chapter 5c sampling effort and collection methods:...

SAMPLING EFFORT AND COLLECTION METHODS 323

ASTM (1995) in standard D 4823 describes many other types of core samplers. The mostcommon sampler is the open tube sampler with a core catcher. This sampler is commonly used inshallow waters where it is manually pushed into the sediment. When the desired penetration depthis reached, the sampler is carefully withdrawn. A leaf core catcher is commonly used to help retainthe sample in the corer (Figure 5.73). The leaves separate and fold against the inside walls of thesampler when the corer penetrates the sediment. The leaves fold closed when the sampler iswithdrawn, holding the sample in the corer. Plastic liners are also commonly used inside the sampler,simplifying core removal from the corer. The liners usually have plastic end caps that can be placedon the liner ends, holding the cores inside until analyses. These conventional core samplers aremost effective with clayey sediments. Sandy sediments tend to easily wash out of most corers uponretrieval, irrespective of the core catcher used. ASTM (1995) mentions excavating around a coresampler and sliding a flat plate under the bottom of the corer before retrieval in shallow water tocapture most of the sample. Forestry Suppliers, Inc. sells the Wildco hand core sediment samplerthat is 2" in diameter and 20" long, made of stainless steel with a plastic core liner tube and eggshellcatcher (catalog #77258 for about $340). Extra plastic liners are also available (catalog # 77260)for about $12 each. They also sell stainless steel liners and core catchers (catalog #77303 for thestainless steel liner for about $70 each and catalog #77304 for the stainless steel eggshell samplecatcher for about $40 each).

Corer samplers also have limitations in some situations (ASTM 1991a). Most corers do notwork well in sandy sediments or in extremely soft (high water content) sediments; dredge samplersor diver-collected material remain the only current alternatives. In general, corers collect lesssediment than dredge samplers that may provide inadequate quantities for some toxicity studies.Small cores tend to increase bow (pressure) waves (disturbance of surface sediments) and com-paction, thus altering the vertical profile. However, these corers provide better confidence limitsand spatial information when multiple cores are obtained (EPA 1983b; Elliott and Drake 1981).As shown by Rutledge and Fleeger (1988) and others, care must be taken in subsampling fromcore samples, since surface sediments might be disrupted even in hand-held core collection. Theyrecommend subsampling in situ or homogenizing core sections before subsampling. Slowing thevelocity of entry of coring equipment also reduces vertical disturbance. Samples are frequently ofa mixed depth, but a 2-cm sample is recommended and the most common depth obtained, althoughdepths up to 40 ft have been used in some dredging studies.

For dredging, remediation, and/or historical pollution studies, it is sometimes necessary toobtain cores of depths up to several meters. This often requires the use of vibracores that aresomewhat destructive to sediment integrity but are often the only feasible alternative for deep orhard sediment sampling (Figures 5.74 through 5.76). In most studies of sediment toxicity, it isadvantageous to subsample the inner core area (not contacting the sampler) since this area is mostlikely to have maintained its integrity and depth profile and not be contaminated by the sampler.Subsamples from the depositional layer of concern, for example, the top 1 or 2 cm, should becollected with a nonreactive sampling tool, such as a polytetrafluoroethylene-lined calibration scoop

Figure 5.73 Leaf core catcher. (From American Society for Testingand Materials). ASTM Standards on EnvironmentalSampling. ASTM Pub Code No. 03-418095-38.ASTM, Philadelphia. 1995. Copyright ASTM.Reprinted with permission.)

LEAVES

324 STORMWATER EFFECTS HANDBOOK

Figure 5.74 Vibratory core collection. Figure 5.75 Lowering vibratory corer.

(Long and Buchman 1989). Subsamples are placed in a nonreactive container and mixed until texture and color appear uniform. Due to the large volume of sediment that is often needed for toxicity or bioaccumulation tests and chemical analyses, it might not be possible to use subsampled cores because of sample size limitations. In those situations, the investigator should be aware of the above considerations and their possible effect on test results as they relate to in situ conditions.

Once sediment samples are collected, it is important, in most situations, to reduce the possibility of sediment oxidation. The majority of fine-grained sediments that are of concern in toxicity assessments are anaerobic below the top few millimeters (Carlton and Klug 1990), and any introduction of oxygen will likely alter the valence state of many ionic chemicals. This alteration may significantly change the bioavailability and toxicity of the sample. To protect sediments from oxygenation, the use of a glove box or bags with an inert gas supply for subsampling and processing, e.g., preparation of sediments for centrifugation, might be necessary.

While coring is preferred for maintaining a sediment’s vertical integrity, care must be taken to reduce the possibility of spillage. Sediment cores should be stoppered immediately upon retrieval

to prevent accidental loss of sediment. During all handling procedures, cores should be kept in an upright position as a general precaution against disturbance of the sediment. This is particularly important to prevent mixing of the uppermost part of the sediment column, which usually consists of very fine, soft, and unconsolidated material. The intact core samples (liners) should also be capped or stoppered and taped closed, secured in an upright position (e.g., rack), and labeled with appropriate information regarding sampling site, location, sample number and/or identification, time and date of collection, method of collection, and name or initials of the collector. When using clear plastic liners, the appearance of each sediment core should be recorded prior to any subsampling, along with other descriptive features such as the length of the core, thickness of various

Figure 5.76 Emptying vibratory corer. sediment units, occurrence of fauna, presence of


oil or noticeable odor, and sediment color, texture, and structure (Environment Canada 1994; Mudroch and Azcue 1995; Figure 5.77).

