The Essential Handbook of Ground Water Sampling - Chapter 5 pptx

5
Ground-Water Sample Pretreatment: Filtration and
Preservation
Gillian L. Nielsen and David M. Nielsen
CONTENTS
Sample Pretreatment Options . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131
Sample Filtration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131
What Fiter Pore Size to Use . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132
Functions of Filtration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134
Which Parameters to Filter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134
Sources of Error and Bias in Filtration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136
Filtration Methods and Equipment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137
Filter Preconditioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139
Sample Preservation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 141
Objectives of Sample Preservation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 141
Physical Preservation Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142
Chemical Preservation Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149
Sample Pretreatment Options
Another group of parameter-specific field protocols that must be evaluated and included
in the SAP are methods for sample pretreatment, including sample filtration and physical
and chemical preservation. Sample pretreatment must be performed at the wellhead at
the time of sample collection to ensure that physical and chemical changes do not occur in
the samples during the time that the sample is collected and after the sample container
has been filled and capped. ASTM International has published Standard Guides that
address both types of sample pretreatment. ASTM Standard D 6564 (ASTM, 2006a)
provides a detailed guide for field filtration of ground-water samples, and ASTM
Standard D 6517 (ASTM, 2006b) discusses physical and chemical preservation methods
for ground-water samples. Each type of sample pretreatment is discussed subsequently.
Sample Filtration
Ground-water sample filtration is a sample pretreatment process implemented in the

field for some constituents, when it is necessary to determine whether a constituent is
truly ‘‘dissolved’’ in ground water. Filtration involves passing a raw or bulk ground-
water sample directly through a filter medium of a prescribed filter pore size either under
131
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negative pressure (vacuum) or under positive pressure. Particulates finer than the filter
pore size pass through the filter along with the water to form the filtrate, which is
submitted to the laboratory for analysis. Particulate matter larger than the filter pore size
is retained by the filter medium. In the case of most ground-water monitoring programs,
this material is rarely analyzed, although it may be possible to analyze the retained
fraction for trace metals or for some strongly hydrophobic analytes such as PCBs or
PAHs. Figure 5.1 illustrates a common vacuum filtration setup, and Figure 5.2 illustrates
one form of positive pressure filtration.
What Filter Pore Size to Use
The most common method for distinguishing between the dissolved and particulate
fractions of a sample has historically been filtration with a 0.45 mm filter (see, e.g., U.S.
EPA, 1991). The water that passes through a filter of this pore size has, by default, become
the operational definition of the dissolved fraction, even though this pore size does not
accurately separate dissolved from colloidal matter (Kennedy et al., 1974; Wagemann and
Brunskill, 1975; Gibb et al., 1981; Laxen and Chandler, 1982). Some colloidal matter is
small enough to pass through this pore size, but this matter cannot be considered
dissolved. For this reason, Puls and Barcelona (1989) reported that the use of a 0.45 mm
filter was not useful, appropriate, or reproducible in providing information on metals
solubility in ground-water systems and that this filter size was not appropriate for
determining truly dissolved constituents in ground water.
The boundary between the dissolved phase and the colloidal state is transitional. There
is no expressed lower bound for particulate matter and no clear cutoff point to allow
selection of the optimum filter pore size to meet the objective of excluding colloidal
FIGURE 5.1
A vacuum filtration system used for ground-water samples. This practice is not encouraged.

132 The Essential Handbook of Ground-Water Sampling
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particles from the sample. The best available evidence indicates that the dissolved phase
includes matter that is less than 0.01 mm in diameter (Smith and Hem, 1972; Hem, 1985),
suggesting that a filter pore size of 0.01 mm is most appropriate. However, filters with
such small pore sizes are subject to rapid plugging, especially if used in highly turbid
water, and are not practical to use in the field. Kennedy et al. (1974) and Puls et al. (1991)
provide a strong case for the use of a filter pore size of 0.1 mm for field filtration to
allow better estimates of dissolved metal concentrations in samples. Puls et al. (1992) and
Puls and Barcelona (1996) also recommend the use of 0.1 mm (or smaller) filters for
determination of dissolved inorganic constituents in ground water. Such filters are
considerably more effective than filters with larger pore sizes (e.g., 0.45 or 1.0 mm) in
terms of removing fine particulate matter. These filters are widely available and practical
for use in the field for most situations, although in some highly turbid water, filter
plugging may make the filtration process difficult and protracted. All factors considered,
0.1 mm field filtration, although it is a compromise, appears to offer the best opportunity
for collecting samples that best represent the dissolved fraction.
Yao et al. (1971) indicate that colloids larger than several microns in diameter are
probably not mobile in aquifers under natural ground-water flow conditions due to
gravitational settling. Puls et al. (1991) also suggest that colloidal materials up to 2 mmare
mobile in ground water systems. With the upper bound for colloidal matter described by
many investigators as being between 1.0 and 10 mm, it seems reasonable to suggest that
a filter pore size of 10 mm would include all potentially mobile colloidal material and
exclude the larger, clearly nonmobile artifactual fraction. However, it should be noted that
using this filter pore size, artifactual colloidal material that is finer than 10 mm in diameter
will be included in the sample. Although this filter pore size is a compromise, it will lead
to conservative estimates of total mobile contaminant load while excluding at least a
portion of the particulate matter that is artifactual in nature. The collection and analysis of
both filtered and unfiltered samples is sometimes suggested as a means of discriminating
between natural and artifactual colloidal material or between dissolved and colloidal

