vitracies
3.5
enochironomus xenolabis
6.6
loto us par
6.1
avrelia spp
6.1
avrelim is spp
18.6
CULICIDAE
no heles spp
18.6
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 41 of 50
8/13/2021

Order
Latin Name
Tolerance Value
DIXIDAE
Dixa spp
2.5
Dixella spp
4.9
RHAGIONIDAE
Atherix lantha
1.8
therix spp
0.9
SIMULIIDAE
Prosimulium mixtum
3.6
Prosimulium spp
4.5
imulium spp
4.9
imulium venustum
7.3
imulium vittatum
9.1
ABANIDAE
Chr so s spp
6.7
Tabanusspp
8.5
ANYDERIDAE
Proto lasa itchii
4.0
IPULIDAE
Antocha spp
4.4
Dicranota spp
0.0
Hexatoma spp
3.5
Limonia spp
9.3
Pol meda/Ormosia spp
6.5
Pseudolimno hila spp
6.2
Ti ula spp
7.5
Oli ochaeta
NAIDIDAE
Dero spp
9.8
Nais spp
8.7
Pristina spp
7.7
lovina appendiculata
8.4
t laria locustris
8.4
UBIFICIDAE
Aulodrilus pluriseta
5.6
eranchiura sowerb i
8.6
II odrilus tem letoni
9.3
Limnodrilus hoffmeisteri
9.4
Limnodrilus spp
8.5
iros erma nikolsk i
6.0
Tubi ex tubi ex
9.9
Gastro oda
ANCYLIDAE
Ferrissia spp
6.6
Laeva ex uscus
6.6
HYDROBIIDAE
Arnnicola spp
4.1
LYMNAEIDAE
Pseudosuccinea columella
7.7
to nicola spp
8.1
PHYSIDAE
Ph so s p
8.7
PLANORBIDAE
Helisoma ance s
6.6
Menetus dilatatus
7.6
PLEUROCERIDAE
Elimia spp
2.7
Le toxis spp
1.7
IVIPARIDAE
Cam eloma decisum
5.8
Bivalvia
CORBICULIDAE
Corbicula luminea
6.6
SPHAERIIDAE
Pisidium spp
6.6
haerium spp
7.2
UNIONIDAE
Elli do com Janata
4.7
Elli tio spp
4.9
Crustacea
ASELLIDAE
Caecidotea spp
8.4
Lirceus spp
7.4
CAMBARIDAE
Cambarus P. sp C
6.3
Cambarus spp
7.5
Orconectes spp
2.7
Procambarus spp
9.3
GAMMARIDAE
Cran on x spp
7.2
Gammarus asciatus
7.0
Gammarus spp
7.1
PALAEMONIDAE
Palaemonetes paludosus
16.1
Palaemonetes spp
18.7
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 42 of 50
8/13/2021

Order
Latin Name
Tolerance Value
ALITRIDAE
H alella spp
7.2
Other
ERPOBDELLIDAE
Er obdella/Mooreobdella spp
8.6
Mooreobdella tetragon
9.4
GLOSSIPHONIIDAE
Desserobdella phalera
6.6
Gloiobdella elon ata
9.1
Helobdella triserialis
9.3
Placobdella papillifera
8.2
Placobdella parasitica
8.9
PLANARIIDAE
Cura oremanii
5.5
Du esia ti rina
17.1
PYRALIDAE
Petro hila spp
13.6
SISYRIDAE
Climacia areolaris
16.5
ETRASTEMMATIDAE
Prostoma graecense
16.6
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 43 of 50
8/13/2021

Appendix 5. Resources for Identification of Freshwater
Macroinvertebrates
The following pages provide the complete visual key for identification of common freshwater
macro invertebrates to the level of Order or Family. It is also available online at
https://stroudcenter.org/macros/key/.
Other recommended resources for identification of freshwater benthic macroinvertebrates:
Louw M, Crowley K, Bartley C, et al. The Atlas of Common Freshwater Macroinvertebrates of
Eastern North America (web site). http://macroinvertebrates.org/. Accessed 3/12/2019.
Stroud Research Center. Macroinvertebrate Resources (web page).
https://stroudcenter.org/macros/. Accessed 3/12/2019.
Virginia Save Our Streams. Stream Insect and Crustacean ID resources (web page).
http://www.vasos.org/monitors-page/stream-insect-id-resources/. Accessed 3/12/2019.
Voshell, JR. 2002. A Guide to Common Freshwater Invertebrates in North America. McDonald &
Woodward Publishing. Newark, OH.
West Virginia Department of Environmental Protection. Benthic Macroinvertebrate Resources
(web page). https://dep.wv.gov/WWE/getinvolved/sos/Pages/Macros.aspx. Accessed
3/12/2019.
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 44 of 50
8/13/2021

