5.2 MassDEP/MassCZM Standard Operating Procedures and Data ...neiwpcc.org/nebawwgold/documents/ManualSections/Section 5_2.pdf · Adrienne Pappal Commonwealth of Massachusetts Executive

5.2 MassDEP/MassCZM Standard Operating Procedures and Data Forms

Standard Operating Procedures: Salt Marsh Assessment, May 8, 2012 DRAFT

Development of a Comprehensive State Monitoring and

Assessment Program for Wetlands in Massachusetts:

Salt Marsh Component

Appendix A

Standard Operating Procedures: Assessment of Salt Marsh

Wetland Communities

Phase 2: 2009-2012

Prepared by:

Marc Carullo

Jan Smith

Adrienne Pappal

Commonwealth of Massachusetts

Executive Office of Energy and Environmental Affairs

Office of Coastal Zone Management

251 Causeway Street, Suite 800, Boston, MA 02114

with

Department of Natural Resources Conservation

University of Massachusetts, Amherst

and

Commonwealth of Massachusetts

Executive Office of Energy and Environmental Affairs

Department of Environmental Protection


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Standard Operating Procedures: Salt Marsh Assessment

1. Scope and Application

This SOP establishes a standard set of procedures to be followed for data collection

toward the development of a Site Level Assessment Method (SLAM) for Massachusetts

salt marshes and to verify and calibrate the Conservation Assessment and Prioritization

System (CAPS) as a mechanism for a landscape level analysis (Level 1) of ecological

integrity. This project will focus on assessment of wetland biological community

condition in salt marshes.

Described below are the procedures that will be followed in collecting data on

macroinvertebrates, vascular plants, and habitat (e.g. tidal hydrology, marsh zonation,

open water patch composition, etc.) to serve as a basis for development of a SLAM and

Indices of Biological Integrity (IBIs) for salt marsh wetlands.

2. Summary

This SOP is applicable for salt marshes throughout Massachusetts. This wetland

community is known as tidal fringe wetland in the hydrogeomorphic (HGM)

classification and estuarine emergent (E2EM) in the Classification of Wetlands and

Deepwater Habitats of the United States. Data collection for the pilot study (phase 2b)

focused on salt marshes from the NH/MA border in Salisbury, MA, south to Hull, and

Cape Cod. The watersheds included in the site selection process were Cape Cod, Charles

(Boston Harbor), Ipswich, Merrimack, Mystic, Neponset, North Coastal, Parker, and

Weymouth & Weir Watersheds. These watersheds provided an appropriate mix of urban,

suburban, and relatively undeveloped coastal areas. They include salt marshes that are

representative of those found throughout Massachusetts. By limiting field work to these

select watersheds, we were able to sample a greater number of sites toward statistical

validation of the CAPS model. Investigators had conducted previous assessments in these

watersheds and were familiar with the landscape. This local knowledge aided logistical

planning. Target sites for the operational study (phase 2c) are open to all watersheds

containing DEP-mapped salt marsh. Additional watersheds include Buzzards Bay,

Islands, Mount Hope Bay, Narragansett Bay, South Coastal, and Taunton Watersheds.

Sampling sites will again be selected using a stratified random process, as further

described in Section 8. Field data collection will involve sampling of several biotic

communities to determine if 1) there is a dose-dependent response in various attributes of

the biological community to stressors within the landscape and 2) to verify and calibrate

the ecological integrity metrics that are utilized in the CAPS model. Characterization of

the wetland and assessment of its biological condition will be conducted in the field by

assessing habitat, hydrology, macroinvertebrates, and vascular plants.


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3. Safety Considerations

Fieldwork will not be conducted during heavy rain events or unsafe conditions

such as electrical storms or high wind events. Practice ―safety first.‖

Sampling will always be conducted by two or more persons, unless otherwise

approved by the field manager.

All persons must carry cell phones or other emergency communication devices

while sampling. It is recommended they be waterproof or stored in a waterproof

case.

If there is no safe access to a plot point, the field sampling will either be moved to

a location with safe access—following protocol as described in Section 8 of this

SOP—or not be conducted for that site if such a location is unavailable.

Flagging tape will be used to mark access point locations for safe exit, in

instances where such locations could be difficult to find as deemed appropriate by

field crew.

Attempts to access a plot point where an onshore breeze of intensity that results in

a ―small craft warning‖ or above for two or more days, and that creates an

atypical tide condition shall be assessed for safety prior to sampling. At any time

during the sampling, should the incoming tide not recede as predicted (i.e. tide is

still in flood stage when it should be in ebb stage), all monitoring shall stop and

the site evacuated until such a time when the tide has receded to a point where

sampling can safely resume.

If a boat, canoe, or kayak is used to access a plot point, all persons must wear

personal flotation devices while on the water.

All watercraft used must comply with USCG required safety equipment per 46

U.S.C, Chapter 43. An online Vessel Safety Check interactive program is

available from USCG at http:/www.uscgboating.org/safety/vsc.htm.

Good judgment will be used in selecting clothes and personal protection items.

Common items needed include: extra clothing, sunshade, sunscreen, hats, insect

repellent, and waterproof knee boots—chest waders or hip waders for highest

anticipated depths. Any staff not dressed appropriately for field work should not

participate in the survey. Proper footwear is a must (e.g., no ―flip-flops‖ for field

work).

Good judgment will be used in walking on marsh surfaces; mosquito ditches will

be circumvented, or when deemed possible, crossed with caution.

Private property will be respected using the following guidelines.

o If property is in close proximity to buildings or other heavily used areas,

landowner permission will be sought

o Posted property will not be accessed without permission of the landowner

o Otherwise, sampling will proceed without any special effort to gain

landowner permission


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4. Sample Collection, Preservation, and Handling

Macroinvertebrates will be collected and preserved in 90% ethyl alcohol solution and

placed in a cooler with ice until ready to be transported to the laboratory for sorting and

identification or to a storage facility for later processing. Samples will be checked

periodically to ensure that they contain ample alcohol solution if they are not sorted

within 2-3 days. Samples will not be held for more than two weeks before sorting to the

family level. Each sample will be placed in a plastic collecting bag that is labeled with

site number, site name, date of sampling, sample number, sampling method, and name of

sampler. The Macroinvertebrate Samples Record Check form (Appendix C) will be used

to catalog the collection and processing of samples.

The collection of vascular plants will be limited to species that cannot be identified in the

field. For species that cannot be positively identified in the field samples will be collected

for lab identification and photographed for digital preservation. Taxonomic identification

at the species level (preferred) or genus level (if species identification is not possible) will

be achieved in the laboratory through the use of field guides, technical keys, and

reference to regional herbaria housed at research universities such as UMass. Samples

will be labeled in the field with the plant ID (e.g., ―unknown sedge #1‖) site location,

date, and person who collected the sample, and assigned a code in the laboratory for use

in digital preservation.

5. Equipment/Apparatus

Before leaving for the field the Field Manager will confirm the following equipment is

available (m = macroinvertebrates, v = vegetation; habitat complexity and water

chemistry sampling are included with vegetation and macroinvertebrate equipment,

respectively, as they will be conducted alongside those sampling efforts).

Auger (2.5-inch diameter) [m]

Baster (pipette) [m]

Bayonet [v]

Binoculars [v]

Bleach solution (2%, for decontamination of gear) [m,v]

Bucket (2x, 5-gallon) [m]

Bucket (small) [m]

Clipboard [m,v]

Compass [m,v]

Cooler with ice [m]

Data forms [m,v]

Digital camera w/extra batteries [m,v]

D-Net (500 micron mesh size) [m]

Ethyl alcohol (90%) [m]

Field guides and technical keys [v]

Field notebook [m,v]

First aid kit [m,v]


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Forceps [m]

GPS (Global Positioning System) [m,v]

Hard-bristle brush (for decontamination of gear) [m,v]

Labels for samples [m,v]

Lens cloth (for refractometer, soft cotton flannel) [m]

Location maps [m,v]

Magnifying lens [m]

Measuring tapes (2x, 100-meter) [m,v]

Pens and pencils [m,v]

Permanent marker [m,v]

Plastic collecting bags [m,v]

PVC stakes (3x) [m,v]

Protective gloves [m]

Quadrat frame (18‖x18‖) [m]

Refractometer [m]

Scissors or jack knife [m,v]

Sieve (No. 8, 2,360 micron) [m]

Sieve bucket (500 micron) [m]

Spatula [m]

SOP [m,v]

Tap water [m,v]

Thermometer [m]

Tide chart [m,v]

Throw rope [m,v]

Trowel [m]

6. Reagents

Bleach solution (2%)

Ethyl alcohol solution (90%)

Tap water

7. Calibration & Training

Equipment calibration procedures for the GPS units and salinity refractometers will be

done according to the manufacturers’ recommendations. See section 2.6 of the QAPP for

details.

Field crew members will have sufficient previous training and experience to reliably

conduct field data collection or they will receive training from the CZM QA Manager,

CZM Project Manager, and/or other project scientists with relevant expertise. The

Macroinvertebrate Field/Lab Managers will ensure that all macroinvertebrate field crew

members receive specific training on macroinvertebrate sample sorting.

All field crew members will receive training from the CZM QA Manager on appropriate

QA/QC procedures.


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8.0 Procedures

Sampling will occur between June 15 and September 30, to ensure adequate assessment

of the targeted wetland biotic communities. Sample locations for open water/low

marsh/high marsh (i.e., inner marsh) will be selected randomly within salt marshes (as

depicted in MassDEP Wetlands mapping data; 1:12,000 based on photography from

1993-1999), stratified into 100 bins (deciles of the first principal component of CAPS

metrics crossed with deciles of the second principal component; metrics include habitat

loss, watershed habitat loss, wetland buffer insults, traffic intensity, sediment intensity,

toxic pollution, edge predators, imperviousness, salt marsh ditch density, tidal restriction,

connectedness, and similarity, among others). All points, including those sampled in

previous years, will be separated by at least 500 meters. Approximately 500 points will

be selected to allow for rejection based on the following criteria:

o The point is not within 200 meters of a salt marsh creek (each sampling point will

be moved not more than 200 m to the nearest tidal creek suitable for sampling);

o The creek is not suitable for auger and D-Net macroinvertebrate sampling (e.g.

channel width is less than 2 m; bank height is less than 1 m; bank height and/or

condition could compromise safety);

o The point is not accessible due to physical barriers;

o The point is not accessible due to legal barriers (i.e. request to access private

property is denied).

o Sites will be assessed on these criteria by CZM and DEP personnel using 2008

and 2009 orthophotos, 2008 oblique photos, various GIS layers, field

reconnaissance, and local area knowledge.

Sample locations for marsh border will be placed at edges of salt marshes, stratified

across quartiles of the habitat loss metric ( development intensity). One hundred fifty

points will be selected, with a goal of visiting 50 points. All marsh border points will be

separated by at least 500 meters.

A random identifier will be assigned to each point to obscure the CAPS metrics

represented by each point. Field personnel will not have access to the original values until

after the field season, thus sampling will be blind with respect to CAPS predictions.

Field personnel will visit points in (randomly assigned) numerical point order as

consistent with field logistics. For example, the first acceptable (not rejected in aerial

photo surveys, without access problems, and with appropriate habitat on the ground) open

water/low marsh/high marsh points (i.e., inner marsh) will be visited.

GPS navigation will be used to locate each wetland plot. GPS precision must be 10 m or

less and the navigator will stop and establish the plot once the distance to plot center is 0

m. In the case of GPS interference from tree-canopy or atmospheric effects wait 10

minutes for satellite reception to improve.

It will not be necessary to hit the plot exactly (since it's randomly selected) it just needs to

be selected without bias. However, a reasonably precise GPS point is needed of where the


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plot actually ends up. The strategy is to (1) do the best we can when locating the plot and

(2) take a precise location (precision ≤ 10 m RMS) once the plot has been

established. Field workers will keep trying until they get good GPS coverage.