contaminant concentrations.
FIGURE 5.2
A positive-pressure filtration system is a better option to use for ground-water samples. Note the removal of
sediment achieved by the cartridge filter.
Ground-Water Sample Pretreatment: Filtration and Preservation 133
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Functions of Filtration
Historically, filtration of ground-water samples has served several important functions in
ground-water sampling programs. Filtration helps minimize the problem of data bounce,
which commonly results from variable levels of suspended particulate matter in samples
between sampling events and individual samples, making trend analysis and statistical
evaluation of data more reliable. In addition, by reducing suspended particle levels,
filtration makes it easier for laboratories to accurately quantify metals concentrations in
samples. Perhaps most importantly, filtration of samples makes it possible to determine
actual concentrations of dissolved metals in ground water that have not been artificially
elevated as a result of sample preservation (acidification), which can leach metals from
the surfaces of artifactual or colloidal particles (Nielsen, 1996). The assumption that the
separation of suspended particulates from water samples to be analyzed eliminates only
matrix-associated (artifactual) constituents may often be incorrect (EPRI, 1985a; Feld et al.,
1987), as at least some potentially mobile natural colloidal material will be retained on
most commonly used filter pore sizes.
Filtration is often performed as a post-sampling ‘‘fix’’ to exclude from samples any
particulate matter that may be an artifact of poor well design or construction,
inappropriate sampling methods (use of bailers, inertial-lift pumps, or high-speed,
high-flow-rate pumps), or poor sampling techniques (agitating the water column in the
well). Filtration may be considered particularly important where turbid conditions
caused by high particulate loading might lead to significant positive bias through
inclusion of large quantities of matrix metals in the samples (Pohlmann et al., 1994).
Alternatively, as discussed earlier, the presence of artifactual particulate matter in
samples may also negatively bias analytical results through removal of metal ions from

solution during sample shipment and storage as a result of interactions with particle
surfaces. However, filtration is not always a valid means of alleviating problems
associated with artifactual turbidity, as it often cannot be accomplished without affecting
the integrity of the sample in one way or another.
Which Parameters to Filter
During the planning phase of a ground-water sampling program, each parameter to be
analyzed in ground-water samples should be evaluated to determine its suitability for
field filtration and the most suitable filtration medium. As a general rule of thumb,
parameters that are sensitive to the following effects of filtration should not be filtered in
the field:
. Pressure changes that would result in degassing or loss of volatile constituents
. Temperature changes
. Aeration and agitation that may occur during filtration processes
Table 5.1 presents a summary of parameters for which filtration may be used and of
parameters for which filtration should not be used in the field.
Samples to be analyzed for alkalinity must be field filtered if significant particulate
calcium carbonate is suspected in samples, as this material is likely to impact alkalinity
titration results (Puls and Barcelona, 1996). Care should be taken in this instance,
however, as filtration may alter the CO
2
content of the sample and, therefore, affect the
results.
Filtration is not always appropriate for ground-water sampling programs. If the
intent of filtration is to determine truly dissolved constituent concentrations (e.g., for
134 The Essential Handbook of Ground-Water Sampling
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geochemical modeling purposes), the inclusion of colloidal matter less than 0.45 mmin
the filtrate will result in overestimated values (Wagemann and Brunskill, 1975; Bergseth,
1983; Kim et al., 1984). This result is often obtained with Fe and Al, where ‘‘dissolved’’
values are obtained which are thermodynamically impossible at the sample pH (Puls

et al., 1991). Conversely, if the purpose of sampling is to estimate total mobile
contaminant load, including both dissolved and naturally occurring colloid-associated
constituents, significant underestimates may result from filtered samples, due to the
removal of colloidal matter that is larger than 0.45 mm (Puls et al., 1991). A number of
researchers have demonstrated that some metal analytes are associated with colloids that
are greater than 0.45 mm in size (Gschwend and Reynolds, 1987; Enfield and Bengtsson,
1988; Ryan and Gschwend, 1990) and that these constituents would be removed by
0.45 mm filtration. Kim et al. (1984) found the majority of the concentrations of rare earth
elements to be associated with colloidal species that passed through a 0.45 mm filter.
Wagemann and Brunskill (1975) found more than twofold differences in total Fe and Al
values between 0.05 and 0.45 mm filters of the same type. Some Al compounds, observed
by Hem and Roberson (1967) to pass through a 0.45 mm filter, were retained on a 0.10 mm
filter. Kennedy et al. (1974) found errors of an order of magnitude or more in the
determination of dissolved concentrations of Al, Fe, Mn, and Ti using 0.45 mm filtration as
an operational definition for ‘‘dissolved.’’ Sources of error were attributed to passage of
fine-grained clay particles through the filter.
Evidence from several field studies (Puls et al., 1992; Puls and Powell, 1992; Backhus
et al., 1993; McCarthy and Shevenell, 1998) indicates that field filtration does not effectively
remedy the problems associated with artifactual turbidity in samples. These and other
studies indicate that filtration may cause concentrations of some analytes to decrease
significantly, due to removal of colloidal particles that may be mobile under natural flow
conditions. Puls and Powell (1992) noted that 0.45 mm filtered samples collected with a
bailer had consistently lower As concentrations than samples obtained using low-flow-rate
pumping. They suggested that the difference may have been due to filter clogging from
excessive fines reducing the effective pore size of the filters or adsorption onto freshly
exposed surfaces of materials brought into suspension by bailing. Puls et al. (1992) found
that high-flow-rate pumping resulted in large differences in metals concentrations
between filtered and unfiltered samples, with neither value being representative of values
obtained using low-flow-rate sampling. Ambiguous sampling results found by McCarthy
and Shevenell (1998) were attributed to analytical values for metals obtained using low-