Identif *icatiollln Guide to
♦��
Freshwater M a croi nverteb rates
OUD
SIR
TER RESEARCH CENTER
970 Spencer Road
Avondale, Pennsylvania 19311
© Stroud Water Research Center
Macroinvertebrate images prepared for
Stroud Water Research Center by Mr. Kevin Gill
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 45 of 50
8/13/2021

Abdomen: posterior body
segment of insect
Filaments: hair -like structures
Jointed leg: true legs, legs
capable of bending
Lateral: at the side
Portable case: structure made Abdomen
of leaves, twigs, or sand that
some caddisfly larvae carry
with them
Posterior: tail end of the body
Prolegs: short, stumpy leg -like
structures (not jointed)
Protrusion: part of the body
that sticks out
Segment: a section of body
Ventral: underside
Wing pads: developing wings,
often W in shape
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0
8/13/2021
Thorax
r
Head
Abdominal gills
Prolegs
Page 46 of 50
Jointed legs
Wing pads
Tail filaments
SROUD
WATER RESEARCH CENTER

Jointed legs No jointed legs
8 legs
ARACHNIDA
water mite
6 jointed legs Worm -like Body enclosed in
Go to Pape 3 (no shell) hard shell
�i
MOLLUSCA—snail, clam
Head and/or fleshy
protrusions
Go to Page 2
cra
CRUSTACEANS
Non -segmented
flat -worm
n
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 47 of 50
8/13/2021
Segmented
worms
ANNELIDAE HIRUDINEA
aquatic worm leech
UD�SIR,
TER RSEARCH (ENTER

Worm -like with distinct head or fleshy protrusion
DIPTERA — true flies
Head small, often hidden from view Distinct head
8 pairs of prolegs; posterior with 2 Fleshy protrusions Head capsule retractile, first 7 abdominal
pointed protrusions longer than prolegs at posterior end segments ringed by fleshy pseudopods
E*_— f—
ATHERICIDAE — water snipe flies TIPULIDAE — crane flies TABANIIDAE — horse flies
One end wider than the other
SIMULIIDAE —blackflies
Both ends similar width
L
CHIRONOMIDAE — midges
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 48 of 50
8/13/2021
Approximate size range
Minimum Maximum
UD�SIR,
TER RSEARCH CRTM

Six jointed legs UD
WATER RESEARCH CENTER
Portable case No portable case
(made of sand, gravel,
or plant material)
No observable wings or wing pads Wing pads or wings
present
Observable hooks at end of body No observable hooks at end of body GO t0 Page 4
Without lateral filaments
II With lateral filaments.
W Branched gills on With lateral Body ends in single,
No branched gills underside of abdomen filaments long filament
��
TRICHOPTERA
caddisflies
MEGALOPTERA
dobsonflies, fishflies, alderfiles
With fleshy
prolegs
LEPIDOPTERA
aquatic moths
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 49 of 50
8/13/2021
Without
prolegs
COLEOPTERA
beetles

Wing pads or wings present QMOUD
WATER RESEARCH CENTER
With tail filaments
Two tail filaments, Two orthree tail Three flat tail
without filaments, with filaments, without
abdominal gills abdominal gills abdominal gills
PLECOPTERA EPHEMEROPTERA
stoneflies mayflies
ODONATA
damselflies
Without tail filaments
Wing pads Wings
Hard wing Wings
Large mouth, coverings with leathery, tips
hinged centerline overlapping
ODONATA COLEOPTERA
dragonflies adult beetles
City of Durham, WQ Unit, Benthic Macroinvertebrate Monitoring SOP, v. 1.0 Page 50 of 50
8/13/2021
HEMIPTERA
true bugs