8.1 Establishing Sampling Areas

Assessment Area A: For areas where vegetation (inner marsh only), macroinvertebrate,

habitat complexity, and human disturbance sampling will occur:

A 50 x 100 m rectangular plot will be used to sample the wetland point (Figure 1). A

baseline transect, Transect A, will be created by extending a 100 m tape along the bank of

the salt marsh creek, bay, or salt pond so that the wetland point occurs at 50 m. Stakes

will be placed along Transect A at 0 m, 50 m, and 100 m. Three additional transects will

be run perpendicular to Transect A, originating at Transect A and ending at 50 m or the

salt marsh border community, whichever is closer. A consistent compass bearing will be

used to run these transects (e.g., Transects 1, 2, and 3 will be laid on a bearing of 225

degrees from the bank (Transect A) to 50 m or the salt marsh border community,

whichever is closer). The field crew leader based on observed vegetation characteristics,

slope, and elevation will delineate the salt marsh border community. Geographic

coordinates will be collected at the base of each perpendicular transect (i.e., Transects 1,

2, and 3) using a handheld GPS unit.

Transect A will be used to collect macroinvertebrate samples. Site conditions may require

shortening the transect to no less than 70 m. The following locations of macroinvertbrate

sampling stations along Transect A are based on a 100 m transect. They will be adjusted

accordingly if Transect A is shortened. Macroinvertebrates will be sampled using the dip

net method along Transect A at 15-21 m and 75-81 m.

The auger method will be used to sample macroinvertebrates at 15 m and 85 m. In

substrate with a high concentration of organic material, determined by visual inspection,

compositing and sub-sampling methods will be used to collect two additional samples—

one from four auger samples collected equidistant from 20 m to 45 m, and the other from

four auger samples collected equidistant from 55 m to 80 m (Figure 4). In sandy

substrate, compositing will not occur due to the potential for damage to soft-bodied

macroinvertebrates, however two single samples will be collected at 40 m and 60 m.

Quadrat sampling will occur on the marsh surface along Transect A at 25 m and 75 m.

Transects 1, 2, and 3 will be used to collect plant and habitat complexity data using point-

and line-intercept methods, respectively. Plant and macroinvertebrate transects will be

marked with flagging to prevent trampling.


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Figure 1. Example plot diagram for inner marsh sampling. Transect A is a baseline transect from which

Transects 1, 2, and 3 are run to collect vegetation and habitat complexity data. Transect A is run along the

bank of a prominent water feature such as a tidal creek, bay, or salt pond. It is also used to sample macro-

invertebrates and surface water salinity. Transects 1, 2, and 3 are run perpendicular to Transect A at 0 m, 50

m, and 100 m. The location for all samples (vegetation, macroinvertebrates, water chemistry, etc.) will be

noted on the plot diagram.

A sampling point will be moved if any of the following conditions are encountered.

o The length of any perpendicular transect (Transects 1, 2, and 3) is <30 m, as

may occur in narrow wetlands (e.g. fingerlike projections, fringe marshes).

o The length of the baseline transect (Transect A) is <70 m, as may occur in

narrow wetlands, particularly in urban landscapes.

The sampling point will be moved to the nearest location that does not violate the

previously stated conditions, but no greater than 200 m away. If a suitable sampling point

cannot be found within 200 m of the original point the site will be dropped and another

sampling point from the same bin selected.

A site will not be sampled if the field manager determines that the wetland is neither a

salt marsh nor is in transition to/from a salt marsh. This may occur with error in wetland

mapping.

Assessment Area B: For areas where vegetation (marsh border only) sampling will occur:

A rectangular plot will be used to sample the wetland point. Transect A will be created by

extending a 70 m tape along the upland/marsh border edge. Three additional transects

will run perpendicular to Transect A at 0 m, 35 m, and 70 m originating at Transect A

and ending at the high marsh/salt marsh border edge or 50 m, whichever is smaller. The

minimum transect length will be 5 m. The salt marsh border community will be

delineated by the field crew leader based on vegetation characteristics. Plants commonly


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found in the salt marsh border communities of New England include Iva frutescens,

Myrica gale, Panicum virgatum, and Solidago sempirvirens, among others.

Figure 2. Example plot diagram for marsh border sampling. Transect A is a baseline transect from which

Transects 1, 2, and 3 are run to collect vegetation data. Transect A is run along the edged shared by upland

and salt marsh border communities. Transects 1, 2, and 3 are run perpendicular to Transect A at 0 m, 35 m,

and 70 m. The location for all samples will be noted on the plot diagram.

8.2 Overview of Wetland Biotic Community and Habitat Assessment

Each point will be sampled for vascular plants and macroinvertebrates. Samples will be

taken within a 50x100 m rectangular plot or a 50 m radius circle-plot. Samples will be

analyzed to determine if the attributes of the biotic communities show a dose-dependent

response to anthropogenic stressors in the landscape as measured by CAPS metrics. In

addition a habitat assessment will be conducted to characterize the assessment area. A

detailed description of the plot (includes hydrology, anthropogenic disturbance, etc.) will

be recorded on field data forms.

8.2.1 Habitat Assessment

(a) Habitat complexity

Patch composition of open water features, plant communities, and marsh zones will

be determined to assist in the characterization of the wetland. Transects 1, 2, and 3

will be walked to observe and record points of transition.

From Transect A, walk three 50 m transects and record the number and types of

natural transitions using the line-intercept method. Habitats are defined as

heterogeneous plant communities (e.g. Spartina patens/Distchlis spicata mix) and

hydrogeomorphic features (e.g. panne, pool, and creek, plus ditch) in each of the


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following marsh zones: low marsh, high marsh, salt marsh border, and brackish

border (Figure 2). Both a plant community and hydrogeomorphic feature can be

recorded at the same location (e.g. Spartina alterniflora-short and panne in the high

marsh zone). Do not record transitions for habitats that measure less than one meter

along the transect.

Record the tape measure reading for each habitat transition to the nearest tenth of a

meter in the appropriate box(s) (e.g. 5.5 m). Separate recorded habitat transitions with

a comma if that habitat occurs more than once on a transect. List no more than three

plant species when describing a community. Record species in order of dominance,

with the first species being the most dominant. For example, record a habitat patch

that includes S. patens (75%), J. gerardii (25%), and P. maritimum (< 1%) as S.

patens / J. gerardii mix. Record start and end distances (range) for all

hydrogeomorphic features (e.g. Panne = 25-30 m). Include patches and features that

are offset up to 1 m from the transect. Complete a Habitat Complexity Form for each

transect.

Figure 3. Plant zonation in northeastern salt marshes. This diagram shows the major plant zones and

dominant species; see text for details and Tiner (1987), Mitsch and Gosselink (1993), or Bertness

(1999) for a description of salt marsh vegetation patterns. (Adapted from Carlisle, et al, 2002).

(b) Tidal hydrology (Sampled in 2009 only)

Tidal flow will be sampled at a subset of known or potential tidal restrictions. The

two main goals are: 1) determine the relative magnitude of tidal restriction at select

sites, and 2) develop a magnitude of tidal restriction data set that will be used to train


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a tidal restriction model developed from remotely sensed data interpretation (for

information about the model see Tidal Restriction – Aerial Photo SOP – Appendix R

of the main project QAPP (Forested Wetlands)). Locations will be identified using a

combination of field observations, remotely sensed data, local knowledge, and best

professional judgment. This effort is being managed under the Tidal Restriction

Assessment Standard Operating Procedures (QAPP, Appendix B).

(c) Salinity

Salinity will be measured concurrently with macroinvertebrate sampling.

Measurements of salinity can help to explain the diversity, distribution, and

abundance of plants (as recognized in zonation patterns) and animals in salt marshes.

Two measurements will be made at each plot using a hand-held refractometer (see the

QAPP, Appendix F for the operation manual).

Take readings from surface water at the start of each D-Net sample collection

(presumably 15 m and 75 m along Transect A, depending on direction of tidal flow).

The minimum distance between readings must be 20 m. Note on the Plot Diagram

form the location along Transect A from which readings were taken.

Remove prism cover, place one or two drops of liquid on the prism surface, and

replace cover. Aim the front end of the refractometer to the direction of bright light,

and adjust the adjusting ring of diopter 5 until the reticle can be seen clearly. The

corresponding reading of the light/dark boundary will give the percent of salt content

of liquid on left and parts per thousand on right.

Adjustment of null: open the cover plate, place one or two drops of distilled water on

the surface of the prism. Close the cover plate, rotate and adjust the correcting screw

to make the light/dark boundary coincide with the null line. Open the cover plate to

clean the water off the prism surface with a soft cotton flannel cloth.

(d) Human disturbance

Visual observations of human disturbance to the wetland will be noted. Investigators

will note the following activities on the Human Disturbance field data form,

describing the type, extent, and severity of each disturbance.

Walk the perimeter of the sample plot and describe on the Human Disturbance Form

the types, extents, and severity of disturbances within a 100 m buffer of the sample

point (200 m diameter circle depicted on the site map). Disturbance types are listed

below. Also record any of these indicators of disturbance when encountered while

implementing other elements of the SOP.

Mark locations of disturbances on the site map according to the Human Disturbance

Form (QAPP, Appendix C).


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Water control structures (culvert, tide gate, dam, weir, storm water input, fill

(road/railroad), ditching, channelization, and other human activity affecting

the hydrology of the site

Soil disturbance (filling, sedimentation, haying)

Obvious spills of pollutants

Direct point or non-point source discharge from agricultural operations, septic

or sewage treatment systems, or storm water affecting water quality of the site

Walking trails, horse trails, and roads (excluding wildlife trails)

Small-craft boating and boat storage

Presence of trash/litter

Presence of garbage dumping

8.2.2 Protocols for Sampling, Sorting, and Identifying Biotic Communities

8.2.2.1 Macroinvertebrates

Macroinvertebrates will be sampled as indicators of changes in tidal flow, vegetation

cover, salinity regime, water quality and community composition, and ecosystem health.

Macroinvertebrates will be sampled from August-September. Three types of samples will

be collected at each assessment area: quadrat samples at the top of the salt marsh creek

bank, and D-Net and auger samples in the creek.

(a) Quadrat sampling

Quadrats are used to sample invertebrates that exist on the upper edge of the

estuarine stream bank or seaward marsh edge. We expect to find crabs, mussels,

barnacles, amphipods, isopods, flies, spiders, grasshoppers, and mites in this

habitat. Two samples will be collected along Transect A at 25 m and 75 m in each

area (Figure 1). If these locations are not typical of the bank condition, sample

collections will be located elsewhere along Transect A, while keeping a minimum

distance of 30 m from each other. Protective gloves are used for this sampling

technique.

At each sampling location, methodically work the hands backwards and forwards

across the surface of the ground within the frame, and identify, count and record

every living invertebrate (to family-level) that you encounter. Since barnacles are

usually too numerous to count, record their abundance with the following

notation: + = rare, ++ = common, and +++ = abundant. Place organisms that

cannot be identified to family-level into a re-sealable plastic bag, and label (see

instructions in Section 8.2.2.1 (d)). Results are recorded on the Quadrat Samples

Record Form (QAPP, Appendix C). If samples are collected, record the

information on the Invertebrate Sample Record Form (QAPP, Appendix C). Mark

actual sample locations on the Plot Diagram. Thoroughly rinse the frame.


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(b) D-Net sampling

D-Nets are used to collect invertebrates from shallow water environments (e.g.,

tidal creeks) at low tide. We expect to find mollusks, polychaete worms,

amphipods, isopods, and other organisms requiring constant inundation with

seawater. Two samples will be collected along Transect A from approximately

15-21 m and 75-81 m in each area (Figure 1). We will aim to collect samples

from different habitat types, such as banks and vegetated margins, different

substrate types, woody debris, and floating alga mats, where possible. Transect

locations may be moved to incorporate these habitats, but a minimum distance of

30 m should be maintained between the D-Net samples, and a minimum distance

of 10 m should be maintained between the D-Net samples and auger samples, as

shown in Figure 1. Sample locations will be recorded.

