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4.3 SOLID MATRIX SAMPLING PROCEDURES
This section is concerned with grab and
(areal or depth) composite samples from solid matrix (soil and sediment).
Since similar procedures and equipment exist for soils and sediments, a
general description of sample handling will be discussed.
EPA
and
the Department define sediments as the solid matrix material that is located
beneath surface waters. Although
terrestrial soil is "sedimentary" material, these matrices are
considered distinct.
4.3.1.1 Sampling
equipment shall be selected based on the type of sample to be collected and the
parameters of interest. See Table
4.1 for specific requirements and Section 4.2.5.2 of the Groundwater section for
a discussion on material.
4.3.1.2 All
equipment shall be decontaminated according to specified protocols in Section
4.1.
4.3.1.3
All general sampling concerns outlined in Section 4.0 shall be followed.
4.3.1.4 Field
activities shall be documented in accordance with requirements in Section 5.
4.3.1.5 Sample
container and holding time requirements listed in Section 4.4.2.3 shall be
followed. The sample containers
shall be cleaned or obtained according to protocols listed in Section 4.4.1.1.
4.3.2
Sample Handling Protocols after Sample Acquisition
General sample handling will fall into
3 main categories; surface, shallow subsurface, and deep subsurface. Each of the three categories will be discussed in general.
Once the sample is acquired, the handling procedures are very similar and
are described below.
1. Select the
appropriate precleaned sampling device and procure the sample from the desired
depth. If using liners to transport the sample to the lab, see
4.3.2.10 below.
2. Select the required
sample container for the parameter group.
3. Split spoons and
Shelby Tubes
a.
Breakdown the sampler (split spoon, Shelby tube).
This should be done with the appropriate tools.
b.
At this time, any portion of the sample that has been disturbed shall be
identified, removed with a stainless steel spatula and discarded.
c.
Slice the sample using a clean, decontaminated stainless steel spatula
from the center portion of the corer, split spoon or bucket auger head.
d.
For VOC analyses, immediately transfer the sliced portion to a suitable
container (the container must be equipped with a teflon-lined septum seal).
1.
Carefully fill the vial (or wide-mouth container) with sample.
2.
Tamp the sample into the vial with a SS, glass, or Teflon rod to reduce
headspace.
3.
Add sample and tamp down until no headspace exists.
4.
After cleaning exterior and rim of vial as described below, cap container
(with teflon side facing sample) tightly, label and place on wet ice
immediately.
e.
For other analyses, slice sufficient amount of sample from the center
portion of the sampling device and transfer it to a tray of appropriate
construction (note restrictions on use in Table 4.1).
4.
Bucket Auger, Dredge or Corer
a.
Remove the sample from the sampler (bucket auger, dredge, corer) with
appropriate (stainless steel, teflon, etc.) tools and place in a stainless
steel, glass or aluminum foil-lined tray (note restrictions on use in Table 4.1.
b.
Remove any portion of the sample that has been disturbed with a stainless
steel spatula and discarded.
c.
If VOCs are required, fill an appropriate container with aliquots that
have been taken from selected areas of the entire sample.
Proceed as described in 4.3.2.3.d above.
5.
Sample Mixing
a.
VOCs shall be collected as discussed above before the sample is mixed.
b.
The sample in the tray shall be homogenized thoroughly:
1.
Appropriate tools shall be used to mix the sample.
2.
Homogenize by alternately mixing, dividing, and remixing the sample.
c.
After thorough mixing, transfer the sample to the appropriate sample
container(s) leaving minimal headspace.
6. Clean the outside of
the sample container to remove excess
soil.
7. The container rim
should also be cleaned of soil and sand particles so that the lid can be sealed.
An improperly sealed container may allow cross contamination from ice
melt or petroleum fumes.
8. Affix sample label,
seal (if applicable), and complete the chain-of-custody forms.
9. Place the sample
containers in a clean, plastic sample bag and preserve by placing in wet ice.
10. Liners
a.
If properly used, liners may be inserted into the sampler and used as the
actual sample container.
b.
Be aware that SW-846 has mandated that all solid samples must be
transported in containers that have screw tops.
This also means that all container and lid requirements are still in
effect.
1.
For inorganic samples, ends of the liner must be covered with
polyethylene, Teflon, or aluminum foil sheeting.
The sheeting should be secured by placing an end cap over the sheeting.
2.
For organic samples, the sheeting must be Teflon or aluminum foil.
3.
With any sample containerized this way, specific instructions must be
sent with the sample so that the laboratory will know how to handle the sample.