flow sampling that fell between filtered and unfiltered values from samples collected using
TABLE 5.1
Analytical Parameter Filtration Recommendations
Examples of parameters that may be field filtered
Alkalinity
Trace metals
Major cations and anions
Examples of parameters that should not be filtered
VOCs
TOC
TOX
Dissolved gases (e.g., DO and CO2)
‘‘Total’’ analyses (e.g., total arsenic)
Low molecular weight, highly soluble, and nonreactive constituents
Parameters for which ‘‘bulk matrix’’ determinations are required
Source: U.S. EPA, 1991.
Ground-Water Sample Pretreatment: Filtration and Preservation 135
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bailing or high-flow-rate pumping. Discrepancies in analytical values for some metals
(Al and Fe) exceeded an order of magnitude in this study. They determined that filtration
of turbid samples may have occluded pores in filters, leading to removal of colloidal
particles that may be representative of the load of mobile contaminants in ground water.
Puls and Barcelona (1989) also point to the removal of potentially mobile species as an
effect of filtration, indicating that filtration of ground-water samples for metals analysis
will not provide accurate information concerning the mobility of metal contaminants.
If the objective of a ground-water sampling program is to determine the exposure risk
of individuals who consume ground-water from private water supply wells, filtration of
those samples would not produce meaningful results. To make this type of exposure risk
determination, it is important to submit samples for analysis that are representative of
water as it is consumed, and, because most people do not have 0.45 mm filters at their

taps, unfiltered samples should be collected. In addition, it is important to remember that
MCL and MCLG values set for drinking-water standards are based on unfiltered samples.
Sources of Error and Bias in Filtration
The very act of filtration can introduce significant sources of error and bias into the results
obtained from analysis of sample filtrate (Braids et al., 1987). Some of these changes in
sample chemistry result from pressure changes in the sample, as well as sample contact
with filtration equipment and filter media. It is critical to evaluate the suitability of
filtration on a parameter-specific basis and to carefully select filtration methods,
equipment, and filtration media when developing site-specific filtration protocols to
minimize sample bias caused by filtration. The following sources of negative and positive
sample bias need to be considered:
. Potential for negative bias to occur due to adsorption of constituents from the
sample (U.S. EPA, 1991; Horowitz et al., 1996). For example, Puls and Powell
(1992) found that in-line polycarbonate filters adsorbed Cr onto the surface of
the filter medium, resulting in an underestimation of Cr concentrations in the
ground-water samples being filtered.
. Potential for positive bias to occur due to desorption or leaching of constituents
into the sample (Jay, 1985; Puls and Barcelona, 1989; Puls and Powell, 1992;
Horowitz et al., 1996). In the Puls and Powell (1992) study, K was observed to
leach from nylon filters that were not adequately preconditioned prior to use.
. Removal of particulates smaller than the original filter pore size due to filter
loading or clogging as filtered particles accumulate on the filter surface
(Danielsson, 1982; Laxen and Chandler, 1982) or variable particle size retention
characteristics (Sheldon, 1965; Sheldon and Sutcliffe, 1969).
. Removal of particulate matter with freshly exposed reactive surfaces, through
particle detachment or disaggregation, that may have sorbed hydrophobic,
weakly soluble, or strongly reactive contaminants from the dissolved phase (Puls
and Powell, 1992). This material itself may have been immobile prior to initiation
of sampling and mobilized by inappropriate sampling procedures.
. Removal of solids (metal oxides and hydroxides) that may have precipitated

during sample collection (particularly where purging or sampling methods that
may have agitated or aerated the water column are used) and any adsorbed
species that may associate with the precipitates. Such precipitation reactions can
occur within seconds or minutes (Reynolds, 1985; Grundl and Delwiche, 1992;
Puls et al., 1992), and the resultant solid phase possesses extremely high reactivity
136 The Essential Handbook of Ground-Water Sampling
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(high capacity and rapid kinetics) for many metal species (Puls and Powell, 1992).
Most metal adsorption rates are extremely rapid (Sawhney, 1966; Posselt et al.,
1968; Ferguson and Anderson, 1973; Anderson et al., 1975; Forbes et al., 1976;
Sparks et al., 1980; Benjamin and Leckie, 1981; Puls, 1986; Barrow et al., 1989).
Additionally, increased reaction rates are generally observed with increased
sample agitation.
. Exposure of anoxic or suboxic ground water (in which elevated levels of Fe
2'
are
typically present) to atmospheric conditions during filtration can also lead to
oxidation of samples, resulting in formation of colloidal precipitates and causing
removal of previously dissolved species (NCASI, 1982; EPRI, 1987; Puls and
Eychaner, 1990; Puls and Powell, 1992; Puls and Barcelona, 1996). The precipita-
tion of ferric hydroxide can result in the loss of dissolved metals due to rapid
adsorption or co-precipitation potentially affecting As, Cd, Cu, Pb, Ni, and Zn
(Kinniburgh et al., 1976; Gillham et al., 1983; Stoltzenburg and Nichols, 1985; Kent
and Payne, 1988).
. During sample filtration, care should be taken to minimize sample handling to the
extent possible to minimize the potential for aeration. If sample transfer vessels
are used, they should be filled slowly and filtration should be done carefully to
minimize sample turbulence and agitation. Stoltzenburg and Nichols (1986)
demonstrated that the use of sample transfer vessels during filtration imparted
significant positive bias for DO and significant negative bias for dissolved metal