At each sampling location, place the flat side of the D-Net on the surface substrate

in approximately 0.3 m of water and hold the net perpendicular to the substrate.

Walk against the direction of tidal flow for 6 meters along Transect A while

pulling the net through and over different habitats. Bring the net containing the

sample to the surface for retrieval. Gently swish the net back and forth in the

stream to allow fine silt and sand to pass through the mesh, being careful not to

lose organisms. Place the contents of the inverted net over a bucket half filled

with water and wash all debris and invertebrates off the net and into the bucket.

Use forceps to remove any organisms that remain on the net, and place these in

the bucket. Pour the contents of the bucket through a standard US No. 30 brass

sieve to remove the water. Place the contents of the sieve into a re-sealable plastic

bag, and label (see instructions in Section 8.2.2.1 (d)). Make sure that no

invertebrates are left on the sieve. Sample labels will include information such as

sample number, field site ID, sample station, date, names of collectors, sampling

method, and preservative used. Record the sample information on the

Macroinvertebrate Samples Record Check Form (QAPP, Appendix C). If a large

number of snails and crabs are collected in a sample, identify them in the field,

record the numbers of each family (and if possible, species) on the datasheets, and

then return them back to the water alive. Wash out the D-Net to remove all

remaining debris.

(c) Auger sampling

A 2.5-inch diameter auger is used to collect a sediment or substrate sample from

the creek. We expect to find a variety of worms, snails, clams, amphipods,

isopods, and other organisms that live on or within the substrate. Where organic

material is the dominant substrate, four samples will be collected along Transect

A in the following manner:

Sample 1: Single sample collected at 15 m.


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Sample 2: Composite sample from four samples collected 5-7 m from each

other beginning at 20 m and ending at 45 m. Composite sample

will be sub-sampled, as described below.

Sample 3: Composite sample from four samples collected 5-7 m from each

other beginning at 55 m and ending at 80 m. Composite sample

will be sub-sampled, as described below.

Sample 4: Single sample collected at 85 m.

Figure 4. Schematic of auger sampling stations along Transect A.

Four samples will be collected along Transect A at 15 m, 40 m, 60 m, and 85 m

when sand is the dominant substrate.

At each sampling location, fill two 5-gallon buckets with seawater to be used

later. Place the No. 8 sieve on the bottom of the sieve bucket. Hold the auger

perpendicular to the water surface above the point from which the sample will be

taken. Push the auger downward into the sediment until the bucket of the auger is

half embedded in the substrate. Turn the auger handle to help force the auger into

the substrate. Carefully pull the auger out of the sediment and quickly place the

sieve bucket beneath the auger so that none of the sample is lost. If the sample is

lost, repeat no less than 5 m away in the direction opposite that which the tide is

flowing (e.g., 5 m upstream if there is an ebb tide). Keep the sieve bucket under

the auger and return to the bank, where the remaining auger contents will be

emptied into the No. 8 sieve.

For composite samples, repeat the above method three times, adding the material

from a total of four auger samples collected from 20-45 m or 55-80 m along

Transect A (see Figure 4) to the No. 8 sieve.

Place the sieve bucket into one of the 5-gallon buckets filled with seawater. With

the spatula, gently scrape large debris off the No. 8 sieve, then stir and mix to

suspend materials and organisms within the sieve bucket. Slowly lift the No. 8

sieve out of the bucket while collecting and mixing the suspended material. Take

the No. 8 sieve out of the sieve bucket and discard its contents.

Replace sediment-laden water with clean seawater if necessary, or switch to a

second 5-gallon bucket filled with seawater. Dip the end of the sieve bucket into


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the 5-gallon bucket of water and gently swirl once to redistribute the remaining

material evenly across the sieve bucket while lifting. Drain water and divide the

contents of the sieve in half using your spatula and the dividing line marked on

the bucket. Gently scoop out ½ of the sample, as labeled on the bucket, and

discard. Repeat mixing, sieving, and halving procedure once again for the

material remaining in the sieve bucket. Again, gently scoop out ½ of the sample,

as labeled on the bucket, and discard. Place the remaining half of the material in a

sample bag with a label including the site ID and name, date, and sample ID (see

instructions in Section 8.2.2.1 (d)) and record the sample information on the

Invertebrate Sample Record Form (QAPP, Appendix C).

For single samples, rinse the material in the manner described above, but place

the entire washed contents of the sieve bucket into a labeled sample bag—do not

sub-sample.

Add alcohol as appropriate to preserve. Clean the auger by swishing it back and

forth in the water. Repeat for all samples.

(d) Sample bagging and labeling

Use the following procedures to bag and label samples:

1. Using a permanent ink marker, label all sample bags with the following

information: sample number (begin each sample number with ―A‖ for

auger samples, ―D‖ for D-Net samples, and ―Q‖ for any quadrat samples

collected; e.g., A1, A2, D1, D2), plot identification, site name, and date.

This can be done before going into the field. In addition, record this

information on waterproof paper labels (e.g., Rite in the Rain® paper) and

place in their respective sample bags as back-up labels.

2. Gently flood all bagged samples with 90% concentration ethyl alcohol and

seal carefully.

3. Record the sample information from Step 1 on the Macroinvertebrate

Samples Record Check Form (QAPP, Appendix C).

4. Place each bagged sample in a separate second bag—so that each sample

is double-bagged—and place in a cooler with ice to prevent heating in hot

weather.

5. Transport and store samples in an air-conditioned laboratory (or similar

workspace) or a refrigerator for no longer than two weeks before sorting.

Samples should be checked every 2-3 days if they are not sorted

immediately to ensure they have ample ethyl alcohol preservative.

(e) Sample sorting

Use the following procedures for sorting samples:


16

1. Empty the contents of a sample into the standard US #30 sieve. You

should place the sieve over a bucket so that sediment is not washed down

the drain.

2. Gently rinse the sample under tap water to remove fine organic detritus,

silt, and clay. Place the sieved sample into a white sorting tray (one small

handful at a time, if necessary). You must be careful to remove any

organisms that may be stuck on the sieve.

3. Place the sorting tray under a desk light or magnifying lamp, and using the

magnifying lens and forceps, remove invertebrates from the sediment and

place them into a large (40 mL) vial two-thirds filled with 70% or higher

concentration of alcohol.

4. Immediately after you have finished sorting, have the macroinvertebrate

lab manager scan the debris in the sorting tray to double check your work.

Tightly seal and label each vial (two for each sampling station: one for D-

Net and one for auger, and register the sample on the Macroinvertebrate

Samples Record Check Form (QAPP, Appendix C).

5. Samples can be identified and counted any time after the sorting has been

completed, but should not be left for more than six months because

alcohol in the vial sometimes evaporates and ruins the sample. Sample

degradation can be prevented by routine checks and preservative added if

necessary.

(f) Sample Identification

Use the following procedures for identifying samples:

1. Pour the vial contents of the D-Net sample or auger sample into petri dish.

Maintain separate D-Net and auger samples. Make sure that no organisms

remain in the vials.

2. Place the petri dish under the dissecting scope set at 10X magnification,

and in a deliberate, systematic manner, scan back and forth, identifying

organisms as you go. You may need to increase the magnification to see

finer details.

3. Using Weiss (1995), Pollock (1998), and other references, identify the

invertebrates to family level.

4. Record and count each taxon on the Laboratory Bench Form (Appendix

C).

5. Immediately after you identify and record a specimen, return it to a

labeled vial two-thirds filled with 70% or higher concentration alcohol.

There should be one vial per sample.

6. Label and safely pack the vials for return to your project coordinator so

that someone can reexamine specimens or identify them to a lower level of

taxonomy at some future date.

7. If you have doubts about an organism’s identity, consult with a marine

invertebrate specialist. Place the specimen in question in a separate vial

with alcohol and a complete label. Send the specimens to a specialist for


17

verification, and add to your records later. Alternatively, arrange to have a

taxonomist present during an identification session to provide assistance.

8. Record the completion of this process for each sample on the

Macroinvertebrate Sampling Record Check Form (Appendix C), which

traces the sample collection, sorting, and identification to this stage.

9. On the Laboratory Bench Form (Appendix C), add the data from the

quadrat sample taken from the same sampling station. Enter the total

number of organisms for each family or taxonomic group, the number of

different types of taxa you identified, and the resulting total abundance for

the completed composite sample.

10. Repeat this process for the remaining two sample stations, using a separate

Laboratory Bench Form for each station.

11. Once collecting, sorting, and identifying samples for a site has been

completed, there will be two completed copies of the Laboratory Bench

Form (one for each sample station).

12. Samples and the Sample Record Check Form are to be returned to the

project leader for archival action. Similarly, return all Quadrat Samples

Record Forms and Laboratory Bench Forms to the project leader once

they are completed.

8.2.2.2 Vegetation

Vegetation data will be collected as an indicator of community composition and species

diversity (proportion of native to invasive), and provide useful information on potential

threats to natural systems. Invasive plants named as such in this assessment are those

currently regulated by the Commonwealth of Massachusetts (Somers et al 2006). Data

collection will occur near average low tide from mid-July to mid-September. The

protocol for sampling inner marsh and marsh border plots is the same.

a. Estimate species richness and relative abundance of all vascular plants in a 50x100m

plot using a modified point-intercept method. A single bayonet (vertical projection)

will be used to record species at fixed intervals along each transect. Walk along the

transect opposite the side used to lie the transect to avoid trampling samples. Set the

transect interval by dividing the transect length by 10; this will produce 11 point

intercept locations along each transect. As stated in section 8.1, the minimum and

maximum vegetation transect lengths will be 30 m and 50 m, respectively. Tally each

plant species intercepted by the bayonet from marsh surface to canopy at each fixed

interval along the transect. Record the transect length and interval along with species

occurrence on the Vegetation Form (QAPP, Appendix C).

b. Following transect sampling conduct a 15-minute walk around (within) the entire plot

and list species not encountered on transects on the Vegetation Form. Enter zero for

intervals 1-11 for these species.

c. Collect unknown species for offsite identification by a second expert. Place each

species in a separate collecting bag labeled with plant ID (e.g., ―Unknown #1, etc.),

plot ID and date. Take digital photographs on site as needed. List PhotoID # next to

unknown plant ID for interval 1 on the Vegetation Form.


18

d. Refer to resources on regional flora if necessary (e.g., Tiner, 1987; Gleason &

Cronquist, 1991).

8.3 Protocol for Decontamination of Field Equipment

Inspect all equipment for debris and remove before leaving a site. Dispose of debris in a

trash bag or on dry, high ground. When possible, leave equipment to air dry and inspect

to remove any remaining plant fragments. Scrub and rinse equipment to remove any

additional debris with a hard-bristled brush and 2% bleach solution. Clean the salinity

refractometer according to manufacturer’s recommendations.

9. Quality Control

Compliance with procedures in this SOP will be maintained by following the items

described below. See sections 2.5 and 2.6 of the QAPP for additional details about

QA/QC measures.

Computer aided use of stratified random sampling procedures for site

selection (accuracy, representativeness)

Use of standardized sampling procedures such as transect and time-

constrained sampling (precision, accuracy, representativeness)

Prompt review and documentation of any changes to the SOPs (precision,

accuracy, comparability)

Use of highly qualified field scientists (precision, accuracy, comparability)

Rigorous training and mentoring of less experienced technicians in both

structured and informal settings, the latter on an as needed basis

(precision, accuracy, comparability)

External validation of taxonomic identification for taxa with which the

field crew has had limited prior experience (90% of samples); minimum of

10% of total samples (precision, accuracy)

Daily checks to ensure that data forms are completely filled out

(completeness)

The Quality Assurance Manager is responsible for periodically inspecting the methods

used and inconsistencies will be documented and if possible, corrected. Any significant

changes will be made in coordination with MassDEP and EPA.