All non-volatile samples must be homogenized by the laboratory prior to
analyses. Also, any disturbed
portions of the sample should be discarded prior to mixing.
The following is not a complete
discussion regarding development of a sample compositing scheme nor all
available sampling protocols. When
a large site area is to be investigated for contamination, it is sometimes
advantageous to composite soil or sediment samples and thus minimize the number
of samples to be analyzed.
1. Sample aliquots (of
identical size) to be composited shall be placed in a tray of suitable materials
(see Table 4.1) and thoroughly mixed with a cleaned spoon, spoonula or spatula
of suitable materials (see Table 4.1). The
sample shall be thoroughly blended by mixing, and dividing into sections.
Each section shall then be mixed separately.
Recombine all mixed sections and mix thoroughly.
Repeat sectioning and mixing process to ensure proper homogenization.
2. The origin and size
of each (sub)sample or aliquot that is used to make the composite shall be
documented in the field notebook along with the other important sampling
details.
a.
Although the size of these subsamples is important and should be
documented, it is critical that these subsamples be of equivalent size, so that
the composite sample is not biased by unequal aliquoting.
b.
There is no level of accuracy here; it is dependent upon the size of the
aliquots.
c.
Aliquoting should be done in a systematic manner.
3. Samples
for VOCs shall not be composited or mixed unless the DER programs or permits
require compositing. This procedure
should only be undertaken if mandated by a formal DER document (permit, Consent
Order, etc.) and shall follow these protocols:
a.
Composite samples for VOCs shall not be mixed as described above.
b.
Equal portions of all samples to be included in the composite shall be
placed in the VOC container (see 4.3.2.3.d above).
c.
The laboratory shall be informed that the sample is a composite and must
be mixed before analysis.
4. Clean
the outside of the sample container to remove excess soil, affix label, seal (if
required), and complete the laboratory transmittal forms.
[[4.3.4.1
Site Selection
a.
Soil sampling locations should be selected such that a representative
portion of the soils are collected with minimal disturbance.
Locations where natural vegetation is stressed or dead and/or areas that
have surficial soil staining may be indicative of improper waste disposal
practices.
b.
An upgradient, undisturbed location should also be selected for obtaining
background and/or quality control samples.
Be aware that differences in soil types may affect these background
samples (e.g. sands vs. clays).]]
4.3.4.2
Surface Soil Sampling - ground surface to 6 inches below ground surface
a.
Leaves, grass and surface debris shall be removed from the area to be
sampled using a clean stainless steel spoon or shovel.
b.
Surface soil samples can then be collected using a precleaned stainless
steel scoop or spoon.
4.3.4.3
Shallow Subsurface Soil Sampling
a.
Shallow subsurface samples may be collected by digging a hole or trench
to the required depth with a stainless steel shovel.
b.
Some situations may require a trench or pit to be dug with a backhoe.
Depending upon the equipment available at the site or the soil type to be
penetrated, this option is acceptable. [[Please
note that any OSHA requirements for in-trench sampling should be followed.]]
1.
In these situations, the trench is first dug to the appropriate depth and
then the sample is exposed by using one precleaned spoon, spatula, or equivalent
to clean away the soil that came in contact with the backhoe bucket and a second
precleaned spoon to actually collect the sample.
c.
Alternatively, shallow subsurface soil samples may be
collected with 2-4 inch stainless steel bucket auger which would minimize
the soil to be removed in order to reach the desired depth.
Using this method, a sampling depth of up to 15 feet may be obtained.
1.
The bucket auger consists of a stainless steel cylinder with flush welded
stainless steel cutting edges. The
cutting edges are hardened surfaces, heat treated and sharpened.
2.
A soil sample is obtained by pushing and rotating the auger into the soil
until the bucket is filled.
3.
The sample can be removed (with some difficulty) from the bucket by
pushing or scraping with an appropriate precleaned stainless steel tool.
4.
This auger method is useful for obtaining large samples of unconsolidated
sediment.
5.
The device is supplied with 3 foot extension rods.
6.
Addition of a sleeve may allow an undisturbed soil sample to be obtained.
a.
The device consists of a standard auger head with a removable
non-contaminating sleeve which is inserted into the auger barrel.
b.
Either a clear butyl acrylate (CAB) plastic sleeve (for inorganic
samples) or stainless steel (for organic samples) may be utilized.
c.
The soil sample is obtained in the normal manner by pushing and rotating
the auger into the soil. In this
case it is the sleeve which fills with soil.
After auger retrieval, the sleeve, which is readily removed from the
auger, is capped (see Section 4.3.2.10 above).