concentrations. For this reason, the use of transfer vessels is discouraged. In-line
filtration is preferred because of the very low potential it poses for sample
chemical alteration.
Filtration Methods and Equipment
After a decision is made to field filter ground-water samples to meet DQOs for an
investigation, decisions must be made regarding selection of the most appropriate field
filtration method. The ground-water sample filtration process consists of several phases:
(1) selection of a filtration method; (2) selection of filter media (materials of construction,
surface area, and pore size); (3) filter preconditioning; and (4) implementation of field
filtration procedures. Information on each part of the process must be presented in detail
in the SAP to provide step-by-step guidance for sampling teams to implement in the field.
A wide variety of methods are available for field filtration of ground-water samples. In
general, filtration equipment can be divided into positive-pressure filtration and vacuum
(negative pressure) filtration methods, each with several different filtration medium
configurations. As discussed previously, ground-water samples undergo pressure changes
as they are brought from the saturated zone (where ground water is under pressure greater
than atmospheric pressure) to the surface (where it is under atmospheric pressure),
potentially resulting in changes in sample chemistry. The pressure change that occurs
when the sample is brought to the surface may cause changes in sample chemistry, which
include loss of dissolved gases and precipitation of dissolved constituents such as metals.
When handling samples during filtration operations, additional turbulence and mixing of
the sample with atmospheric air can cause aeration and oxidation of Fe
2'
to Fe
3'
.Fe
3'
rapidly precipitates as amorphous iron hydroxide and can adsorb other dissolved trace
metals (Stolzenburg and Nichols, 1986). Vacuum filtration methods further exacerbate
pressure changes and changes due to sample oxidation. For this reason, positive-pressure

filtration methods are preferred (Puls and Barcelona, 1989, 1996; U.S. EPA, 1991).
Ground-Water Sample Pretreatment: Filtration and Preservation 137
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Table 5.2 presents equipment options available for positive pressure and vacuum
filtration of ground-water samples.
When selecting a filtration method, the following criteria should be evaluated on a site-
by-site basis:
. Possible effect on sample integrity, considering the potential for the following to
occur:
a. Sample aeration, which may result in sample chemical alteration
b. Sample agitation, which may result in sample chemical alteration
c. Change in partial pressure of sample constituents resulting from application of
negative pressure to the sample during filtration
d. Sorptive losses of components from the sample onto the filter medium or
components of the filtration equipment (e.g., flasks, filter holders, etc.)
e. Leaching of components from the filter medium or components of the filtration
equipment into the sample
. Volume of sample to be filtered
. Chemical compatibility of the filter medium with ground-water sample chemistry
. Anticipated amount of suspended solids and the attendant effects of particulate
loading (reduction in effective filter pore size)
. Time required to filter samples. Short filtration times are recommended to
minimize the time available for chemical changes to occur in the sample
. Ease of use
. Availability of an appropriate medium in the desired filter pore size
. Filter surface area
. Use of disposable versus nondisposable equipment
. Ease of cleaning equipment if not disposable
. Potential for sample bias associated with ambient air contact during sample
filtration

. Cost, evaluating the costs associated with equipment purchase price, expendable
supplies and their disposal, time required for filtration, time required for
decontamination of nondisposable equipment, and QC measures
TABLE 5.2
Examples of Equipment Options for Positive-Pressure
and Vacuum Field Filtration of Ground-Water Samples
Positive-pressure filtration equipment
In-line capsules
Á
/Attached directly to a pumping device discharge hose
Á
/Attached to a pressurized transfer vessel
Á
/Attached to a pressurized bailer
Free-standing disk filter holders
Syringe filters
Zero headspace extraction vessels
Vacuum filtration equipment
Glass funnel support assembly
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The filtration method used for any given sampling program should be documented in
the site-specific SAP and should be consistent throughout the life of the sampling
program to permit comparison of data generated. If an improved method of filtration is
determined to be appropriate for a sampling program, the SAP should be revised in lieu
of continuing use of the existing filtration method. In this event, the effect on
comparability of data needs to be examined and quantified to allow proper data analysis
and interpretation. Statistical methods may need to be used to determine the significance
of any changes in data resulting from a change in filtration method.
Filtration equipment and filter media are available in a wide variety of materials of

construction. Materials of construction should be evaluated in conjunction with
parameters of interest being filtered with particular regard to minimizing sources of
sample bias, such as adsorption of metals from samples (negative bias) or desorption or
leaching of constituents into samples (positive bias). Materials of construction of both the
filter holder or support and the filter medium itself need to be carefully selected based on
compatibility with the analytes of interest (Puls and Barcelona, 1989). Filter holders that
are made of steel are subject to corrosion and may introduce artifactual metals into
samples. Glass surfaces may adsorb metals from samples.
Table 5.3 presents a summary of the most commonly used filtration media available for
field filtration of water samples. The potential for sample bias for these filter media
materials is variable, therefore, filter manufacturers should be consulted to determine
recommended applications for specific filtration media and for guidelines on the most
effective preconditioning procedures.
Large-diameter filter media (!47 mm) are recommended for ground-water sample
filtration (Puls and Barcelona, 1989). Because of the larger surface area of the filter,
problems of filter clogging and filter pore size reduction are minimized. High-capacity in-
line filters have relatively large filter media surface areas, which may exceed 750 cm
2
.
This can improve the efficiency of field sample filtration.
Filter Preconditioning
Filter media require proper preconditioning prior to sample filtration (Jay, 1985; U.S. EPA,
1995; Puls and Barcelona, 1996; ASTM, 2006a). The purposes of filter preconditioning are:
(1) to minimize positive sample bias associated with residues that may exist on the filter
surface or constituents that may leach from the filter, and (2) to create a uniform wetting
front across the entire surface of the filter to prevent channel flow through the filter and
increase the efficiency of the filter surface area. Preconditioning the filter medium may
not completely prevent sorptive losses from the sample as it passes through the filter
medium.
In most cases, filter preconditioning should be done at the wellhead immediately prior