Once samples are collected, some form of subsampling and/or compositing is often performed. Removal of a portion of the collected sediment from the grab sampler (i.e., subsampling) can be performed using a spoon or scoop made of inert, noncontaminating material (e.g., Teflon, titanium, or high-quality stainless steel). It is recommended when subsampling to exclude sediment that is in direct contact with the sides of the grab sampler as a general precaution against any potential contamination from the device. Each subsample may be placed into a separate clean, prelabeled container. As a general rule, each labeled sample container must be tightly sealed and the air excluded. However, if Figure 5.77 Vertical layers of a sliced core.

the sample is to be frozen, it is advisable to leave a small amount of headspace in the container to accommodate expansion and avoid breakage.

Compositing of core samples or subsamples, if necessary, can be done in the field or laboratory, such as by using a drill auger mixer shown in Figure 5.78. The quality of the core sample must be acceptable and only sediment depths with similar stratigraphy should be combined. Although there might be occasions when it is desirable to composite incremental core depths, it is recommended that only horizons of similar stratigraphy be composited. Depending on the study objectives and desired sampling resolution, individual horizons within a single core can be homogenized to create one or more depth composites for that core, or corresponding horizons from two or more cores might be composited. Thorough homogenization of the composite sample, by hand or using a mechanical mixer, is recommended prior to analysis or testing.

The type of sediment characterization needed will depend on the study objectives and the contaminants of concern; however, a minimum set of parameters should be included which are known to influence toxicity and will aid data interpretation. At a minimum, the following physical and chemical characterization of sediment is recommended: total solids (dry weight), total organic carbon (TOC), acid volatile sulfides (AVS) (when metals are of concern), ammonia, and grain size fractionation. The following parameters are also frequently useful in characterization and data interpretation of contaminant effects: pH, ORP (oxidation–reduction potential), temperature, salinity-conductivity, hardness, total volatile solids (ash free weight), nitrogen and phosphorus species, cation exchange activity (CEC), sediment or suspended solids biochemical oxygen demand (BOD), and/or chemical oxygen demand (COD). Many of the characterization methods have been based on analytical techniques for soils, wastewaters, and waters, and the literature should be consulted for further information (EPA 1977; Black 1965; Figure 5.78 Mixing sediment with a drill auger. USGS 1969; ASTM 1989; Page et al. 1982).


Interstitial Water and Hyporheic Zone Sampling

Interstitial water (pore water) is defined as the water occupying space between sediment or soil particles and is often isolated to provide either a matrix for toxicity testing or provide an indication of the concentration and partitioning of contaminants within the sediment matrix. U.S. EPA sediment quality criteria are based on the assumption that the primary route of exposure to benthic organisms is via the interstitial water (Di Toro et al. 1991). However, this route of exposure does not include uptake from ingestion of contaminated sediment particles. In addition, contaminants in interstitial waters can be transported into overlying waters through diffusion, bioturbation, and resuspension processes (Van Rees et al. 1991). The usefulness of interstitial water sampling for determining chemical contamination and/or toxicity will depend on the study objectives and nature of the sediments at the study site. Sediments that are either very large grain-sized (such as gravel or cobble) or hard, compacted clays will likely not have interstitial waters that are significantly contaminated. Therefore, sampling of interstitial waters should be restricted to sediments ranging from sandy to noncompacted clays. Interstitial waters from depositional zones containing smallersized sediments (clays) are usually the most contaminated.

Frequently, surface waters and groundwaters intermix via upwelling or downwelling transition zones (TZ). The ecosystem associated with this transition zone is sometimes referred to as the hyporheic zone or hyporheous. It can be a very important zone for many reasons: provides essential habitat and refugia for micro-, meio-, and macrofauna or flora; affects contaminant attenuation, removal, or transport; cycles nutrients and carbon; and provides trophic links between the microbes and invertebrates and their macrofaunal predators (Duncan 1999). To date these zones have largely been ignored in environmental contaminant assessments and conceptual models, even though they are quite common. They provide a challenge in that their assessment requires collaboration of hydrogeologists, hydrologists, ecologists, chemists, and toxicologists.

The biological and physicochemical conditions within the groundwater and surface water are different, and hence may affect the partitioning (e.g., bound or freely dissolved), mobility, and bioavailability of sediment-associated contaminants. For example, changes in pH may affect the binding of metals, whereas the rate and extent of microbial processing of sediment organic matter may affect the partitioning of persistent organic contaminants. Upwelling zones (where groundwater and interstitial water move up toward surface water) are generally anoxic, with low pH. Anaerobic microbial processes dominate and may include reductions, denitrification, ammonification, and methanogenesis. Dissolved organic carbon (DOC) is of low quality and species diversity is often quite low in upwelling zones. However, benthic consumers are attracted to this habitat. Downwelling zones (the downward movement of surface water into the stream bed) are generally higher in oxygen content and pH. Aerobic microbial processes such as oxidation and nitrification are dominant. DOC quality, species diversity, and productivity are high in downwelling habitats. The hydrological interface between upwelling groundwater and downwelling surface water within the stream bed contains large gradients for a variety of physicochemical parameters (e.g., temperature, dissolved oxygen, pH, and pE). Previous studies have shown that organic contaminant and metals concentrations can vary over several orders of magnitude (Benner 1995).