7.
If the auger hole is prone to collapse, due to low cohesion in some
soils, a temporary rigid PVC casing should be inserted into the hole.
The casing prevents hole collapse and minimizes cross-contamination
between soil zones as the auger is advanced.
d.
Upon sample collection, the temporary casing (if used) must be removed
and the hole filled with the excavated soil.
[[e.
If a confining layer has been breached during sampling, the hole shall
should be grouted to land surface with Type-1 Portland cement.
NOTE: this requirement may
be different throughout Florida. Contact
the local Water Management District office for local requirements.]]
4.3.4.5
Deeper Subsurface Soil Sampling
a.
A drill rig is normally required if soil samples are taken from boreholes
greater than 15 feet BLS (below land surface).
There are a number of sampling devices used in conjunction with the drill
rig for retrieving the samples; Shelby tubes, split spoon samplers and standard
core barrels.
b.
Shelby Tube Sampler
1.
The Shelby tube sampler is used to sample unconsolidated soils and
consists of a stainless steel tube approximately 30 inches long and 2 inches, or
larger, in diameter.
2.
One end of the tube has edges beveled into a cutting edge. The other end
can be mounted to an adapter which allows attachment to the drill rig assembly.
3.
After drilling to the required depth with an auger or rotary drill bit, a
soil sample is obtained through the auger or directly in the borehole.
4.
The Shelby tube is pushed into the soil using the drill rig's hydraulic
ram or manually with a sledge hammer.
5.
When the tube is retrieved, the soil sample taken from the center and
away from the sides, can be transferred into the appropriate container (for
VOCs) and/or mixed using a stainless steel spoon handle or spatula when other
parameters are of interest.
c.
Split Spoon Sampler
1.
A split spoon sampler, useful for sampling unconsolidated soils, consists
of two carbon steel half cylinders (spoons) that fit together to form a tube
approximately 2 feet in length and 2 inches in diameter.
2.
The cylindrical arrangement is maintained by a retaining head and bit
rings that screw on at either end of the split spoon.
3.
The bit ring has beveled edges to facilitate sampling as the split spoon
is forced into the ground.
4.
As with the Shelby tube, either the weight of the drill stem and rods or
a mechanical hammer is used to advance the sampler.
5.
A catcher device is inserted in the head ring to prevent loss of
unconsolidated sample during recovery.
6.
After retrieving the split spoons, the soils can be withdrawn by
unscrewing the bit and head rings and splitting the barrel.
7.
The top 2 to 3 inches of the sample will be normally disturbed and should
be discarded.
8.
A cleaned stainless steel spatula is used to collect a subsample for VOCs
and/or transfer the contents into an appropriate tray for mixing and
containerizing.
d.
Standard Core Barrel
1.
A standard core barrel is utilized when consolidated samples (such as
limestone or dolomite) are to be sampled.
2.
The core barrel is carbon steel cylinder approximately 3 feet long and 2
inches in diameter.
3.
The barrel has a removable head ring with small embedded diamonds which
allow the device to cut through rock or consolidated soils as the drill rods are
rotated.
4.
The sample core can be retrieved by unscrewing the head ring and sliding
the sample into the container.
4.3.5.1
General Overview
a.
Sediment samples are taken from material underlying streams/rivers,
lagoons, ponds/lakes, and estuaries.
b.
The actual sampling location
is dependent upon project scope.
c.
Sediment samples may be taken as an adjunct to surface water samples.
d.
They may be taken as a compositing series to define water or sediment
quality in a system.
e.
They may be taken above and below an outfall to document degradation.
f.
Similarly, if stressed shore vegetation or visible surface water
contamination is evident, sediment samples may be taken.
g.
Decisions for sample location will not be discussed in this document.
h.
All surface water samples shall be taken prior to any sediment samples
(see 4.2.3.2).
4.3.5.2
Sample Collection Protocols
a.
Sediment samples are taken via three groups of equipment:
scoops; corers and dredges.
b.
Soil sampling equipment is generally not applicable to sediments because
of low cohesion of sample.
c.
Sample location (edge or middle of lagoon), depth of water and sediment,
sediment grain size (fineness), water velocity, and analytes of interest all
must be considered when choosing equipment.
d.
Stainless steel equipment shall be used if trace contaminants are to be
sampled.
e.
Dredges must be used for hard or rocky substrates.
They are heavy enough to use in high velocity streams.
f.
Coring device may be used for softer substrates.
Coring devices must be used in soft substrate if the fine particles are
to be included. Coring devices
should be used in quiescent waters.
g.