to use (Puls and Barcelona, 1989). In some cases, filter preconditioning must be done in a
laboratory prior to use (e.g., GFuF filters must be baked prior to use). Some manufacturers
‘‘preclean’’ filters prior to sale. These filters are typically marked ‘‘precleaned’’ on filter
packaging and provide directions for any additional field preconditioning required prior
to filter use.
The procedure used to precondition the filter medium is determined by the following:
(1) the design of the filter (i.e., filter capsules or disks); (2) the material of construction of
the filter medium; (3) the configuration of the filtration equipment; and (4) the parameters
of concern for sample analysis. Filtration medium manufacturers’ instructions should be
followed prior to implementing any filter preconditioning protocols in the field to ensure
that proper methods are employed and to minimize potential bias of filtered samples.
Ground-Water Sample Pretreatment: Filtration and Preservation 139
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TABLE 5.3
Examples of Common Filter Media Used in Ground-Water Sampling
Filter Medium
Acrylic Copolymer Glass Fiber Mixed Cellullose Esters Nylon Polycarbonate Polyethersulfone Polypropylene
Analytes XX
Major ions X — — — X X X
Minor ions — — — — X X X
Trace metals X — — X X X X
Nutrients X — X — X — —
Organic compounds — X — — — X —
Filter effective area (cm
2
)
17 X X X XXX—
20 X X X XXX—
64 — X — ——X—
158 X X X — — X —

250 — — — — — X —
600 — — — X — X —
700 X — — — — — —
770 — — — X — X X
Pore size (mm)
0.1 — — X — X X —
0.2 — — X — X X —
0.45 X — X X X X —
1.0 X X X X — X X
5.0 X — X X — X X
Filter type
Flat disk X X X X X X —
Capsule X — — X X X X
Syringe X X — X — X —
Funnel X — X X X X —
Source: ASTM, 2006a.
140 The Essential Handbook of Ground-Water Sampling
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These instructions will specify filter-specific volumes of water or medium-specific
aqueous solutions to be used for optimum filter preconditioning.
The volume of water used in filter preconditioning is dependent on the surface area of
the filter and the medium’s ability to absorb liquid. Many filter media become fragile
when saturated and are highly subject to damage during handling. Therefore, saturated
filter media should be handled carefully and are best preconditioned immediately prior
to use in the field.
Disk filters (also known as plate filters) should be preconditioned as follows: (1) hold
the edge of the filter with filter forceps constructed of materials that are appropriate for
the analytes of interest; (2) saturate the entire filter disk with manufacturer-recom-
mended, medium-specific water (e.g., distilled water, deionized water, or sample water)
while holding the filter over a containment vessel (not the sample bottle or filter holder)

to catch all run-off; (3) then place the saturated filter on the appropriate filter stand or
holder in preparation for sample filtration; (4) complete assembly of the filtration
apparatus; (5) pass the recommended volume of water through the filter to complete
preconditioning; (6) discard preconditioning water; and (7) begin sample filtration using a
clean filtration containment vessel or flask. When preconditioning disk filters, care should
be taken not to perforate the filter. The filter medium should not be handled with
anything other than filter forceps. Otherwise, there may be a reduction in the porosity and
permeability of the filter medium. In addition, care should be taken to avoid exposure of
the filter medium to airborne particulates to minimize introduction of contaminants onto
the filter surface.
Preconditioning of capsule filters requires that liquid be passed through the filter prior
to sample filtration and collection. A volume of manufacturer-recommended, medium-
specific water (e.g., distilled water, deionized water, or sample water) should be passed
through the filter while holding the capsule upright, prior to sample collection. In general,
large-capacity capsule filters require that 1000 ml of water be passed through the filter
prior to sample collection, while small-capacity filters require approximately 500 ml of
water to be passed through the filter.
Sample Preservation
The second form of pretreatment of ground-water samples is physical and chemical
preservation. As described in ASTM Standard D 6517 (ASTM, 2006b), ground-water
samples are subject to unavoidable chemical, physical, and biological changes relative to
in situ conditions when samples are brought to ground surface during sample collection.
These changes result from exposure to ambient conditions, such as pressure, temperature,
ultraviolet radiation, atmospheric oxygen, and atmospheric contaminants, in addition to
any changes that may be imparted by the sampling device as discussed earlier in this
chapter.
Objectives of Sample Preservation
The fundamental objective of physical and chemical preservation of samples is to
minimize further changes in sample chemistry associated with sample collection and
handling from the moment the sample is placed in the sample container to the time it is