10. Interferences

Inclement weather (heavy rain) may interfere with our ability to collect representative

data on a variety of parameters. Severe weather may delay field data collection due to

safety concerns. Access may be a challenging aspect of data collection in more developed

areas of the study area. Posted property or sites that are too difficult to access or unsafe to

sample will be replaced with alternative sites from the same stratified sampling bin.

11. Preventative Maintenance


19

Field equipment will be inspected by the CZM Field Manager each day before going out

to collect field data. At the field site equipment will be tested prior to data collection to

ensure that it is working properly. Equipment will be subject to regular maintenance as

needed and as recommended by the manufacturer. GPS accuracy will be assessed once a

month by a check of any units used in the field with a known location. See section 2.6 of

the QAPP for more detail.

12. Corrective Actions

Data quality control ensures high quality data, however we are prepared to re-measure

any plots within the same season or period of monitoring as needed (e.g., data are

missing, samples are lost or compromised, etc.). Any plots that contain data that cannot

be resolved will be removed from the data set.

13. Waste Minimization and Pollution Prevention

Care will be taken to avoid transport of vegetation and soil to other sites. This will be

done by thorough inspection of all equipment and clothing prior to departure from a site.

Invasive plant samples will be disposed of in a way to avoid accidental release into the

environment.

14. References

Bertness, M.D. 1999. The Ecology of Atlantic Shorelines. Sinauer Associates, Inc.

Sunderland, MA

Brinson, M. M. 1993. A hydrogeomorphic classification for wetlands. Technical Report

WRP-DE-4, U.S. Army Engineer Waterways Experiment Station, Vicksburg, MS. NTIS No.

AD A270 053.

Carlisle, B.K., A.M. Donovan, A.L. Hicks, V.S. Kooken, J.P. Smith, and A.R. Wilbur. 2002.

A Volunteer’s Handbook for Monitoring New England Salt Marshes. Massachusetts Office of

Coastal Zone Management, Boston, MA.

Carey, J.M. and M.J. Keogh. 2002. Compositing and subsampling to reduce costs and

improve power in benthic infaunal monitoring programs. Estuaries 25(5):1053-1061.

Connors, B. 2006. Protocols for Decontaminating Biomonitoring Sampling Equipment. ME

Department of Environmental Protection DEPLW0641.

Cowardin, L.M., V. Carter, F.C. Golet, and E.T. LaRoe. 1979. Classification of wetlands and

deepwater habitats of the United States. FWS/OBS-79/31. U.S.D.I. Fish and Wildlife

Service, Washington D.C.

Gleason , H.E. and A. Cronquist. 1991. Manual of the Vascular Plants of Northeastern

United States and Adjacent Canada. 2nd

edition. New York Botanical Garden Press.


20

Jackson, Scott. 1995. Delineating Bordering Vegetated Wetlands Under the Massachusetts

Wetlands Protection Act: A Handbook. MA Department of Environmental Protection.

Boston, MA. 89 + 5 pp.

Karr, J.R. and E.W. Chu. 1999. Restoring Life in Running Waters: Better Biological

Monitoring. Island Press, Washington, DC.

McGarigal, K., B.W. Compton, S.D. Jackson, K. Rolih, and E. Ene. 2005. Conservation

Assessment Prioritization System (CAPS) Highland Communities Initiative PHASE 1.

Final Report. Landscape Ecology Program, Department of Natural Resources

Conservation, University of Massachusetts, Amherst. URL:

http://www.umass.edu/landeco/research/caps/reports/caps_reports.html

Mitsch, W.J. and J.G. Gosselink. 1993. Wetlands. Van Nostrand Reinhold, Inc. New

York, NY.

Neckles, H.A. and M.Dionne, Editors. 2000. Regional standards to identify and evaluate

tidal wetland restoration in the Gulf of Maine. Wells National Estuarine Research

Reserve Technical Report, Wells, ME. URL:

http://www.pwrc.usgs.gov/resshow/neckles/gpac.htm

Pollock, L.W. 1998. A Practical Guide to the Marine Animals of North America. Rutgers

University Press, New Brunswick, New Jersey.

Scanlon. J. 2006. MassWildlife Landcover Mapping Decision Rules. Internal Document.

Massachusetts Division of Fisheries and Wildlife, Westborough, MA.

Somers, P. K. Lombard, and R. Kramer. 2006. A Guide to Invasive Plants in

Massachusetts. Massachusetts Division of Fisheries & Wildlife, Natural Heritage &

Endangered Species Program, Rabbit Hill Road, Westborough, MA 01581.

Swain, P.C. and J.B. Kearsley. 2001. Classification of the Natural Communities of

Massachusetts. DRAFT - 2001- (version 1.3) Reprinted 2004. Massachusetts Division of

Fisheries and Wildlife, Westborough, MA.

Tiner, R.W. 1987. A Field Guide to Coastal Wetland Plants of the Northeastern United

States. The University of Massachusetts Press, Amherst, MA.

U.S. EPA, 2002. Methods for Evaluating Wetland Condition: Developing an Invertebrate

Index of Biological Integrity for Wetlands. Office of Water, U.S. Environmental Protection

Agency, Washington , DC. EPA-822-R-02-019

Weiss, H.M. 1995. Marine Animals of Southern New England and New York: Identification

keys to common nearshore and shallow water macrofauna. Bulletin 15. State Geological and

Natural History of Connecticut, Department of Environmental Protection, Hartford, CT.

http://www.umass.edu/landeco/research/caps/reports/caps_reports.html

http://www.pwrc.usgs.gov/resshow/neckles/gpac.htm

Salt Marsh QAPP: Appendix C Version 2.2, May 2012 DRAFT

CAPS/SLAM: Salt Marsh Assessment Inner Marsh

Version 2.2 - May 2012

Vegetation Site Form

Site ID (Bin-Index): Site Name/Town:

Bank condition (for length of plot):

Bank vegetation (for length of plot):

Comments:

Plot Diagram Sketch a diagram of the sample plot. Include the following sampling information: baseline length (in meters); transect IDs; transect

lengths (in meters); location of origin sample point; and locations of macroinvertebrate sampling stations if they are flagged for

future sampling. Also, sketch marsh features, including creeks, marsh zones/dominant plant communities, and invasive species in

and around the plot. In addition to the plot diagram, sketch the location of the sample point and all transects on the Site Map (aerial

photo).


Vegetation Form: Site ID ______ Transect ID______

Record the transect number, lat/long, length, compass bearing (circle M for magnetic north or T for true north), distance from sample point, and

point interval. Walk along the transect and record the presence of plants touching the bayonet at each interval. Use the table below to mark a

tic in each cell for intervals 1 に 11 where a plant is present. Use the blank cells at the end of the table to record the genus and species for plants

not listed. Additional plant species are listed on the next page.

Date: Time: Evaluators:

Lat/Long (ground condition; DD; WGS84)

Transect 1 Transect 2 Transect 3

N N N

W W W

Transect length (m): Distance from center Sample Point (m):

Transect compass bearing (°): M / T Point interval ( = length/10) (m):

Genus species 1 2 3 4 5 6 7 8 9 10 11

Distichlis spicata

Iva frutescens

Juncus gerardii

Limonium nashii

Phragmites australis

Salicornia europaea

Salicornia virginica

Schoenoplectus* americanus

Schoenoplectus* pungens

Schoenoplectus* robustus

Solidago sempirvirens

Spartina alterniflora - Short

Spartina alterniflora - Tall

Spartina patens

Suaeda linearis

Typha angustifolia

Typha latifolia

AQUATIC ALGAL MAT

BARE SOIL

DITCH

MARSH DIEBACK

ROCK

WRACK

* Formerly Scirpus


Genus species 1 2 3 4 5 6 7 8 9 10 11

Agalinis maritima

Agropyron pungens

Agrostis stolonifera

Amaranthus cannabinus

Aster subulatus

Aster tenuifolius

Atriplex patula

Baccharis halimifolia

Carex paleacea

Carex salina

Euthamia graminifolia

Festuca rubra

Glaux maritima

Juncus balticus

Myrica gale

Onoclea sensibilis

Panicum virgatum

Plantago maritima

Pluchea purpurascens

Potentilla anserina

Rosa rugosa

Salsola kali

Spartina cynosuroides

Spartina pectinata

Toxicodendron radicans

Triglochin maritimum

Triglochin palustre

Comments


Habitat Complexity Form: Site ID______ Transect ID______

After completing the Vegetation Form, walk back along the transect towards the water feature. Use the tape measure to observe

and record the さゲデ;ヴデざ locations (distance in meters) and types of habitat transitions along the transect. Do not record transitions for

habitats that measure less than one meter along the transect. Habitats are defined as heterogeneous plant communities (e.g.

Spartina patens / Distchlis spicata mix) and hydrogeomorphic features (e.g., panne, pool, and creek, plus ditch) associated with the

marsh. Both a plant community and hydrogeomorphic feature can be recorded at the same location (e.g. Spartina alterniflora-short

and panne). Record the tape measure reading for each habitat transition to the nearest tenth of a meter in the appropriate box(s) (e.g. 5.5 m).

Separate recorded habitat transitions with a comma if that habitat occurs more than once on a transect. List no more than three

plant species when describing a community. Record species in order of dominance, with the first species being the most dominant.

For example, record a habitat patch that includes S. patens (75%), J. gerardii (25%), and P. maritimum (< 1%) as S. patens / J. gerardii

mix. Record start and end distances (range) for all hydrogeomorphic features (e.g. Panne = 25-30 m). Include patches and features

that are offset up to 1 m from the transect. Complete a Habitat Complexity Form for each transect. See the next page for a marsh

zonation reference diagram (Figure 1).

Plant Community Transition Distance (m) Hydrogeomorphic Feature Transition Distance (m)

Distichlis spicata Creek

Juncus gerardii Ditch

Spartina alterniflora-short Panne

Spartina alterniflora-tall Pool

Spartina patens

S. alterniflora-short / S. patens mix

S. patens / S. alterniflora-short mix

S. patens / D. spicata mix

S. patens / J. gerardii mix

D. spicata / S. patens mix

MARSH DIEBACK


Comments

Figure 1. Plant zonation in

northeastern salt marshes.

This diagram shows the major

plant zones and dominant

species; see text for details and

Tiner (1987), Mitsch and

Gosselink (1993), or Bertness

(1999) for a description of salt

marsh vegetation patterns.

(Adapted from Carlisle, et al,

2002).


Human Disturbance Form: Site ID______

Walk the perimeter of the sample plot and describe in the table below the types, extents, and severity of disturbances within a

100 m buffer of the sample point (200 m diameter circle depicted on the site map). Disturbance types are listed below. Use the

bracketed letters preceding each disturbance type to mark locations of disturbances on the provided site map. Also record any

of these indicators of disturbance when encountered while implementing other elements of the SOP.