There are several scenarios in which data on groundwater–surface water interactions would be useful in evaluations of the fate and dynamics of sediment contaminants and the in situ exposure of biota. Upwelling groundwater can affect benthos and surface water biota if either or both the groundwater and sediments are contaminated. Aqueous phase chemicals (e.g., freely dissolved, colloidbound) in the upward flowing groundwater and/or the mobilization of sediment-bound contaminants by upwelling groundwater are the potential inputs to the surficial environs under these conditions. Downwelling surface water can affect benthic, hyporheic, and phreatic (groundwater-associated) biota if either or both the surface water and sediments are contaminated. Under such conditions, the potential exists for the transport of sediment contaminants to deep layers within the stream bed and the


contamination of groundwater by the downward-flowing contaminant load. We have observed this at sites contaminated by PCBs and chlorinated benzenes (Greenberg and Burton 1999).

Selection of Measurement Methods for Interstitial Water

Isolation of sediment interstitial water can be accomplished by a wide variety of methods, which can be grouped as laboratory or field (in situ) based. The common laboratory-based methods can be categorized as (1) centrifugation, (2) pressurization, or (3) suction. Field-based methods include suction and “peepers” (for reviews, see Adams et al. 1991; ASTM 1994; Burton et al. 2001; Environment Canada 1994). Peepers are small chambers with membrane or mesh walls, which are buried in sediments, and surrounding interstitial water then equilibrates within the chamber. Chambers are typically retrieved from 2 to 20 days after deployment.

It is important to work with the analytical and toxicity testing laboratories to determine the least amount of sample needed, because of the difficulty of obtaining large amounts of interstitial water for analyses. As an example, the use of an anodic stripping voltammeter is suitable for direct analyses (undigested) of heavy metals in interstitial water using only about 5 mL of water for several metals (at least copper and lead) simultaneously, instead of about 50 mL typically required. Organic analyses may be conducted with about 250 mL of water, using the modified methods described in Chapter 6, instead of the typically required 1-L sample sizes, but with loss of sensitivity. The use of an automated water analyzer (such as the TrAAcs 2000 analyzer from Bran+Luebbe) can dramatically reduce the water volume needed for conventional nutrient analyses. Ion chromatography also requires only a very small amount of sample for complete cation and anion analyses. Microtox, from Azur Environmental, is also a very useful indicator of toxicity and requires only a very small amount of sample (about 1 mL). Bacteria tests can also be conducted using small sample volumes (using methods from IDEXX, Inc., for example), especially if the bacteria densities are high, as is likely in contaminated urban streams, allowing dilution of the samples.

When relatively large volumes of water are required (such as 20 mL or greater), only grab and core sampling with subsequent centrifugation and sediment squeezing methods are typically used. Other methods such as suction and in situ samplers do not easily produce sufficient volumes for most required analyses. However, larger-sized peepers (500 mL volume) have been used for collecting samples for chemical analyses and for exposing test organisms in situ (Burton 1992a,b; Sarda and Burton 1995; see also Chapter 6).

Most sediment collection and processing methods have been shown to alter interstitial water chemistry (e.g., Schults et al. 1992; Bufflap and Allen 1995a,b; Sarda and Burton 1995) and, therefore, can potentially alter contaminant bioavailability and toxicity. Some important interstitial water constituents, e.g., dissolved organic carbon, dimethylsulfide, ammonia, major cations, and trace metals can be significantly altered by the collection method (e.g., Martin and McCorkle 1993; Carignan et al. 1994; Bufflap and Allen 1995a,b; Sarda and Burton 1995). Increased sample handling associated with methods such as grab or core sampling and centrifugation, squeezing, or suction may cause significant increases in key constituents, such as ammonia, sulfide, and DOC concentrations, as compared to those collected via in situ “peepers” or core-port suction. Other constituents, such as salinity, dissolved inorganic carbon, sulfide, and sulfate, might not be affected by collection, providing oxidation is prevented. If sediments are anoxic, as most depositional sediments are, all steps involved in sample processing should be conducted in inert atmospheres or by limited contact with the atmosphere to prevent oxidation (and subsequent sorption/precipitation) of reduced species. When anoxic sediments are exposed to air, volatile sulfides will be lost which may increase the availability of sulfide-bound metals. In addition, iron and manganese oxyhydroxides are quickly formed which readily complex with trace metals, thus altering metalsrelated toxicity (e.g., Bray et al. 1973; Troup et al. 1974; Burton 1991). There is no need for maintaining anoxic processing conditions when the study objectives are concerned only with exposures to oxic sediments, or if target contaminants are unaffected by oxidation in short-term


toxicity or bioaccumulation testing. For example, often studies of dredged material toxicity do not consider ammonia-related toxicity, and oxidation is actually promoted to remove ammonia from overlying waters of the toxicity test beakers.

Immediate collection and analysis of interstitial water is recommended since chemical changes might occur even when sediments are stored for short periods (e.g., 24 h) at in situ temperatures (Sarda and Burton 1995). Coagulation and precipitation of the humic material was noted when interstitial water was stored at 4°C for more than 1 week. Oxidation of reduced arsenic species in interstitial water of stored sediments was unaffected for up to 6 weeks when samples were acidified and kept near 0°C, without deoxygenation. When samples were not acidified, deoxygenation was necessary. Others have recommended interstitial waters be frozen after extraction, prior to toxicity testing, to prevent changes, but others have recommended against freezing samples that will undergo toxicity testing. The optimal collection method will depend upon the purpose of the sample (e.g., acidification for metal analysis and not toxicity testing), characteristics of the sediment, and the contaminants of concern. Sediments that are highly contaminated with strongly nonpolar organics (such as PCBs) are not likely to change in toxicity during storage.