removed from the container for extraction or analysis in the laboratory. Sample
preservation methods are determined on a parameter-specific basis and must be specified
in the SAP prior to sample collection. Requirements for sample container type, storage
and shipping temperature, and chemical preservatives are specified in the analytical
method used for each individual parameter selected for analysis. Sampling team
Ground-Water Sample Pretreatment: Filtration and Preservation 141
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members are encouraged to speak with a laboratory representative prior to the sampling
event to ensure that the correct types and numbers of sample containers (along with a few
spares) and necessary chemical preservatives are shipped to the field site in sufficient
time for the scheduled sampling event. Sampling team members must also learn from the
laboratory what the parameter-specific holding times are (the amount of time that can
transpire from the moment the sample container is filled to the time the sample is
extracted or analyzed by the lab) for each parameter to ensure that samples are received
by the laboratory in a timely fashion. This is particularly critical when sampling is
conducted late in the work week for parameters that have a short holding time (e.g., 48 h).
Physical Preservation Methods
Physical preservation methods for ground-water samples include: (1) use of appropriate
sample containers for each parameter being analyzed; (2) use of appropriate packing and
packaging of samples to prevent damage during transport to the laboratory; and (3)
temperature control of samples during delivery to the laboratory. Sample containers are
specified on a parameter-specific basis by the chosen analytical method for the sampling
program (ASTM, U.S. EPA SW846 [U.S. EPA, 1996], AWWA Standard Methods [APHA,
AWWA, and WPCF, 1985]), as well as in Federal (40 CFR Part 136), state, and local
regulatory guidelines on ground-water sample collection and preservation. Containers
are specified with a number of design criteria in mind, to protect the integrity of the
analytes of interest, including shape, volume, gas tightness, materials of construction, use
of cap liners, and cap seal or thread design (Figure 5.3). Table 5.4 presents a summary of
some of the more common ground-water sample containers used. These required
containers are subject to change as methods are revised.

Sampling team members must also be aware that how they package sample containers
for either hand delivery to the laboratory or commercial shipping is a critical aspect of
physical preservation procedures. Field personnel should package and ship samples in
compliance with applicable shipping regulations as discussed in ASTM Standard D 6911
FIGURE 5.3
Sample containers vary in design based on the analytes to be measured in the sample. This example depicts
containers used for a full suite of parameters included in the RCRA detection monitoring program as collected
from one well.
142 The Essential Handbook of Ground-Water Sampling
© 2007 by Taylor & Francis Group, LLC
(ASTM, 2006c). Shipping regulations such as the U.S. Department of Transportation Title
49 Code of Federal Regulations Part 172 and the International Air Transport Association
(IATA) regulations should be consulted by sampling team members prior to a sampling
event where ground-water samples may be sufficiently contaminated to require
classification as dangerous goods for shipping purposes or where concentrated chemical
preservatives require shipment. These regulations will provide definitive instructions on
the correct packaging of regulated samples for shipment to the laboratory. Sample
containers should be shipped in a manner that will ensure that the samples are received
intact by the laboratory at the appropriate temperature as soon as possible after sample
collection, to permit sufficient time for the laboratory to perform the requested analyses
within the prescribed holding time for each analyte. Care must be taken by sampling
team members to ensure that sample containers are packed sufficiently tight within the
outer shipping container and to prevent movement during shipment that could result in
container breakage. It is also a good practice to avoid packing glass containers against
glass containers whenever possible (plastic against glass is a better configuration). Special
shock-absorbing sleeves and containers with a plastic coating have been designed to help
reduce the incidence of container breakage during shipment and handling. Use of bubble
wrap around containers can also minimize container breakage. Commercial carriers often
recommend that absorbent pads be placed in the bottom of sample shipping containers
TABLE 5.4

Examples of Frequently Used Containers for Ground-Water Samples
Parameter of Interest Container Volume (ml)
Inorganic tests Chloride P 125
Cyanide (total and amenable) P 1000
Nitrate P 125
Sulfate P 250
Sulfide P 500
Metals Cr
6'
P 500
Mercury P 500
Metals except Cr
6'
and Hg P 1000
Organic tests Acrolein and acrylonitrile G, PTFE ls 40
Benzidines G am, PTFE lc 1000
Chlorinated hydrocarbons G am, PTFE lc 1000
Dioxins and furans G am, PTFE lc 1000
Haloethers G am, PTFE lc 1000
Nitroaromatics and cyclic ketones G am, PTFE lc 1000
Nitrosamines G am, PTFE lc 1000
Oil and grease G am, wm 1000
Total organic carbon G am PTFE ls 40
Organochlorine pesticides G am, PTFE lc 1000
Organophosphorus pesticides G am, PTFE lc 1000
PCBs G am, PTFE lc 1000
Phenols G am, PTFE lc 1000
Phthalate esters G am, PTFE lc 1000
Polynuclear aromatic hydrocarbons G am, PTFE lc 1000
Purgeable aromatic hydrocarbons G, PTFE ls 40

Purgeable halocarbons G, PTFE ls 40
TOX G am, PTFE lc 250
Radiological tests Alpha, beta, and radium P 1000
Notes: P, high density PE; G, glass; G am, amber glass; wm, wide mouth; PTFE, polytetrafluorethylene
(Teflon
†
); lc, lined cap; ls, lined septum; TOX, total organic halides.
Source: U.S. EPA, 1996.
Ground-Water Sample Pretreatment: Filtration and Preservation 143
© 2007 by Taylor & Francis Group, LLC
and on the top of sample containers after the shipper is filled, to absorb shock during
transit.
Another important consideration during handling of samples in the field, following
collection and during transport to the laboratory, is temperature control. Many
parameters require that samples be stored at 48C in the field between sample collection
locations, during sample shipment (or delivery if by hand), and upon arrival at the
laboratory. (48C is the temperature at which water is at its maximum density and is most
chemically stable.) To accomplish this, sample temperatures should be lowered
immediately after sample containers have been filled, labeled, and had any required
security seals affixed to them.
Cooling can be accomplished using on-site refrigeration systems if they are available in
the field or, more commonly, using wet (natural) ice. Wet ice is the preferred method to
cool samples to 48C. It is inexpensive, readily available, and will not get samples so cold
that they will freeze. Wet ice will, however, require replenishment throughout the day to
maintain sample temperatures, especially when sampling in warm ambient temperatures.
Wet ice should be double bagged to prevent leakage into the shipping container as the
ice melts.
Reusable chemical ice packs (also called blue ice packs) (Figure 5.4) are neither suitable
for lowering sample temperatures to 48Cfromin situ temperatures nor suitable for
maintaining sample temperatures at 48C during field handling and shipment. Thus, they