[A] Water control structures (culvert, tide gate, dam, weir, storm water input, fill (road/railroad), ditching, channelization,

and other human activity affecting the hydrology of the site

[B] Soil disturbance (filling, sedimentation, haying)

[C] Obvious spills

[D] Direct point or non-point source discharge from agricultural operations, septic or sewage treatment systems, or storm

water affecting water quality of the site

[E] Walking trails, horse trails, and roads (excluding wildlife trails)

[F] Small-craft boating (presence of docks, moorings, or observed boating) and boat storage

[G] Presence of trash/litter

[H] Presence of garbage dumping

Type Presence Description of type, areal extent, and severity of impact

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Development of a Comprehensive State Monitoring and Assessment Program for Wetlands in Massachusetts

Appendix H Revised December 30, 2009

Standard Operating Procedures: Assessment of Wetland

Communities

Phase 2c: 2009

Prepared by:

Theresa Portante, Wetland Field Manager Kasey Rolih, Wetland Field Manager

Brad Compton, Computer Data QA Manager and

Scott Jackson, UMass QA Manager

Department of Natural Resources Conservation, Holdsworth Hall University of Massachusetts, Amherst, MA 01003

Standard Operating Procedures: Assessment of Forested Wetland Communities December 30, 2009

Standard Operating Procedures: Assessment of Wetland Communities

1. Scope and Application This SOP establishes a standard set of procedures to be followed for data collection toward the development of a Site Level Assessment Method (SLAM) for MA freshwater forested wetlands and to validate/calibrate the Conservation Assessment and Prioritization System (CAPS) as a mechanism for a landscape level analysis (Level 1) of ecological integrity. This project will focus on assessment of wetland biological community condition in forested wetlands. Described below are the procedures that will be followed in collecting data on algae, macroinvertebrates, vascular plants, bryophytes, epiphytic macrolichens and habitat characterization (e.g. water chemistry, hydroperiod, etc.) to serve as a basis for development of a SLAM, which will incorporate the use of Indices of Biological Integrity, for freshwater forested wetlands. 2. Summary This SOP is applicable for freshwater deciduous/coniferous forested wetlands that have the hydrogeomorphic (HGM) classification of a slope or flat throughout Massachusetts (hereafter referred to as forested wetland). Data collection for phase 2c will focus on forested wetland communities in the Miller’s and Concord (Sudbury-Assabet-Concord) Watersheds, however this SOP can be applied to all forested wetland communities. Sampling sites will be selected via a stratified random process. Field data collection will involve sampling of several biotic communities to determine if 1) there is a dose-dependent response in various attributes of the biological community to stressors within the landscape and 2) to validate/calibrate the ecological integrity metrics that are utilized in the CAPS model. Characterization of the wetland and assessment of its biological condition will be conducted in the field by assessing habitat, algae, macroinvertebrates, vascular plants, bryophytes, epiphytic macrolichens and habitat characterization. 3. Safety Considerations

• Fieldwork will not be conducted during heavy rain events or unsafe conditions such as electrical storms or high wind events. Practice “safety first”.

• If there is no safe access to a plot point, the field sampling will not be conducted for that site.

• Private property will be respected using the following guidelines.

o If property is in close proximity to buildings or other heavily used areas, landowner permission will be sought

o Posted property will not be accessed without permission of the landowner


o Otherwise, sampling will proceed without any special effort to gain landowner permission

o If asked to leave private property by the landowner, samplers will

discontinue work and leave.

• Each field technician will carry a personal first aid kit and a wilderness first aid guide

• Field personnel will not access sites alone without the instruction of a field manager

• No chemicals (other than ethanol) will be handled by personnel in the field

4. Sample Collection, Preservation, and Handling Macroinvertebrates collected using the stovepipe sampler will be preserved in 95% ethyl alcohol solution. 70% ethanol will be used to preserve macroinvertebrates collected in the emergence traps. Macroinvertebrates collected in the pitfall traps will be preserved initially in a 50:50 propylene glycol/water solution and a drop of dishwashing liquid soap. The samples will be rinsed with tap water in the lab and transferred to a 70% ethyl alcohol solution. Samples will be labeled with the plot ID, date, surveyor, and collection method. They will be sorted and identified to order in the lab. Samples will be preserved and held in the lab until resources are available to identify the macroinvertbrates to genus and species (if possible). Earthworms will be collected into 70% isopropyl alcohol and kept cool until transfer to the lab for permanent preservation in 10% formalin. Samples will be labeled in the field with plot ID, data, and name of surveyor. Transfer of worms into formalin will occur in a fume hood using safety glasses and gloves. Worms will remain in formalin for at least 24 hours before being permanently stored in 70% isopropyl alcohol. Tentative species IDs and counts may be made in the field. Official counts and IDs will be made in the lab using a dissecting microscope. Earthworm species identifications will follow Schwert (1990) and Reynolds (1977). Algae will be collected and labeled with the plot ID, date, surveyor, and collection method. Algae samples will be preserved with M3 fixative (Potassium Iodide, Iodine (optional), glacial acetic acid, formalin) and stored until resources are available to identify them to genus and species. Vascular plant, bryophyte and lichen collections will be limited to species that cannot be identified in the field. For species that cannot be positively identified in the field samples will be collected for lab identification and photographed for digital preservation. Taxonomic identification at the species level (preferred) or genus level (if species


identification is not possible) will be achieved in the laboratory through the use of field guides, technical keys, and reference to regional herbaria housed at research universities such as UMass. Samples will be labeled in the field with the plant ID (e.g., “unknown sedge #1”) site location, date, and person who collected the sample, and assigned a code in the laboratory for use in digital preservation. 5. Equipment/Apparatus Before leaving for the field the Field Manager will confirm the following equipment is available:

Backpack sprayer Beaker Bleach solution (1/2 cup bleach per gallon tap water) Clipboard Compasses Cooler with ice Data sheets Deionized water Digital camera w/extra batteries Dip net, small, 500 micron mesh Dishwashing soap solution Emergence traps Ethanol (95%, 70%) Field notebook Flagging Forceps GPS (Global Positioning System) Hand lens Hanna ph/conductivity meter Hip chain HOBO Pendant Temperature/Light Data Logger iButtons Isopropyl alcohol Labels for algae samples Labels for earthworm samples Labels for macroinvertebrate samples Labels for vascular plant, bryophyte & lichen samples Lids, closed Liquid dish soap or hand soap (phosphate-free and biodegradable) Location maps Meter stick Meter tape M3 preservative Nalgene bottle (500ml) Palm Tungsten E2 Handheld (PDA) Pencils


Permanent markers pH/CON 10 pH/Conductivity/Co Meter Plastic collecting bags Plastic cups Plastic containers (32 oz and 16 oz) Plastic amber bottles (100 ml-250 ml) PVC pipe (2 ½” diameter) Rite-in-rain paper and pen Scissors or jack knife Screens Stakes String Soil auger SOP Spoonulet Squirt bottle Standard solutions for calibration of pH/Conductivity/Temp meter Stovepipe sampler Tap water Trowel or bulb planter Turkey baster (large Pipette) Vials Water/detergent solution White bowl

6. Reagents

Bleach solution (1/2 cup bleach per gallon tap water) Deionized water Ethanol Formalin solution (10%) * Glacial acetic acid * Isopropyl alcohol Liquid dish soap or hand soap (phosphate-free and biodegradable) Potassium Iodide * Propylene glycol/water solution Standard solutions for calibration of pH/Conductivity/Temp meter Tap water

* M3 solution 7. Calibration & Training Equipment calibration procedures for the GPS units, Oakton pH/CON 10 pH/Conductivity/Co Meter, Hanna portable ph/EC/TDS/Temperature Meter, Thermocron ibutton, and HOBO Pendant Temperature/Light Logger will be done according to the manufacturers’ recommendations. See section 2.6 of the QAPP for details.


Field crew members will have sufficient previous training and experience to reliably conduct field data collection or they will receive training from the UMass QA Manager and/or other project scientists with relevant expertise. The QA Manager will ensure that all field crew members receive specific training on macroinvertebrate sample sorting and identification (to order), plant identification, and delineation of a Bordering Vegetated Wetland. All Field Managers and Field Scientists will receive training from the QA Manager on appropriate QA/QC procedures. 8.0 Procedures Sampling will occur between May 11 and September 30, to ensure adequate assessment of the targeted wetland biotic communities. Forested wetlands in the Millers and Concord Watersheds will be identified using the MassDEP Wetlands Mapping data (1:12,000 based on photography from 1993 and 1999). Sample locations will be randomly stratified across deciles of buffer zone insults (one of the landscape metrics used in CAPS) and deciles of ecological integrity (results from CAPS analysis) from the CAPS assessment of 2009. This will create 100 buffer zone insults x IEI bins. Up to five random points that fall within deciduous or mixed forested wetlands (as depicted in MassDEP wetlands; 1:12,000 based on photography from 1993 and 1999) will be selected for each bin. Samples within 100 m of a fourth order or larger stream will be excluded to avoid areas that might potentially be floodplain forests. All points will be separated by at least 500 meters. The 150 (75 in each watershed) sampling plots will be selected randomly from among the 100 bins. Within each bin, potential plots are ordered. If a plot needs to be dropped, the next-higher plot in the same bin will be used. Note that some bins will have fewer than five points or may be entirely empty because some combinations of IEI and wetland buffer insults are rare or absent in the landscape. A random identifier will be assigned to each bin to obscure the IEI/wetland buffer insults class that each bin represents. Field personnel will not have access to the original classes, thus sampling will be blind with respect to CAPS predictions. Plots will be compared to aerial photographs (1:5000, 2005 Color Orthophotos available from MassGIS) and GIS data for hydrography (MassGIS, 2005), Potential Vernal Pools (NHESP, 2000) and Certified Vernal Pools (NHESP, 2008). Plots that fall within 30 m of potential or certified vernal pools, dominated by conifers, or fall within 30 m of a 3rd order stream or greater will be dropped. Areas in close proximity to vernal pools and larger (> 2nd order) streams will be dropped to avoid sampling invertebrates too close to areas characterized by longer hydroperiods than our target wetland community. Likewise, areas dominated by conifers will be avoided because they do not match the target wetland community (freshwater deciduous/coniferous forested wetlands that have the hydrogeomorphic (HGM) classification of a slope or flat).


GPS navigation will be used to locate each wetland plot. GPS precision must be 10 m or less and the navigator will stop and establish the plot once the distance to plot center is 0m. In the case of GPS interference from tree-canopy or atmospheric effects two procedures may be followed. The first is to wait 10 minutes for satellite reception to improve. If a dense forest canopy appears to be the problem use triangulation to locate the plot. We will approach the plot from three different locations where the canopy is mainly deciduous. Using compass and distance measurements provided by the GPS (precision must be 10 m or less), the plot will be located. It will not be necessary to hit the plot exactly (since it's randomly selected) it just needs to be selected without bias. However, a reasonably precise GPS point is needed of where the plot actually ends up. The strategy is (1) do the best we can when locating the plot and (2) take a precise location (precision ≤ 10 m RMS) once the plot has been established. Field workers will be on the plot for 2-3 hours and will be able to keep trying until they get good GPS coverage. 8.1 Establishing Sampling Area A 30 m radius plot will be used to sample the wetland point (Figure 1). A reserved 5 m radius area will be established in the center of the plot. Eight 25 m transects will be run from plot center at 0o, 45 o, 90o, 135 o, 180o, 225 o, 270o, and 315 o compass bearings. Vascular plants and bryophytes will be surveyed on transects run at, 45 o,135 o,225 o, and 315 o. Plant transects (transects 2, 4, 6, 8) and bryophyte plots will be denoted to prevent trampling, by flagging the transects and marking them on the Plot Information A form (Appendix L). The plot will be subdivided into 4 quarters, A-D.. They will be established in a clockwise direction beginning with transect 1 (Quarter A between the N and E transect, etc.)


Figure 1.