The conditions for isolation of interstitial waters by centrifugation have varied considerably. For toxicity testing, interstitial waters have been isolated over a range of centrifugal forces and temperature ranges (Ankley and Schubauer-Berigan 1994; Schults et al. 1992) with centrifuge bottles of various compositions. When centrifugation followed by filtration has been compared with in situ dialysis, higher speed centrifugation followed by filtration with 0.2 membrane filters has produced results that were more similar for metals and organic carbon. Centrifugation at low speeds or use of a larger pore size filtration membrane (e.g., 45 µm mesh) will result in retention of dissolved contaminants, colloidal materials, and aquatic bacteria in the pore water sample. Highspeed centrifugation (e.g., 10,000 × g) is necessary to remove colloids and dispersible clays (Ankley and Schubauer-Berigan 1994). Typically, toxicity is reduced with high-speed centrifugation or filtration due to the removal of particle-associated contaminants (Ankley and Schubauer-Berigan 1994; Schults et al. 1992; Bufflap and Allen 1995a). While the duration of the centrifugation has been variable, 30 min is relatively common. The temperature for the centrifugation should reflect the ambient temperature of collection to ensure that the equilibrium between particles and interstitial water is not shifted.

Filtration through glass fiber or polycarbonate membranes may cause the loss of some dissolved metals and organics (Schults et al. 1992). If filtration is employed, a nonfiltered sample should also be tested for toxicity and contaminant concentrations. The effects of centrifugation speed, filtration, and oxic conditions on some chemical concentrations in interstitial waters have been well documented (e.g., Ankley and Schubauer-Berigan 1994; Schults et al. 1992; Bufflap and Allen 1995b; Bray et al. 1973). It is recommended that, for routine toxicity testing of interstitial waters, sediments should be centrifuged at 10,000 × g for a 30-min period at 4°C. It is difficult to collect interstitial water from sediments that are predominantly coarse sand. A modified centrifuge bottle has been developed with an internal filter which can recover 75% of the interstitial water as compared to 25 to 30% from squeezing.

Polytetrafluoroethylene (PTF) bottles will collapse at 3000 g but have been used successfully up to 2500 g when filled to 80% of capacity. Isolation of interstitial water in this case should be at the temperature of collection, at a slower speed of 2500 × g for 30 min. This material will contain colloidal material as well as dissolved compounds. At low centrifugation speeds, without filtration, removal of the colloids may not be possible. The influence of dissolved and colloidal organic carbon may be estimated by measuring the organic carbon content. If small volumes of water are required for testing, higher speed centrifugation can be performed with glass tubes (up to 10,000 × g). If metal analysis of toxicity is not a concern, then high-speed centrifugation in stainless steel centrifuge tubes is an option. When working with samples contaminated with organics, efforts should be made to reduce sample exposure to light to reduce photo-related degradation or alteration of any potentially toxic compounds. This can be accomplished by using amber bottles and yellow lights.


Isolation of interstitial water by squeezing has been performed with a variety of procedures. In all cases, the interstitial water is passed through a filter that is a part of the apparatus. Filters have different sorptive capacities for different compounds. Numerous studies have shown filters reduce toxicity and contaminant concentrations by retaining contaminant-associated particles and also by contaminant sorption onto the filter matrix (Schults et al. 1992; Bray et al. 1973; Troup et al. 1974; Sasson-Brickson and Burton 1991). The characteristics of filters and the filtering apparatus should be carefully considered. Squeezing has been shown to produce a number of artifacts due to shifts in equilibrium from pressure, temperature, and gradient changes (e.g., Schults et al. 1992; Troup et al. 1974; Mangelsdorf et al. 1969; Fanning and Pilson 1971; Figure 5.79). Squeezing can affect the electrolyte con- Figure 5.79 Pore water squeezer — stainless steel centration in the interstitial water with a decrease with Teflon liner.

near the end of the squeezing process. It is therefore recommended that moderate pressures be used with electrolyte (conductivity) monitoring during extraction. Significant alterations to interstitial water composition occurred when squeezing was conducted at temperatures different from ambient (e.g., Mangelsdorf et al. 1969). Other sources of alteration of interstitial water when using the squeezing method are contamination from overlying water, internal mixing of interstitial water during extrusion, and solid-solution reactions as interstitial water is expressed through the overlying sediment. As interstitial waters are displaced into upper sediment zones, they come in contact with solids with which they are not in equilibrium. This intermixing causes solid-solution reactions to occur. The chemistry of the sample may be altered due to the fast kinetics (minutes to hours) of these reactions. Most interstitial water species are out of metastable equilibrium with overlying sediments and are rapidly transformed, as observed with ammonia and trace metals. Bollinger et al. found elevated levels of several ions and dissolved organic carbon in squeezed samples as compared to samples collected by peepers. The magnitude of the artifact will depend on the element, sediment characteristics, and redox potential. It is unlikely that reactive species gradients can be established via squeezing of sediment cores.