Scoops
1.
Scooping is generally most useful around the margin or shore of the water
body.
2.
Stainless steel spoons or grain scoops work very well.
The scoops can be attached to an extendible pole for obtaining samples
several feet from shore or boat.
3.
The sampler may also wade into the water body to obtain a scooped sample.
4.
The sampler must stand facing the direction of flow and approach the
location from the downstream direction.
5.
Precautions must be taken not to disturb the bottom prior to scooping.
6.
The sample shall be scooped in the upstream direction of flow.
h.
Corers
1.
Coring devices can be easily fabricated from many materials.
Although stainless steel, glass or teflon must be used for sampling trace
organics, other inexpensive material (PVC, carbon steel, etc.) may be used for
demands, nutrients, metals as appropriate.
2.
Some corers are simple "push tubes", whereas other more
sophisticated models may be finned, gravity driven devices.
3.
Not only are they useful in sampling fine grain sediments, they can also
present or preserve the historical layering of sediments.
4.
Upon descent, water displacement is minimal, which also minimizes the
shock wave produced by other equipment (dredges).
5.
The corer is the equipment of choice for fine sediments in static waters,
especially trace organics and metals.
6.
Corer diameter, grain size, and sample consistency will determine if the
sample will remain in the corer upon withdrawal.
7.
Sample washout can be a problem and there are several ways to reduce or
prevent it.
a.
The leading edge of the corer can be fitted with a nosepiece or core
catcher which physically keeps the sample from slipping back out the corer.
1.
The core catcher material must also be compatible with analytes of
interest.
b.
A second option is fitting the top or back end with a check valve which
first creates negative pressure on the back of the sample as it is being pulled
from the substrate and second, prevents surface water from washing out the top
portion of the sample.
8.
The corer shall be rotated as it is pushed in.
a.
Rotation should be around its axis, not rocked back and forth.
b.
Rotation improves penetration and prevents compaction of the sample as it
is pushed to the full length of the corer.
9.
Upon withdrawal from the water surface, a cap shall be placed on the
bottom to prevent the sample from sliding out.
10. The
core should then be extruded out into a pan or tray and sample processed as
described in Section 4.3.2 above.
11.
Corers can also be fitted with liners.
This is advantageous if a complete core is desired that has not been in
contact with the atmosphere. It is
also advantageous if the coring device is not constructed of the proper material
(e.g. PVC) and one of the analytes requires a sampler of inert construction
(glass, SS, or Teflon).
i.
Dredges
1.
The three main types are the Peterson, Eckman, and Ponar.
2.
The Peterson and Ponar dredges are suitable for hard or rocky substrates.
a.
The Peterson and Ponar are virtually the same, except the Ponar has been
adapted with a top screen and side plates to prevent sample loss upon ascent.
For this reason, the Ponar is the dredge of choice for rocky substrates.
These dredges are heavy enough to use in streams with fast currents.
3.
The Eckman is designed for softer substrates of sand, silt, or mud.
a.
The Eckman is too light to use in fast currents.
4.
Follow the manufacturer's suggestions for setting
and operating the weighted messenger devices.
4.3.6.1
Decontamination
a.
It is important to properly decontaminate sampling equipment prior to
going into the field and between individual fish, as heavy metals and organic
compounds are usually the analytes of interest in fish tissues and fish slime
contains these analytes. Therefore,
all equipment used for tissue sampling shall be decontaminated using the
following procedure:
1.
Rinse equipment with tap water.
2.
Wash with Liquinox or Acationox (or a comparable detergent).
A brush may be required to remove scales and other tissue remnants.
3.
Rinse with deionized water.
4.
Rinse with isopropanol.
5.
Use analyte-free water for a final rinse.
6.
Decontaminated equipment should be wrapped in aluminum foil or untreated
butcher paper to protect it from contamination if it will be stored or
transported prior to use.
NOTE:
If analyte-free water is unavailable, allow isopropanol rinsed equipment
to dry thoroughly before use.
4.3.6.2
Sample Collection/Sample Preparation
a.
Fish shall be captured using the means appropriate for the situation
(electroshock, seine net, hook and line, etc.) and immediately placed on wet ice
in coolers.
b.
Balances, cleaned by the method above and rezeroed after weighing each
fish, should be used to weigh the fish as soon as possible after capture.
c.
NOTE: if the fish are to be used for the analysis of dioxins by
Method 8090, the following procedures for filleting SHALL NOT be followed.
WHOLE fish shall be frozen and transported to the laboratory for
processing.
d.