Diagram of sampling area. Eight 25 m transects run at 0o, 45 o, 90o, 135 o, 180o, 225 o, 270o, and 315 o compass bearings. The location for all samples (algae, water chemistry, etc.) will be noted on the plot diagram. A sampling point will be moved if any of the following conditions are encountered.

o The dominant tree cover in the plot area is <30% as determined by visual estimation

N

Plot Diagram 2009 – Forested Wetlands 1

2

3

4

5

6

7

8

A

BC

D

Transects for Macroinvertebrates, Algae and Hydro Profile (30m)

Bryophyte plots (0.5m2)

Plant transect (25m)

Reserved area 5m radius

Macroinvert sample point

A Quarters


o Any transect length is <15 m, as may occur in narrow wetlands (e.g. fingerlike projections, narrow bands of wetland along streams)

o Plot area is inundated due to beaver dams o Point falls within 30 m of a mapped 3rd order stream (or larger)

The sampling point will be moved to the nearest location that does not violate the previously stated conditions, but no greater than 30 m away. If a suitable sampling point cannot be found within 30m of the original point the site will be dropped and another sampling point from the same bin selected. 8.2 Overview of Wetland Biotic Community and Habitat Assessment Each point will be sampled for algae, macroinvertebrates, vascular plants, bryophytes and epiphytic macrolichens. Samples will be taken within a 30 m radius plot. Samples will be analyzed to determine if the attributes of the biotic communities show a dose-dependent response to anthropogenic stressors in the landscape as measured by CAPS metrics. In addition a habitat assessment will be conducted to characterize the assessment area. A detailed description of the plot (includes hydrology, anthropogenic disturbance, etc.) will be recorded in a field notebook by each surveyor. Data will be recorded with a PDA and paper forms. Tungsten E2 Handheld PDAs will be used to record vegetation, bryophyte and lichen data in the field. Paper data sheets will also be completed to serve as backups. Data from the PDAs will be downloaded to the master database on a daily basis. 8.2.1 Habitat Assessment (a) Topographic complexity

Topographic complexity will be determined to assist in the characterization of the wetland. Each odd numbered transect will be walked to observe and record variations in slope/elevation. From the center point of the plot walk four 30 m transects and count the number of micro-topographic depressions (“pits”) at least 1 m2 in size encountered along each transect. Counts will be recorded on a data sheet Topographic Complexity form (Appendix L) Depressions will only be counted if they are sufficiently obvious that they could be recognized even if groundcover vegetation is dense. If a pit is divided along the transect line by a mound it will be counted as two separate pits. A mound is defined as ≥ 15cm in height relative to the base of a pit and has the development of soil. Vegetation (e.g. tussock sedge) will not count as a mound. Topographic complexity will be expressed as the number of micro-topographic depressions per 100 m of transect length.

(b) Hydrology


Hydroperiod A HOBO Pendant temperature/light data logger will be placed in the water for the duration of the study period (about 4 months) to determine the relative hydroperiod of the wetland surface water. The HOBO will record temperature at two hour intervals. Place the data logger in a location within the plot that is judged by the field manager likely to remain inundated longest whether or not there is any standing water at the time. Place the logger inside a plastic white container to protect it from direct sunlight. Holes will be drilled into the sides of the cup to allow water to flow through. The cup will be held flush to the surface of the ground with a plant stake with a metal ring at the top to keep the cup from moving. Label the container with the serial number of the HOBO. Measure the depth of the water where the HOBO is placed at each plot visit (7 measurements) and record on the Hydrological Characterization form (Appendix L). An ibutton will be hung against the North side of the closest tree to the location of the HOBO. The ibutton will record ambient air temperature every two hours in sync with the HOBO. The ibutton will also be protected by direct sunlight with a white plastic container and holes will be drilled to allow air passage. Label the container with the serial number of the ibutton. Record the placement location and the serial number of the loggers on the Plot Information A form. Collect data loggers upon the completion of the biotic community assessment.

The temperature data from the loggers will be uploaded following procedures according the manufacturer’s instructions (See QAPP Appendix J). The temperature data will be used to determine the relative hydroperiod (i.e. the duration of the sampling period).The coefficient of variation (CV) in temperature for each 24 hour period will be calculated for both the ambient air temperature (AAT) and water temperature (WT). The assumption is the ratio of AAT(CV)/WT(CV) will approach 1 as the depth of the water decreases. This will be verified with the recorded water depth of the HOBO location (recorded at 7 dates throughout the sampling period). This relationship will be used to estimate the depth of water for each day based on the temperature data. This data will be used to characterize the relative hydroperiod of surface water for each plot (method to be determined).

Hydologic Profile/Characterization A hydrologic profile along odd numbered transects will be taken using a point intercept method each time a site is visited (eg. trap deployment, trap collection, etc.) The profile will be used to characterize the surface hydrology during the field season.


At the first site visit, odd numbered transects will be flagged every 5m. At each 5m point intercept along the transect, the presence of saturated soil, surface water (>2.5cm), or dry surface will be recorded on the Hydrologic Characterization form. The percent cover of each category will be determined for each visit and for the duration of the field season. Hydrologic features such as a single channel or braided stream channel that is located in the plot will be described (direction of flow, etc.) and recorded on the Plot Information A form. Groundwater

Groundwater will be monitored using shallow groundwater monitoring wells to determine the fluctuation in the water table throughout the field season. Readings will only be taken 6 or 7 times and will not be monitored daily. This information will provide information to characterize the influence of groundwater to the wetland point. A PVC pipe, 1.2 m in length and 6.35 cm in diameter, will be installed to monitor groundwater (Fig. 2). A single pipe will be installed at the lowest point in the wetland, based on topography and depth of surface water. This will be determined after setting up the hydrologic profile transects and walking around the plot. The hole for the pipe will be dug using a soil auger. 0.90 m will be placed below the surface. Slits will be cut every 4.8 cm along the length of the pipe on each side through about a quarter of the pipe. The slits will allow the passage of water while preventing the soil from entering the pipe. The bottom of the pipe will be capped with a water tight seal. A 4.8 cm diameter cap will cover the top of the pipe for ease of removal to take water measurements. A meter stick lined with chalk will be used to measure the depth to groundwater. First determine the measuring point (MP) by measuring the length of the pipe above the surface. Insert the meter stick lined with chalk above the well and record when it crosses into the pipe (held value). Remove the stick and note where the chalk is wet (wet value). To determine the depth to groundwater first subtract the wet value from the held value to determine the water level below MP. Then subtract MP to determine the level below the land surface. (/personal correspondence/, R. S. Socolow, USGS) Measurements will be taken each time the site is visited. The data will be recorded on Hydro Profile form.


Figure 2. Groundwater well measurements.

(c) Water geochemistry

Conductivity, temperature and pH will be measured for surface water (if present) using a portable pH/Conductivity meter at 4 locations in the plot. Take readings from surface water closest to the midpoint of each of the odd numbered transects running in cardinal directions (location of algae samples). If there is no standing water present along a transect move in a clockwise direction to find the closest area with standing water. If there is no standing water present within the quarter plot keep moving clockwise until readings are collected from four locations within the plot. The minimum distance between readings must be 3 m. Take a reading from any major stream channel in the plot if present. Note on the Plot Information A form the transects and/or quarters from which readings were taken. Record pH, conductivity, and temperature on the Plot Information B form.

(d) Human disturbance


Visual observations of human disturbance to the wetland will be noted. Surveyors will note the following activities in the field notebook, describing the type and extent of each disturbance. Walk the four odd numbered transects running in cardinal directions and record in the field notebook the type and extent of disturbance for each of the following.

• Water control structures (culvert, dam, weir, storm water input, fill (road/railroad), ditching, channelization, beaver dam, and other human activity affecting the hydrology of the site

• Soil disturbance (filling, plowing, grading, grazing, dredging, sedimentation, vehicle use.

• Obvious spills. • Direct point or nonpoint source discharge from agricultural operations, septic

or sewage treatment systems, or storm water affecting water quality of the site • Walking trails, horse trails, logging roads, ATV trails, old cart paths, and

roads (excluding wildlife trails) • Evidence of mowing, burning, or timber harvesting. • Presence of trash/litter. • Presence of garbage dumping.

Also record any of these indicators of disturbance when encountered while implementing other elements of the SOP.

(e) Soils A soil pit will be used to characterize the soil for each plot. Select a location for the soil pit within 5 m of the groundwater well and 1 m distant from tree stems, animal holes, or other disturbances. Using a spade dig a soil pit 12 inches in diameter to a minimum depth of 16 inches; increase depth if more information is needed to characterize the soil. Dig a second soil pit for plots lacking uniform topography, where a change in the soil may be present. Conduct work only when the light allows for accurate color classification of the soil and its features. Remove a clean slice of soil from the soil pit. If saturated conditions prevent a pit from being dug use an auger to sample the soil and collect the necessary data. Turn the auger no more than 4 times so that the core is not mixed and an accurate profile can be documented. Repeat this step until the profile reaches a depth of at least 16 inches. Record on the Soils data form a description of the soil profile, including soil horizons, redoximorphic features and the associated colors. A Munsell Soil Color Chart (Munsell 2000) will be used as a guideline when describing the color and redoximorphic features of the soil pit. Soil Taxonomy, tenth edition (USDA 2006) will be used to define the soil horizons. Document on the data form additional


information useful for classification, such as hydric indicators (USDA 2002), texture, depth to groundwater, stoniness, and slope. This information will be analyzed in order to classify the soil using Keys to Soil Taxonomy, tenth edition.

8.2.2 Protocols for Sampling Biotic Communities 8.2.2.1 Algae

Algae will be sampled as a indicator of water quality, community composition, and ecosystem health. Algae are an integral component to the wetland community and are a primary food source to many macroinvertebrates. Samples will be collected in June before water draw down occurs. Four samples, each 50 ml, will be collected from each microhabitat within the wetland (benthic, including leaf litter and surface sediments, and surface water) for a total of 12 samples per site. Algae samples will be preserved in M3 fixative (Potassium Iodide, Iodine (optional), glacial acetic acid, 25% formalin). One ml of M3 will be added per 50 ml sample. All algae samples will be recorded on the algae sample login form before storage in the lab. Protocols for sampling algae were adapted from Danielson, 2006, Hawkins et al., 2003, and Vermont DEP, 2003.

(a) Benthic algae

Leaf litter samples will be collected. Leaf litter will be collected from areas within the plot with surface water present. In the absence of surface water, leaf litter will be collected from wet depressions. Collect leaf litter from areas of standing water closest to the midpoint of odd numbered transects If there is no standing water present along a transect move in a clockwise direction to find the closest suitable sampling location within the quarter plot. If standing water is lacking within a quarter plot collect leaves from a wet depression closest to the midpoint of the transect. If there are no suitable locations (surface water or wet depressions) present within a quarter keep moving through the plot until four samples have been collected. The minimum distance that samples must be spaced is 3 m. Note on the Plot Information A form the transects and/or quarters from which samples were taken and a description of the sampling location. Record the depth of the surface water if present on the Plot Information B form. From each sampling location collect red maple leaves to cover the bottom of a small bowl (10.5 cm2). Scrape the leaf surfaces using a metal spoonulet to scrape off the algae. If red maple leaves are not available collect other deciduous leaves of similar size and make a note of the species used. Rinse each leaf with DI water after scraping. Collect all scrapings from the small bowl into a 50 ml vial. Keep rinsing the pan with DI water until there is 50ml in the vial. Add 1ml of M3 per 50ml of benthic leaf scrapings for preservation. Clean the pan and spoonula with tap water after sampling.


(b) Water grab sample (adapted from ME DEP)

Water samples will be collected to sample algae. Take samples from surface water closest to the midpoint of the four odd numbered transects. If there is no standing water present along a transect move in a clockwise direction to find the closest suitable sampling location. If there is no suitable location present within the quarter plot keep moving clockwise until samples are collected from four locations within the plot. The minimum distance between samples must be 3 m. Note on the Plot Information A form the transects and/or quarters from which samples were taken. Record the depth of the surface water on the Plot Information B form Use a clean and dry 50 ml vial to collect sample. Submerge the water sampler to collect the surface water taking care to minimize the collection of organic material. Water samples will not be collected in areas where the leaf litter must be depressed in order to collect a sample. Add 1ml of M3 per 50ml of the water sample for preservation. Repeat for each transect.

(c) Surface substrate sampling

Surface substrate samples will be collected to sample algae. Using a turkey baster (large pipette) collect a 50 ml sample of the surface substrate from areas with surface water at the same location as leaf samples (see (a) above). To collect the sample, stick the end of the baster into the substrate and suck up a sample from the surface. If necessary, loosen up the substrate by moving around the tip of the baster before taking a sample. Pour the 50 ml sample into a 50 ml vial. Add 1ml of M3 per 50ml of the water sample for preservation. Note on the Plot Information A form the transects and/or quarters from which samples were taken. Repeat for each transect. Record the depth of the surface water if present on the Plot Information B form. Clean the turkey baster with deionized water after sampling.