Many studies have demonstrated the usefulness of in situ collection methods (e.g., Barnes 1973; Belzile et al. 1989; Bottomley and Bayly 1984; Buddensiek et al. 1990; Howes et al. 1985; Jahnke 1988; Mayer 1976; Murray and Grundmanis 1980; Sayles et al. 1973; and Whiticar 1982). These methods of interstitial water collection are superior to more traditional methods in that they are less likely to alter the chemistry of the sample. The principal methods of interstitial water collection are through the use of peepers (e.g., Bufflap and Allen 1995a,b; Carignan 1984; Bottomley and Bayly 1984) or in situ suction techniques. These methods have the greatest likelihood of maintaining in situ conditions and have been used to sample dissolved gases (Sarda and Burton 1995) and volatile organic compounds.

Suction using an aquarium air stone recovered up to 1500 mL from 4 L of sediment suctioned in an anoxic environment (Galli 1997). Hand vacuuming using an aquarium stone has shown to be an effective method of collecting interstitial water (Sarda and Burton 1995). The air stone is attached to a 50-mL syringe via plastic tubing. The stone is inserted in the sediment to the desired depth and then suction applied. Clogging of the air stone is a problem in some sediments; however, it is effective in most tested. The collection system can be purged of oxygen prior to leaving the laboratory. Ammonia concentrations in water obtained by this system were similar to those collected


Figure 5.80 Disassembled small-volume, high-reso- Figure 5.81 Small-volume peeper assembly showlution peepers. ing 75-mm nylon screening.

with in situ peepers (Sarda and Burton 1995). Problems common to suction methods are loss of equilibration between the interstitial water and the solids, filter clogging, and oxidation. However, in situ suction or suction via core ports has been shown to accurately define small gradients of some sediment-associated compounds, including ammonia, the concentrations of which can change by an order of magnitude over a 1-cm depth interval. However, these small-scale suction methods may not provide an adequate volume for conducting most standard toxicity test procedures.

Small-volume, high-resolution peepers, made by the University of Alabama at Birmingham, were designed for chemical and bacteriological analyses of interstitial water (Lalor and Pitt 1998). These peepers were made from Delrin and are about 10 to 15 cm wide and 45 to 60 cm long, with one end tapered to a point (Figures 5.80 through 5.83). The main body is made of 20-mm-thick stock and has numerous deep and wide slots (not cut through), spaced 1 cm apart, that hold about 5 to 10 mL of water each. This common peeper design enables vertical stratification of pore water quality to be determined. However, because the water volume for each separate chamber is very small, special laboratory analysis procedures are needed that minimize water volume requirements. In order to collect larger volumes of water, these peepers are frequently placed in a cluster arrangement allowing compositing from similar depth slots from adjacent peepers.

The slots should not extend any closer than about 20 mm from the edge, to prevent cracking of the thinner cover piece (common in peepers made from Plexiglas, for example). A nylon screen having 75-µm apertures is placed over this thick piece and is then covered with a thinner sheet of

Figure 5.82 Peeper placement near shore in urban Figure 5.83 Ten replicate high-resolution peepers (to lake. obtain larger water composite samples).


Delrin that is 6 mm thick. This cover piece has identically located slots cut through the material and has countersunk holes matching tapped holes in the main body. For use, the cavities in the main body are filled with distilled or deionized water, covered with the nylon screen, and the two Delrin pieces are screwed together using plastic screws, sandwiching the nylon screen (Figures 5.80 and 5.81). The unit is then pushed into the stream or lake sediment, gently pushing down on the unit until resistance prevents further penetration, leaving about five slots above the sediment/water interface (Figure 5.82). The unit is left in place until equilibrium is established, and is then removed (several hours using the large aperture screening). The unit may require up to 2 weeks for equilibrium to become established when using small aperture screenings (such as 0.45 or 2 µm membrane filter material). A recent modification has added a thin stainless steel cover to the peeper that slides over the front slots to protect them while inserting or withdrawing the peepers in sediment. The cover is slid off after the peepers are pushed into the sediment to the appropriate depth. In addition, the water is extracted from the peeper wells after disassembling the units and carefully rolling back the nylon screening, instead of puncturing the screening and inserting a syringe for sample withdrawal. These modifications have significantly reduced the disturbance to the sediments when using the peepers and have reduced contamination of the sample water.

The optimal equilibration time for in situ peepers is a function of membrane aperture, sediment type, contaminants of concern, and temperature. There are several artifact problems associated with peepers which use dialysis membranes. Total organic carbon may be elevated in peepers (4 to 8 µm pore size) due to biogenic production; however, colloidal concentrations are lower than centrifuged samples. Cellulose membranes are unsuitable because they decompose too quickly. A variety of polymer materials have been used, some of which may be inappropriate for studies of certain nonpolar compounds.

More recently, larger pore sized mesh has been used (Figures 5.84 through 5.87) which dramatically shortens equilibration time (Fisher 1992; Sarda and Burton 1995), as illustrated in Figure 5.88 during tests at UAB. In this test, 75-µm nylon screening was used on a peeper placed in a bucket of saline water (about 5.5 mS/cm). Every few minutes, the peeper was removed, and a syringe was used to remove water from an individual cell. This was then measured for conductivity.

Water Sample Extraction Tubes

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Figure 5.84 Large-volume peeper with large aper- Figure 5.85 Withdrawing interstitial water sample ture mesh. (From Burton, G. A., Jr., Ed. from large-volume peeper. Sediment Toxicity Assessment. Lewis Publishers. Boca Raton, FL. 1992b. With permission.)