The filleting procedure should take place on a stable surface.
The surface shall be constructed of stainless steel or other suitable
materials (see table 4.1) which can be decontaminated initially and after each
fish is prepared.
e.
Using a clean stainless steel fillet knife with either a wooden or
stainless steel handle, the individual fish should be filleted carefully, so as
not to puncture any visceral organs.
f.
The sampler should wear disposable gloves (see Section 4.0.2) which must
be changed between fish.
g.
Fillets shall be scaled, but not skinned, as both skin and muscle tissue
are considered edible portions of the animal.
(This is especially important if organic analytes are being sampled, as
these compounds are fat soluble and a layer of fat lies directly below the
skin.)
h.
Duplicate samples shall be collected from the same regions of opposite
fillets, labeled, stored and analyzed as duplicates.
i.
Samples may be of any size and may include the whole fillet, but
must be taken through the entire width of the fillet if smaller.
j.
All samples taken shall be wrapped in clean aluminum foil, labeled and
placed in a plastic bag.
k.
At a minimum, samples shall be preserved by keeping them on wet ice while
in the field for no more than 24 hours and shall be frozen upon return to the
laboratory. It is important to guarantee that the moisture content does not drop
through repeated freeze-thaw cycles.
l.
Optimal preservation of fish tissue samples should include lyophilization
(freeze-drying) combined with determination and recording of the sample's
moisture content (weight loss) after freeze-drying.
The latter technique is often beyond the scope of many laboratories and
most consultants.
4.3.6.3
References:
1.
Mid-America Fish Contaminants Group, August 1989, Fish Sampling
Guidelines.
2.
DER Water Quality Monitoring and Quality Assurance Section, Bureau of
Water Analysis, Tallahassee, FL, July 1984 Draft, Interim Method For The
Sampling and Analysis Of Metals In Fish Tissue With Emphasis On Mercury.
3.
U.S. E.P.A., Environmental Monitoring and Support Laboratory, Cincinnati,
OH. October 1980, Interim Methods For The Sampling And Analysis Of Priority
Pollutants In Sediments and Fish Tissues.
This protocols pertain only to the
collection of whole organisms which will be processed by the laboratory. This means that the shellfish must be collected with shells
INTACT.
4.3.7.1
Sample Collection
a.
Shellfish may be brought to the surface using divers, clam rakes, tongs
or any other suitable means.
b.
Unless specific species are required, the sample should include samples
of all species.
c.
Select only live organisms. This
means that the shells are tightly closed and cannot be manually pried open.
d.
The laboratory must be consulted for the number of shell fish to submit.
Since many shells will open in transit, collect at least twice as many
specimens as the laboratory required.
4.3.7.2
Sample Transport and Preservation
a.
Wrap all specimens from one location securely in aluminum foil.
b.
Place in a sealable plastic bag, seal bag, identify contents with a field
ID number and place immediately on ice.
c.
Complete documentation which must include:
1.
Description of sampling location
2.
Depth that specimens were found
3.
Species identification (if possible)
4.
Number of different types of specimens (if applicable)
5.
Collection time
6.
Collection equipment and method
d.
Samples must be transported to the laboratory within 24 hours of
collection. They may be frozen for
transport, but must be frozen immediately upon receipt in the laboratory.
NOTE:
If samples are frozen for transport, sufficient ice and/or dry ice must
be packed to that they remain frozen during transit.
Residuals matrix is defined as domestic
waste sludge residuals. All
sampling must be conducted by following EPA's POTW Sludge Sampling and Analysis
Guidance Document, 1989.
4.3.9
Hazardous Waste Sampling
Hazardous Waste Sampling shall follow
the protocols outlined in the following sections of the EPA Region IV
"Standard Operating Procedures and Quality assurance Manual" (February
1991):
a.
Waste: Pits, Ponds and
Lagoons - Section 4.12.3, pp. 3 through 5.
b.
Waste: Open and Closed
Container Sampling - Section 4.12.4, pp. 5 through 8.
c.
Waste: Waste Piles and
Landfills - Section 4.12.5, pp. 8 through 9.
4.3.10
Sampling Protocols for Macrobenthic Invertebrate Identification
Sample collection macrobenthic
organisms shall follow protocols specified in these documents:
a.
"Macroinvertebrate Field and Laboratory Methods for Evaluating the
Biological Integrity of Surface Waters", ORD, Washington, D.C., November
1990;
b.
Standard Methods for the Examination of Water and Wastewater, Part 10500,
17th Edition, APHA, 1989.