are not recommended for use during ground-water sampling (Kent and Payne, 1988;
ASTM, 2006b). There is also some concern about what chemicals may be released into the
shipping container should a chemical ice pack be punctured or leak during sample
shipment.
Dry ice is sometimes specified for use for sample cooling. Unfortunately, when dry ice
is used, samples often become too cold and smaller volume containers commonly freeze,
resulting in container breakage and sample loss. Dry ice is also relatively expensive
and difficult to obtain in the field. It requires special handling procedures in the field and
is a regulated substance under shipping regulations.
FIGURE 5.4
Reusable chemical ice packs, such as those used in this sample shuttle, are not recommended for ground-water
sampling because they are incapable of achieving the desired sample temperature (48C) in most cases and they
may contribute contaminants to samples if they rupture or leak in transit to the laboratory.
144 The Essential Handbook of Ground-Water Sampling
© 2007 by Taylor & Francis Group, LLC
To verify appropriate temperature control of samples, it is recommended that samplers
include a QC sample referred to as a temperature blank (ASTM, 2006b) in the same
shipping container as the ground-water samples. Upon receipt at the laboratory, a
laboratory representative will check the temperature blank to determine whether samples
were approximately preserved with respect to temperature. If the temperature blank is
not at the required temperature (4 928C), the laboratory representative will contact the
sampling team to notify them of the sample arrival temperature and to determine an
appropriate course of action.
Chemical Preservation Methods
Chemical preservation is an important field procedure that samplers must implement to
ensure that chemical change in samples is minimized during sample handling and
shipment. Chemical preservation involves the addition of one or more chemicals (reagent
grade or better) to the ground-water sample during sample collection. Chemicals can be
used to adjust sample pH to keep constituents in solution or to inhibit microbial
degradation of samples. Chemical preservatives are specified by each analytical method

for each parameter and the preservatives are typically provided by the laboratory.
Table 5.5 provides examples of common chemical preservatives used for ground-water
samples.
Ground-water samples can be chemically preserved in one of several ways: (1) titration
of pH-adjusting compounds (e.g., nitric acid) while monitoring pH change with a pH
meter or narrow-range litmus paper; (2) addition of a fixed volume of liquid preservative
(e.g., sulfuric acid contained in glass vials or ampoules) to the sample container; (3)
addition of a fixed amount of pelletized preservative (e.g., sodium hydroxide) to the
sample container; or (4) placement of preservatives in empty sample containers prior to
shipment of the containers to the field (i.e., prepreserved sample containers). Titration
methods for sample preservation, while theoretically a valid approach, are not always
practical under field conditions where samplers are required to handle large volumes of
concentrated preservatives and work with glass titration apparatus under less than ideal
conditions (wind, rain, dust, etc.). A modified version of this method is to use calibrated
dropper bottles of preservative rather than glass burettes for titration. This ensures that
the correct preservative is titrated into the sample while monitoring pH changes, but in a
safer fashion in the field.
Vials or ampoules of preservatives are commonly used for sample preservation in the
field (Figure 5.5). The laboratory provides the vials or ampoules containing a fixed
volume of the required preservative for each sample container requiring chemical
preservation. The sampling team should be provided with directions on which
preservative must be added to which container on a parameter-specific basis, as well
as guidance on whether the preservative should be added to the container before or after
filling. One common error made by sampling teams is to assume that the amount of
preservative provided in vials or ampoules will always be sufficient to reach the required
end pH for the analyte. This is not a safe assumption, especially in situations where
ground-water pH may be abnormally high or low based on contaminant chemistry or the
natural pH of formation-quality water. For this reason, it is essential that both an initial
and final pH measurement be taken to check for pH anomalies and to ensure that the
required end pH for the sample has been reached (ASTM, 2006b). To take these

measurements, a small aliquot of sample should be decanted into another container (e.g.,
a clean empty VOC vial without preservative that will not be used for sample collection
or a clean small-volume beaker) and the pH measured using either a calibrated pH probe
or narrow-range litmus paper. If the sample pH is not at the required endpoint, additional
Ground-Water Sample Pretreatment: Filtration and Preservation 145
© 2007 by Taylor & Francis Group, LLC
preservative must be added until it is reached. For this reason, sampling teams must ask
the laboratory to provide additional preservative, preferably in a vial that can be resealed
if only a few extra drops of preservative are required. Sampling teams must resist the
temptation to double the preservative required by the method simply for the sake of
convenience. This can result in the samples becoming a corrosive liquid for shipping
TABLE 5.5
Examples of Commonly Used Ground-Water Sample Chemical Preservatives and Holding Times
Parameter of Interest Preservative
Holding
Time
Inorganic tests Chloride Cool to 48C ASAP
Cyanide (total and
amenable)
Cool to 48C
a
; if oxidizing agents present, add 5 ml
0.1 N NaAsO
2
/l or 0.06 g ascorbic acid/l; pH ! 12
with 50) NaOH
14 days
Nitrate Cool to 48C; boric acid for method 9210 ASAP
Sulfate Cool to 48C ASAP
Sulfide Cool to 48C; pH ! 9 with NaOH; Zn acetate; no