Figure 5.86 Medium-volume peeper with large aper- Figure 5.87 Medium-volume peeper buried in sediture mesh for water sampling. ment.

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Figure 5.88 Equilibrium plots for 75-µm nylon screening in small-volume peeper.

Effective equilibrium was reached after about 20 min. In comparison, Figure 5.89 is an equilibrium plot for a 0.22-µm polyethersulfone membrane filter used in a diffusion peeper (Easton 2000). This test was conducted in a small laboratory flume with water flowing about 1 ft/s. Saline water was placed in the peeper (about 18 mS/cm), and the flume water was regular tap water (about 200 µS/cm). Samples were withdrawn from the peeper frequently at the beginning of the test, and at longer intervals later, and analyzed for conductivity. In this case, it required about 20 hours to reach equilibrium, although about 90% of the equilibrium was established at 10 hours.

When using sampler peepers and 75-µm membrane material, we commonly leave the peepers in place for about 2 to 24 hours to ensure equilibrium. Solids that pass through the mesh tend to settle to the bottom of the peeper chamber. Long exposure times may be impractical due to security problems and high flows in streams. The samplers need to be taken to the laboratory where the water

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Figure 5.89 Equilibrium plot for 0.22-mm polyethersulfone membrane filter in diffusion peeper (From Easton, J. The Development of Pathogen Fate and Transport Parameters for Use in Assessing Health Risks Associated with Sewage Con/tamination. Ph.D. dissertation, Department of Civil and Environmental Engineering, University of Alabama at Birmingham. 2000. With permission)

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is immediately analyzed. It is also possible to remove the samples from the slots in the field (using a syringe and needle), transferring the water into sealed and full bottles (such as small VOC vials). Four or five high-resolution peepers located close together can provide a 20 to 50 mL composite sample of pore water in 1-cm depth increments for chemical analyses (as shown on Figure 5.83).

When ionizable compounds, e.g., metals, are to be collected, it is important to preequilibrate the samplers with an inert atmosphere to avoid introducing oxygen into the sediments, thereby changing the equilibrium. Plastic samplers can contaminate anoxic sediments with diffusable oxygen and should be stored before testing in inert atmospheres (Carignan et al. 1994). In addition, when samples are collected and processed, they should also be kept under an inert atmosphere and processed quickly. Metals sampling of interstitial waters can be accomplished using a polyacrylamide gel probe (Krom et al. 1994) More recently, semipermeable membrane devices (SPMDs) filled with a nonpolar sorbant have been used effectively to show potential for bioaccumulation of nonpolar organic compounds.

Recently, test organisms have been exposed within peeper chambers where larger mesh sizes of 149 µm were used successfully in oxic sediments. Chambers can be buried several centimeters or in surficial sediment depending on the study objectives (Figures 5.90 and 5.91). Equilibration of conductivity was observed within hours of peeper insertion into the sediment (Fisher 1992). Replicate peepers revealed extreme heterogeneity in sediment interstitial water concentrations of ammonia and dissolved oxygen (Frazier et al. 1996; Sarda and Burton 1995; Sherman et al. 1994). Sediments that were high in clay and silt fractions usually were anoxic and did not allow for organism exposure in situ (Fisher 1992).

The Birmingham SSO (sanitary sewer overflow) evaluation project is a recent example of the use of peepers with large apertures. Enterococcus, Escherichia coli, total coliform bacteria, Micro-

Figure 5.90 Medium-volume peepers in situ with Figure 5.91 Surficial sediment chambers. sampling tubes exposed.


tox toxicity screening, heavy metals (copper and lead), major ions, and nutrients are being analyzed on most of the pore water samples by combining water from three adjacent 10-cm chambers, and by using five replicate peepers located close together. This allows a total of about 150 mL of water for analysis. The careful selection of test methods (and dilution of water for the bacteria analyses) allows a relatively comprehensive evaluation of pore water chemical and bacteriological conditions. Changes in pore water chemical and bacteriological quality for different depths can be used to calculate diffusion coefficients and kinetic rate coefficients.

In situ and real-time chemical measurements of interstitial water are also possible using continuously recording in situ water quality sondes. The University of Alabama at Birmingham is currently using YSI 6000 monitoring probes to continuously monitor interstitial water pH, ORP, conductivity, DO, and temperature in urban streams as part of an EPA-sponsored research project investigating SSO impacts. These instruments are capable of unattended operation for several weeks. The probe end of the instrument is wrapped with a nylon screen having 150-µm apertures. Equilibrium should be obtained within a few hours using this large aperture. The instrument can be placed vertically with the probe end buried several hundred mm in the sediment in slow-moving streams for short periods. The instrument is completely buried horizontally for longer periods or for higher flows. The use of a direct readout (hand-held readout from YSI, or a portable computer) is useful in determining equilibrium times during preliminary trials. The available turbidity probe is also used to indicate the effects of placement of the probe by measuring the exchange of water in the probe chamber. A similar unit placed simultaneously in the water column can be used to measure the lag time of any chemical changes (such as conductivity) in response to storm events and to directly determine diffusion coefficients. Of course, this method does not provide accurate vertical placement of the analytical results, but it is expected to be generally representative of nearsurface conditions where most of the benthic organism activity occurs. These probes are extremely useful to illustrate the variation of these parameters with time, especially during wet weather events, and to measure the recovery of conditions after events.