8.2.2.2 Macroinvertebrates Macroinvertebrates are will be sampled as an indicator of water quality and community composition, and ecosystem health. Macroinvertebrates will be sampled from June-August. Stovepipe sampler and emergence traps will be used in June; pitfall traps to collect epigeal macroinvertebrates and soil pits to collect earthworms will be conducted from July-August.

(a) Earthworms


Earthworms will be sampled in forested wetlands from August through November using a combination of liquid extraction and midden counts (Lawrence and Bowers 2002, Hale et al 2005): For midden counts place 1m2 sampling frame on soil surface at 15m along each odd-numbered transect and count number of middens inside the frame. Establish one earthworm sampling plots at the most suitable location (not standing water) within the assessment area. Place sampling frame (11’ diameter or 613 cm2) on top of soil and carefully remove any vegetation from within frame. Collect any earthworms found on soil surface or in vegetation and place in small plastic sampling tray with lid. Count number of juveniles, adults, and middens within the plot. Push sampling frame into soil. Pour ½ gallon liquid mustard solution into sample area and begin collecting worms as they surface. Wait three minutes before pouring remaining ½ gallon into soil. Liquid extraction sampling time for each plot is 10 minutes. Earthworms encountered during the excavation pit traps or soil pits will be collected and preserved. Kill all worms in 70% isopropyl alcohol. Place worms into alcohol-filled vial labeled with plot ID, subplot ID, and date, and collector’s name. Keep earthworms cool until transfer into 10% formalin solution for permanent preservation at the end of the field day.

(b) Aquatic macroinvertebrates: Stovepipe sampler (adapted from ME DEP)

Macroinvertebrates will be collected using a stovepipe sampler (5 gallon plastic bucket with the bottom cut off). Collections will be made in two locations dispersed within the plot where surface water and/or wet depressions are present. Samples will be taken from two locations within the plot where surface water is most suitable for sampling based on water depth and areal extent of inundation. If surface water is not present within the plot, sample in locations (depressions) with the wettest substrate. If possible locate the sampling locations in diagonal quarters of the plot (e.g. quarters 1 & 3 or quarters 2 & 4). If suitable sampling conditions are not present in diagonal quarters try to use sampling locations in each of two adjacent quarters. If necessary place both sampling locations in the same quarter. The minimum distance between samples must be 3 m. Note on the Plot Information A form the transects and/or quarters from which samples were taken. At each sampling location place the stovepipe sampler firmly into the substrate (few cm deep) and hold it in place. Agitate the water in the sampler for 10 seconds to dislodge organisms from the substrate and vegetation. If surface water (>1.27 cm) is present take five sweeps within the sampler with a 500 micron mesh hand net (10.5x12.5 cm). After each sweep, transfer all material into a 32 oz


collecting jar. Inspect the net, remove any clinging organisms and add them to the sample. The jar should only be filled halfway with sample material and additional jars may be used if necessary. Fill container with 95% ethanol. Record depth of surface water on the Plot Information B form. For wet depressions (with little or no standing water) collect three, one-hand leaf litter grab samples from within the stovepipe. Distribute grabs evenly throughout the stovepipe area. Preserve the sample the same as for the dipnet samples. Record on the Plot Information B form. Label containers with site ID, date of collection, surveyor ID, and description of microhabitat. Samples will be strained and preserved with fresh ethanol within four months of collection. Containers will be stored in the lab for up to five years until they are processed.

(c) Insects: Emergence Traps

Four emergence traps per plot will be set and collected after 7 days. Emergence traps will be set on the water surface or on the surface of the soil in the wettest depressions in the absence of surface water. Site selection for trap placement will follow the protocol previously described for benthic algae, but will be placed 1m apart from areas that were disturbed while sampling for algae or using the stovepipe sampler. Set emergence traps in areas of standing water closest to the midpoint of each transect. If there is no standing water present along a transect move in a clockwise direction to find the closest suitable sampling location within the quarter plot. If standing water is lacking within a quarter plot set the trap in a wet depression closest to the midpoint of the transect. If there are no suitable locations (surface water or wet depressions) present within a quarter keep moving through the plot until four trap locations are selected. The minimum distance that samples must be spaced is 3 m. Note on the Plot Information A form the transects and/or quarters where emergence traps are set. Fill a jar (with funnel top) with 70% ethanol and place it upside down at the top of the emergence trap to collect emerging insects. Tie the traps with string to nearby vegetation or with stakes to prevent drifting. Make sure that there is enough slack in the string to ensure the trap will stay flush with the water surface if draw down or flooding occurs. Upon collection of the traps replace the jar lids with fully enclosed lids and add ethanol as needed. Samples will be kept separately. Label jars with site ID, start and end date of collection, surveyor ID, and description of microhabitat. If surface water is present record the depth at the time of placement and collection on the Emergence Trap Log form (Appendix L). In addition, record the setter and collector ID, microhabitat, condition of the trap, and the amount of ethanol in the jar when collected. Jars will be stored in the lab for up to five years until processed.

(d) Epigeal macroinvertebrates


Pitfall traps will be set out in July to collect epigeal macroinvertebrates. Traps will be 16 oz clear cups placed in the ground with the top of the cup flush with the ground surface. Cups will be filled with ~150ml of a 50:50 propylene glycol/water solution and a drop of dishwashing soap. A small screen made of hardware cloth (1x1 cm squares) will be placed inside the cups to prevent small vertebrates from entering the killing solution. A plastic plate held up with small stakes will be placed over the pitfall trap to serve as a roof. Place eight pitfall traps, 2 on each transect at 10 and 15m. Place traps in areas where the chance of flooding by surface water (avoid pits) is reduced. Collect the contents of pitfall traps after 7 days. If the trap is >1/2 full of water it will be discarded. Each trap will be collected separately in a small container. Record the setter and collector ID, microhabitat, amount of water in the trap, and the condition on the Pitfall Trap Log (Appendix L). The samples will be rinsed with tap water in the lab (to remove the soap) and 70% ethanol will be added. Label jars with site ID and start and end date of collection. Samples will be stored for up to five years in the lab until they are processed.

8.2.2.3 Vascular plants Vascular plant data will be collected as an indicator of community composition and species diversity (proportion of native to invasive), will contribute to the understanding of the status of species of conservation concern (rare, endangered, or invasive), and provide useful information on potential threats to natural systems. Invasive plants named as such in this assessment are those currently regulated by the Commonwealth of Massachusetts (Somers et al 2006). Data collection will occur throughout the field season, June – September 2008.

a. Estimate species abundance of all vascular plants in a 30 m radius plot using a point

intercept method. Estimate percent cover as the proportion of the line directly intercepted by each species by vertical projection on four 25 m transects (excluding reserved area) placed in the four directions (even numbered transects). Tally each plant species that touches the transect line or is intercepted by a vertical projection from forest floor to canopy every 1m along the transect. Record tallies every 5 m to ensure an accurate count.

b. Following transect sampling conduct a 20-minute walk around (within) the entire plot and list species not encountered on transects. Assign these additional species a percent cover class of <1%. Record data on the vascular plant data form.

c. Estimate basal area using a wedge prism (10 or 15-factor). Stand near plot center, hold prism over plot center, view trees through prism at breast height (1.4 m) and tally trees, moving in a full circle starting north. List the species of each tallied tree.

d. Assign a forested landcover class according to MassWildlife Landcover Mapping Decision Rules (March 1996) and a natural community type according to the


Massachusetts Natural Heritage & Endangered Species Program (Swain & Kearsley 1999).

e. Collect unknown species for lab identification under dissecting scope. Place each species in a separate collecting bag labeled with plant ID (e.g., “Unknown #1, etc.), plot ID and date. Take digital photographs on site as needed. List PhotoID # next to unknown plant ID on the vascular plant form.

f. Refer to resources on regional flora if necessary (Gleason & Cronquist 1991, Magee & Ahles 1999). Assistance from the herbaria and staff at the UMass herbarium will be requested as needed.

8.2.2.4 Epiphytic macrolichens Epiphytic macrolichen data will be collected as an indicator of forest health, community composition, and species diversity. Stand at center of established 30 m radius plot. Starting due north, use a 10 or 15-factor prism to select trees for lichen sampling. Identify and estimate percent cover for macro-lichens on all trees and shrubs with a diameter at breast height (dbh) of four inches or greater. Estimate percent cover on the trunk in the area between from base of tree up to 2m from base. On the Epiphytic Macrolichens form number and list each tree, record the tree species and dbh, and list macrolichen species present. Estimate percent cover for each macro-lichen species using the following cover classes: 0.1=<1%, 1=1-5%, 2=6-25%, 3=26-50%, 4=50-75%, 5=>75%. Collect samples as needed into paper herbarium packets labeled with plot ID, date, collector, and sample number. Mark any samples collected with a “V” for voucher on the data sheet next to its tentative name or as “Unknown #1, Unknown #2, “ etc. Nomeclature will follow (Esslinger 2007).

8.2.2.5 Bryophytes Bryophytes have important roles in mineral cycling, water dynamics (some species may hold 10 times their weight in water), regulation of microclimate, and provide food and habitat to a host of invertebrates. Many are sensitive to human disturbance including forest management, and bryophytes may comprise a major component of the biomass and net productivity in wetland systems. Ground-dwelling moss and liverwort data will be collected on 4-0.5 m2 plots located in representative areas along the vascular plant sampling transects. Estimate percent cover for each bryophyte species in each quadrat using the following cover classes: 0.1=<1%, 1=1-5%, 2=6-25%, 3=26-50%, 4=50-75%, 5=>75%. Follow quadrat sampling with a 20-minute walk around the plot and list additional species not found in quadrats; species documented during the walk around will be assign a percent cover of 0.01%. Collect a voucher specimen in herbarium packets for each species found


across all study plots. Nomenclature for mosses follows Anderson (1990) and Anderson et al (1990), for liverworts follows Schuster (1974). 8.7 Protocol for Decontamination of Field Equipment

Inspect all equipment for debris before leaving a site. Dispose of debris in a trash bag or on dry, high ground. When possible, leave equipment to air dry and inspect to remove any remaining plant fragments. Spray equipment with a bleach solution, scrub, and rinse with tap water to remove any additional debris. Clean the pH/conductivity meter according to manufacturer’s recommendations.