headspace
7 days
Metals Chromium
6'
Cool to 48C 1 day
Mercury pH B 2 HNO3 28 days
Metals Except Cr
6'
and
Hg
pH B 2 HNO3 6 months
Organic tests Acrolien and acrylonitrile Cool to 48C; pH 4 Á
/5
a
Na2S2O3 14 days
Benzidines Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Chlorinated hydrocarbons Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Dioxins and furans Cool to 48C; 0.008) Na2S2O3
a
30e/45ae
Haloethers Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Nitroaromatics and cyclic
ketones
Cool to 48C; 0.008) Na2S2O3

a
dark 7e/40ae
Nitrosamines Cool to 48C; 0.008) Na2S2O3
a
dark 7e/40ae
Oil and grease Cool to 48C; pH B 2 HCl ASAP
Organic carbon, total
(TOC)
Cool to 48C; pH B 2 H2SO4 or HCl; dark ASAP
Organochlorine pesticides Cool to 48C 7e/40ae
Organophosphorous
pesticides
Cool to 48C; 0.008) Na2S2O3
a
,pH5Á/8 7e/40ae
PCBs Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Phenols Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Phthalate esters Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Polynuclear aromatic
hydrocarbons
Cool to 48C; 0.008) Na2S2O3
a
7e/40ae
Purgeable aromatic

hydrocarbons
Cool to 48C; Na2S2O3 pH B 2 HCl or H2SO4 or
NAHSO4
14 days
Purgeable aromatic
halocarbons
Cool to 48C; Na2S2O3 pH B 2 HCl or H2SO4 or
NAHSO4
14 days
TOX Cool to 48C; pH B 2 H2SO4; dark; no headspace 28 days
Radiological
tests
Alpha, beta, and radium pH B 2 HNO3 6 months
a
Only add a reducing agent if the sample contains free or combined chlorine. A field test kit needs to be used for
this determination.
7e: sample extraction must be completed within 7 days of sample collection.
40ae: analysis must be completed within 40 days after sample extraction.
Note: ASAP, analysis should be performed as soon as possible. Samplers should discuss with the laboratory how
ASAP is to be interpreted on a project-specific basis. In many cases, if the parameter can be analyzed with
accuracy and precision in the field that is preferred. Otherwise, many laboratories use a 24 h holding time.
Source: U.S. EPA, 1996.
146 The Essential Handbook of Ground-Water Sampling
© 2007 by Taylor & Francis Group, LLC
purposes as described in ASTM Standard D 6911 (ASTM, 2006c) and can detrimentally
affect the chemistry of the sample.
It is generally accepted that the sample dilution attributed to the addition of chemical
preservatives should be limited to a maximum of 0.5) (ASTM, 2006b). The pH of
samples should be checked upon arrival at the laboratory to ensure that appropriate
sample preservation procedures were implemented in the field. If the pH is not where it is

required to be, the laboratory will consider the sample to be inappropriately preserved
and the sampling team will be contacted to discuss an appropriate course of action.
The most convenient method of chemically preserving ground-water samples is to use
prepreserved containers (Figure 5.6). Prepreserved containers are either purchased by the
laboratory already prepared or they can be prepared by the laboratory. These containers
hold a fixed volume of the parameter-specific preservative and are shipped to the
sampling team with information about which preservatives have been added to which
containers. The advantages of this method of sample preservation are: (1) the sampling
team does not have to handle preservatives; (2) theoretically, no errors associated with
adding an incorrect preservative to a sample can be made by field personnel; and (3) time
savings. There are also several limitations to using prepreserved containers. As when
using vials and ampoules to add preservative to samples, the volume of preservative is
fixed. Thus, difficulties can arise in the field if field verification of end pH determines that
the volume of preservative provided in the container is insufficient. In this situation,
sampling teams must have available additional preservative (the same as that used to
prepare the container), so the required end pH can be achieved prior to shipping the
sample to the laboratory. This may be impossible for the laboratory to provide if they
have purchased prepreserved containers from a supplier. From a practical perspective,
FIGURE 5.5
Chemical preservation using vials.
Ground-Water Sample Pretreatment: Filtration and Preservation 147
© 2007 by Taylor & Francis Group, LLC
one common complaint of sampling team members using prepreserved containers is that
it is easy to lose preservative from the container, either through accidentally knocking
over the container during filling or through overfilling, especially when attempting to
collect zero-headspace samples in 40 ml vials.
Another concern over prepreserved containers is related to the fact that concentrated
preservatives may react with the empty container during storage prior to sample
collection. For example, nitric acid, when in long-term storage in a high-density PE
container, will chemically react with the container, resulting in alteration of the container

walls (evidenced by orange staining inside the container) and, ultimately, as the authors
have observed, failure of the container (the plastic container will crack linearly when
squeezed). In glass containers stored in hot ambient conditions, acid preservatives will
commonly evaporate to form an acid vapor, which is released to ambient air when the
container is opened, leaving little preservative available to lower the pH of the sample
and creating a breathing hazard for samplers. In addition, the vapor can deteriorate
sample container lid threads — a problem that may not be detected until the container is
taken into the field and the cap crumbles into pieces when it is removed for container
filling. For these reasons, some sampling protocols do not permit the use of prepreserved
containers. In other programs, prepreserved containers are allowed but with storage time
restrictions that can vary from days to hours. It is recommended that sampling teams
work with the laboratory and regulatory agencies during the planning phases of the
ground-water sampling program to determine how long a prepreserved container can
remain in storage prior to sample collection.
FIGURE 5.6
A prepreserved container used for chemical preservation of samples.
148 The Essential Handbook of Ground-Water Sampling
© 2007 by Taylor & Francis Group, LLC
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© 2007 by Taylor & Francis Group, LLC
© 2007 by Taylor & Francis Group, LLC