Mini-Piezometer Measurements of Pore Water Conditions

Mini-piezometers (Lee and Cherry 1978) are useful tools because they allow for the detection of upwelling groundwater and downwelling surface water on a local scale (i.e., cm to m). Additionally, these simple, inexpensive devices allow for samples of pore water to be withdrawn from desired depths within the stream bed for chemical analysis. Mini-piezometers are comprised of lengths of 1/8" ID plastic tubing that is perforated and screened with 300-µm mesh along the bottom 5 cm (Figures 5.92 through 5.94). A nest is a group of mini-piezometers of different lengths attached to a 1-m dowel rod that will sample at desired levels beneath the sediment surface (e.g., 10, 25, 50, 75, and 100 cm). Once piezometers are installed, they can be left in place indefinitely for repeated sampling and measurements. To detect areas of upwelling and downwelling, transects of nested mini-piezometers are installed in the riffle and pool areas of in situ test sites. Hydraulic heads (in cm) are determined by measuring the heights of water columns drawn simultaneously from the inserted mini-piezometer and overlying surface water into a manometer (Winter et al. 1988; Figure 5.94). Relative to surface water, a positive or negative hydraulic head indicates an upwelling or downwelling zone, respectively.

The hydrologic data from mini-piezometer pore water samples and hydraulic head measurements have improved our ability to interpret often complex exposure–effects relationships that result from in situ toxicity tests. We have found that contaminant concentrations in samples of sediments and pore water are not always predictive of in situ chamber (actual) exposure levels and observed effects in the test species. For example, in an in situ study of three sites in a stream system with similar levels of sediment contamination by chlorinated benzenes, one site was downwelling at all mini-piezometer nest locations and two sites had no net hydraulic pressure differences. Total chlorinated benzenes in water samples taken from the piezometer nests ranged


Figure 5.92 0Placement of mini-piezometers into sup- Figure 5.93 Placement of mini-piezometer array into port tubing. sediments via temporary support pipe.

from 100 to 1300 µg/L at all sites. The highest concentrations generally occurred in piezometers installed 30 cm or deeper into the stream bed. Concentrations of total chlorinated benzenes in water samples taken from the chambers used during 4-day in situ exposures of Ceriodaphnia dubia, Hyalella azteca, and Chironomus tentans to surficial sediments were near 100 µg/L at the two no-exchange sites, whereas the level was only 3 µg/L at the downwelling site. Survival of all three test species was significantly higher at the downwelling site (>80%) than at the noexchange sites (<20%). For C. dubia and H. azteca, survival between the downwelling and reference sites was not significantly different. It appears the downward flow of surface water through the sediments might have removed bioavailable contaminants in the surficial sediments to deeper zones within the stream bed (Greenberg and Burton 1999). However, this condition places transition zone species and groundwater resources at risk.

Sediment chemists, toxicologists, and risk managers have primarily focused their research efforts and the development of sediment quality guidelines on the effects of contaminants on benthic and water column organisms associated with the surficial sediments (0 to 10 cm depth). Implicit in this approach is that the historical contamination buried beneath the top sediment bed layer is biologically unavailable and hence poses little to no ecological risk. However, deeper sediments (ca. 10 to 100 cm depth), and more specifically sediments within the transition zone, serve important ecosystem functions and therefore may be sensitive to chemical perturbation. Vertical transport of dissolved or colloid-bound contaminants within the sediment interstices can potentially exert deleterious effects in the surficial sediments, surface water, or groundwater, or it can exacerbate preexisting degraded conditions. Therefore, ecosystem integrity can be more effectively evaluated if the scientific and regulatory community adopts a holistic approach to stream health that includes focusing on the transition zone. At the present time, we have begun to incorporate this added hydrologic perspective in our in situ sediment toxicity research program through the use of mini-piezometers. Continuing this line of research by developing assessment tools capable of measuring biological Figure 5.94 0Field manometer connected to mini-pie

zometer to measure ver tical floweffects within the transition zone is the next step. through sediments.


Case Example 1. Sediment Sampling for Interstitial Pore Water in an Ice-Covered Lake

A site was sampled in northern Minnesota in January which had depositional sediments (nonconsolidated silts and clays) and was ice-covered with water depths of 50 to 60 ft. Site conditions prevented use of peeper sampling and no in situ core-port sampling equipment was available. The study design required collection of 30 L of sediment. Based on these restrictions, a Ponar grab sampler was most appropriate for sediment collection.

Replicate Ponar grabs were collected through holes drilled in the ice and were deposited into a 20-L high-density polyethylene bucket and gently stirred to homogenize. Nitrogen gas was bubbled into any overlying water and added to the headspace prior to lid closure. Sediments were placed in ice chests at approximately 4°C and returned to the laboratory for processing.

Interstitial waters were collected using centrifugation. Sediments were distributed to the appropriate type of centrifuge bottles under a nitrogen atmosphere and centrifuged at 10,000 × g at 4°C for 30 min. The supernatant was gently decanted under nitrogen atmosphere. Note: if solids are resuspended with the supernatant, a second centrifugation of the interstitial water should be conducted. The interstitial water from all bottles was combined under nitrogen and then split for chemical analyses and toxicity testing. Chemical samples were preserved and stored as appropriate. Toxicity testing was initiated within 48 hours, at which time the sample temperature was raised from 4°C to the required test temperature and dissolved oxygen checked to ensure adequate levels.