9. Quality Control Compliance with procedures in this SOP will be maintained through monthly internal reviews. Personnel will conduct periodic self-checks by comparing their results with similarly trained personnel working on the project. See sections 2.5 and 2.6 of the QAPP for details about QA/QC measures. 10. Interferences Inclement weather (heavy rain) may interfere with our ability to collect representative data on a variety of parameters. Severe weather may delay field data collection due to safety concerns. Access may be a challenging aspect of data collection in more developed areas of the study area. Posted property or sites that are too difficult to access or unsafe to sample will be replaced with alternative sites from the same stratified sampling bin. 11. Preventative Maintenance Field equipment will be inspected by the UMass Field Manager each day before going out to collect field data. At the field site equipment will be tested prior to data collection to ensure that it is working properly. Equipment will be subject to regular maintenance as needed and as recommended by the manufacturer. GPS accuracy will be assessed once a month by a check of any units used in the field with a known location. See section 2.6 of the QAPP for more detail. 11. Corrective Actions Data quality control ensures high quality data, however we are prepared to re-measure any plots within the same season or period of monitoring which contain data anomalies. Any plots that contain anomalous data that cannot be resolved will be removed from the data set. 12. Waste Minimization and Pollution Prevention Care will be taken to avoid transport of vegetation and soil to other sites. This will be done by thorough cleaning and inspection of all equipment and clothing prior to


departure from a site. Invasive plant samples will be disposed of in a way to avoid accidental release into the environment. 13. References Anderson, L.E., H.A. Crum, and W.R. Buck. 1990. List of the mosses of North America north of Mexico. The Bryologist 93:448-499. Anderson, L.E. 1990. A checklist of Sphagnum in North America north of Mexico. The Bryologist 93:500-501. Brinson, M. M. 1993. "A hydrogeomorphic classification for wetlands," Technical Report WRP-DE-4, U.S. Army Engineer Waterways Experiment Station, Vicksburg, MS. NTIS No. AD A270 053. Cobb, B., E. Farnsworth, and C. Lowe. 2005. A Field Guide to the Ferns and their Related Families. Houghton Mifflin Co., Boston, MA. 281 + xviii pp. Connors, B. 2006. Protocols for Decontaminating Biomonitoring Sampling Equipment. ME Department of Environmental Protection DEPLW0641. Cowardin, L.M., V. Carter, F.C. Golet, and E.T. LaRoe. 1979. Classification of wetlands and deepwater habitats of the United States. FWS/OBS-79/31. U.S.D.I. Fish and Wildlife Service, Washington D.C. http://wetlands.fws.gov/Pubs_Reports/Class_Manual/class_titlepg.htm Danielson, T. 2004. Protocols for Collecting Water Grab Samples in Rivers, Streams, and Freshwater Wetlands. ME Department of Environmental Protection DEPLW0637. Danielson, T. 2006. Protocols for Sampling Algae in Wadeable Rivers, Streams, and Freshwater Wetlands. ME Department of Environmental Protection DEPLW0634 DiFranco, J. 2006. Protocols for Sampling Aquatic Macroinvertebrates in Freshwater Wetlands. ME Department of Environmental Protection DEPLW0640. Esslinger, T.L. 2007. A cumulative checklist for the lichen-forming fungi, lichenicolous, and allied fungi of the continental United States and Canada. North Dakota State University: http://www.ndsu.nodak.edu/instruct/esslinge/chcklst/chcklst7.htm. Gleason, H.A. 1952. The New Britton and Brown Illustrated Flora of the Northeastern United States and Adjacent Canada. Hafner Press, NY, NY. 3 volumes, 1,732 + lxxv pp. Gleason , H.E. and A. Cronquist. 1991. Manual of the Vascular Plants of Northeastern United States and Adjacent Canada. 2nd edition. New York Botanical Garden Press. Hale C.M., L.E. Frelich, and P. B. Reich. 2005. Exotic earthworm invasion dynamics in northern hardwood forests of Minnesota, USA. Ecological Applications 15(3):848-860.


Jackson, Scott. 1995. Delineating Bordering Vegetated Wetlands Under the Massachusetts Wetlands Protection Act: A Handbook. MA Department of Environmental Protection. Boston, MA. 89 + 5 pp. Lawrence, A. P., and M. A. Bowers. 2002. A test of the ‘hot’ mustard extraction method of sampling earthworms. Soil Biology and Biochemistry 34:549–552. Magee, D.W. and H.E. Ahles. 1999. Flora of the northeast: A Manual of the Vascular Flora of New England and Adjacent New York. 1213 pages. University of Massachusetts Press, Amherst, MA. McGarigal, K., B.W. Compton, S.D. Jackson, K. Rolih, and E. Ene. 2005. Conservation Assessment Prioritization System (CAPS) Highland Communities Initiative PHASE 1. Final Report. Landscape Ecology Program, Department of Natural Resources Conservation, University of Massachusetts, Amherst (URL: http://www.umass.edu/landeco/research/caps/reports/caps_reports.html Merrit, R.W., K.W. Cummins (editors). 1996. An introduction to the aquatic insects of North America, Third Edition. Dubuque, IA: Kendall/Hunt Publishing Company. 862 pp. Munsell Color.2000. Munsell Soil Color Chart Soil. Grand Rapids, MI. Newcomb, L. 1977. Newcomb’s Wildflower Guide. Little, Brown & Co., Boston, MA. 490 + xxii pp. Petrides, G.A. 1972. A Field Guide to Trees and Shrubs. Houghton Mifflin Co., Boston, MA. 428 + xxxii pp. Reynolds, J. W. 1977. The earthworms (Lumbricidae and Sparganophilidae) of Ontario. Royal Ontario Museum Miscellaneous Publication, Toronto, Ontario, Canada. Scanlon. J. 2006. MassWildlife Landcover Mapping Decision Rules. Internal Document. Massachusetts Division of Fisheries and Wildlife, Westborough, MA. Schauff, M.E. 1986. Collecting and Preserving Insects and Mites: Techniques and Tools. Systematic Entomology Laboratory, USDA, Washington, D.C. Somers, P. K. Lombard, and R. Kramer. 2006. A Guide to Invasive Plants in Massachusetts. Massachusetts Division of Fisheries & Wildlife, Natural Heritage & Endangered Species Program, Rabbit Hill Road, Westborough, MA 01581. Schuster, R.M. 1974. The Hepaticae and Antherocerotae of North America: East of the Hundredth Meridian Volume 3. Columbia University Press, New York, NY, USA and London, UK. Schwert, D. P. 1990. Oligochaeta: Lumbricidae. Pages 341–356 in D. L. Dindal, editor. Soil biology guide. John Wiley and Sons, New York, New York, USA.


Stevenson, R.J. 2001. Using Algae to Assess Wetlands in Multivariate Statistics, Multimetric Indices, and Ecological Risk Assessment Framework. pages 113-140 in Bader, R.B., Batzer, D.P. and Wissinger, S.A. (eds.), Bioassessment and Management of North American Freshwater Wetlands. John Wiley & Sons, NY, NY. Swain, P.C. and J.B. Kearsley. 2001. Classification of the Natural Communities of Massachusetts DRAFT - 2001- (version 1.3) Reprinted 2004. Massachusetts Division of Fisheries and Wildlife, Westborough, MA. U.S. Department of Agriculture, Natural Resources Conservation Service. 2002. Field Indicators of Hydric Soils in the United States, Northeastern & Northcentral Regions, Version 5.0. G.W. Hurt, P.M Whited and R.F Pringle (eds). USDA-NRCS in cooperation with the National Technical Committee for Hydric Soils, Fort Worth, TX. U.S Department of Agriculture, Soil Survey Staff. 2006. Soil Taxonomy, Tenth Edition. Handbook No. 463. U.S Government Printing Office, Wasington, D.C U.S. EPA, 2002. Methods for Evaluating Wetland Condition: Developing an Invertebrate Index of Biological Integrity for Wetlands. Office of Water, U.S. Environmental Protection Agency, Washington , DC. EPA-822-R-02-019 U.S. EPA. 2002 Methods for Evaluating Wetland Condition: Using Algae to Assess Environmental Conditions in Wetlands. Office of Water, U.S. Environmental Protection Agency, Washington , DC. EPA-822-R-02-021

_________________________________________________________________________________________________________ 2009 CAPS – Forested Wetlands Assessment

FORM Plot Information A Point ID: GPS ID:

Digital Photo ID: Surveyor ID: Dates visited

Plot setup ET set: Algae Stovepipe PT set: Plants ET collect: PT collect:

ibutton serial number: HOBO serial number: Landowner Information:

Plot Diagram Include the following info: Number of transects, transect lengths, transect azimuth, location of emergence traps (ET), pitfall traps (PT), stovepipe, algae samples, and location of data loggers. Denote vascular plant transects, and bryophyte plots. Draw in location and flow of any hydrologic features (streams) present in plot or nearby. Note local disturbances or structures in plot. Add comments below.

N W E S Comments:

_________________________________________________________________________________________________________ 2009 MA Assessment of Forested Wetlands Communities

FORM Plot Information B Point ID: Surveyor: Date: Water geochemistry (Measure where algae is sampled. Dig a small pit if there is no surface water. If a stream is present in plot, take a measurement.)

Surface Water (SW), Ground water (GW), or Flowing Stream

pH Conductivity Temp.

1

2

3

4

5 (stream)

Water depth (cm) *note wet depression for stovepipe samples if no standing water is present Locations Date 1 2 3 4

Stovepipe sample (2 locations in plot)*

Algae: Leaf litter sample

Algae: Water sample

Algae: Surface sediment

Comments:

2009 CAPS – Forested Wetlands Assessment

FORM HYDROLOGICALCHARACTERIZATION POINT ID: GPS ID:

Visit # 1 (Plot

establishment)

2 (ET set)

3 (ET collect)

4 (PT set)

5 (PT collect)

6 (Plant survey)

Date:

Surveyor Initials

HOBO H20 depth

(cm)

Transect

On 4-30m transects tally whether W, M, or D at every 5m flag for a max of 6 per transect. W=Water >2.5 cm deep; M=Muck (Saturated to Surface, water < 2.5cm); D=Dry (Not saturated to surface)

N

W M D

W M D

W M D

W M D

W M D

W M D

E

W M D

W M D

W M D

W M D

W M D

W M D

S

W M D

W M D

W M D

W M D

W M D

W M D

W

W M D

W M D

W M D

W M D

W M D

W M D

Totals By Date:

(max=24)

W M D

W M D

W M D

W M D

W M D

W M D

Tally stream crossings or dug channels with an asterisk * Comments:

Emergence Trap Log CAPS 2009 Point ID: Date Set: Date Collected: Transect

Setter ID H2O

Depth (m) (set)

Habitat Condition Ethanol level

H2O Depth (m) (collect)

Collector ID

N

E

S

W

Comments: Habitat descriptors: W=surface water, M=saturated soil, D=Dry Condition: F=Fallen over, D=Damaged, G=Good Ethanol level: P=Present, N=None left (record at time sample is collected)

Pitfall Trap Log CAPS 2009 Point ID: Date Set: Date Collected: Transect

Location Setter ID Habitat Water

content Condition Collector

ID N 10 m

15 m E 10 m

15 m S 10 m

15 m W 10 m

15 m Comments: Microhabitat descriptors: D=Depression, M=Mound, F=Flat. Describe the depression, mound or flat by using M=Moss, L=Leaf cover, of V=Vegetation. (e.g.: DM=depression with moss) Water content: F=flooded (greater than half full of water), PF=partial flooding (less than half full of water), N= no flooding. Condition: G=no damage, BR=bowl removed, D=Destroyed

FORM TOPOGRAPHIC COMPLEXITY* CAPS 2009 POINT ID: Date:

Surveyor:

Transect # depressions > 1m2 Totals

N

E

S

W

Comments:

* One-time count of microtopographic depressions ≥ 1m2, elevation change ≥ 15cm (6”) high, along each transect during a dry period.

FORM Vascular Plants 2009 PointID: Pg of Surveyors: Date: NHESP Natural Community:

________________________________________________________________________________________________________________________ Draft 30 April 2009 MassCAPS & MA DEP Rapid Assessment – Forested Wetlands

Tran-sect (0-360º)

List and tally all plants, unvegetated surface (bare soil, litter, rock, and wood under 4”) every 1 meter, from ground to canopy

1-5 6-10 11-15 16-20 21-25

FORM BRYOPHYTES CAPS 2009

POINT ID GPS ID:

Surveyor: Date: Cover class +=%, 1=1-5%, 2=6-25%, 3=26-50%, 4=50-75%, 5=<75% Subplot Species # % cover Substrate Comments

Area Search Start Time: Specimens collected in area search ? Yes or No

FORM Epiphytic Macrolichens 2009 PointID: Pg of Surveyors: Date: factor prism (D)=Dead (V)=Voucher Start Time:

________________________________________________________________________________________________________________________ Draft 30 April 2009 MassCAPS & MA DEP Rapid Assessment – Forested Wetlands

Cover class +=%, 1=1-5%, 2=6-25%, 3=26-50%, 4=50-75%, 5=<75% Tree# Tree species Dbh(in.) Lichen species % cover Species found on litterfall etc:

Documents

5.2 MassDEP/MassCZM Standard Operating Procedures and Data ...neiwpcc.org/nebawwgold/documents/ManualSections/Section 5_2.pdf · Adrienne Pappal Commonwealth of Massachusetts Executive