Case Example 2. Shallow Stream with Contaminated Sediments

A shallow stream in Ohio with sediment contamination was studied to develop site-specific sediment quality criteria. Site conditions allowed the placement of peeper samplers. The sediment depth of concern was from 0 to 5 cm. Peepers were constructed from high-density polyethylene bottles with 70- to 140-µm PTF mesh windows on the chamber walls, 1 to 5 cm from the top of the chamber (similar to Figure 5.84). Chambers were filled with sterile deionized water and placed in a nitrogen atmosphere for 24 to 48 hours prior to site placement. Five replicate (total volume approximately 2.5 L) chambers were placed at the site by removing a plug of sediment the size of the chamber, inserting the chamber and gently packing the sediment around the chamber so that only the lid was exposed. Equilibration time can be reduced and time series sampling of the interstitial water is possible by constructing an outlet tube into the chamber lid (Sarda and Burton 1995). Degassed syringe samplers can then be attached to the outlet port and interstitial water removed without disturbing the peeper unit. Equilibration time with 140-µm mesh windows occurs within several hours. However, it may take days for the sediment gradients to reestablish adjacent to the chamber. Sampling of interstitial waters at the sediment surface (0 to 1 cm depth) is not readily feasible when large samples are required. However, toxicity may be determined on surficial sediment using in situ toxicity test chambers which expose organisms either directly to the sediments or via mesh barriers (Burton 1992a,b; see also Chapter 6). Microanalytical sampling of near-surface sediments is possible using narrow plate chamber designs (see reviews in Adams 1991, Burton 1991, and above citations). Samples are returned to the laboratory on ice and then processed by the appropriate chemical and toxicity test methods.

SUMMARY: BASIC SAMPLE COLLECTION METHODS

This chapter presented methods to determine the needed sampling effort, including the number of samples and the number of sampling locations. These procedures can be utilized for many different conditions and situations, but some prior knowledge of the conditions to be monitored is


Table 5.19 General Sampling Guidelinesa

Location

1. Locate stations at sites representative of least and greatest impact from each pollutant source and for the total system, considering each ecosystem component (e.g., substrate, flow, biota).

2. Sample depositional areas and critical habitats such as riffles and spawning areas. 3. Collect replicate samples at each station which characterize the site spatially. 4. Sample during baseflow and various stormflow conditions. 5. Sample during different seasons. 6. Sample during recovery periods (following storm events) noting different periods of disturbance (i.e., storm

recurrence period). 7. Note diurnal, weekly, monthly, and seasonal cycles of various ecosystem components-endpoints (e.g., DO,

redox, tissue residues, toxicity, life stage).

Type

8. In areas where effects are uncertain, use a “weight-of-evidence” integrated approach (see Chapter 8). Characterize the inputs and receiving water system both physically (e.g., flow, solids, temperature, habitat) and chemically (e.g., oxygen, hardness, organics, metals). Measure key indigenous biological communities (indices), indicators (e.g., trout), and endpoints (e.g., fish abnormalities). Measure toxicity of effluents, waters, and sediments using sensitive and relevant species representing multiple levels of biological organization (e.g., fish, zooplankton, algae, benthic macroinvertebrates). In situ toxicity testing is the preferred approach.

Method

9. Process samples quickly (refrigerate and/or preserve immediately upon collection). 10.Reduce sample manipulation whenever possible (e.g., mixing, sieving, aeration, filtration). 11.Maintain sample integrity when possible (e.g., using core rather than grab [dredge] collection). 12.Characterize key components of all sample replicates. 13.Follow proper QA/QC practices.

a All sampling issue decisions must be based on the study objectives and their associated data quality objectives.

needed. A phased sampling approach is therefore recommended, allowing some information to be initially collected and used to make preliminary estimates of the sampling effort. Later sampling phases are then utilized to obtain the total amount of data expected to be needed.

Descriptions of data quality objectives and associated QA/QC requirements are also given. The use of different sample blanks and other quality control samples are described, along with dealing with typical problems associated with detection limits.

The main component of this chapter covers sampling methods, including water, source area, sediment, and pore water sampling options. Numerous examples are given illustrating the use of the many sampling methods and approaches. There are few universal methods that can be used for all sampling activities, and much discretion and professional judgment is needed to select the most appropriate methods for any specific project. However, there are some general guidelines for sampling streams and lakes which should apply to most studies, as listed in Table 5.19. Each of the points listed in this table are also discussed in greater detail elsewhere in this handbook, especially in Chapter 6 and the appendices.

There are a number of factors to consider when selecting a sampling site after preliminary surveys and design elements are completed. The selection factors and their relative importance are often study specific, but some general considerations do exist, as shown in Table 5.20. The factors that influence the representativeness of a sample are numerous and cross many disciplines, as do all ecosystem evaluations. Therefore, it is important to select sampling stations based on professional judgment(s) from an individual(s) with expertise in aquatic ecosystem assessments (hydrology, environmental chemistry, biosurveys, and ecotoxicology), taking into account spatial and temporal variation and the characteristics of base- and stormwater flow; habitat; pollutant loadings, fate and effects; aquatic communities; and sensitive indicator species.

These same selection criteria should then be used to establish reference area sampling, if preexisting reference data are not available. The reference station (upstream), stream or lake, and


Table 5.20 Sampling Size Selection, Sampling Media, and Sampling Frequency Considerations

Consideration Sample Influencing Factors

Heterogeneity Ambient water, sediment effluent, runoff, biotic communities

Pollutant sources Upstream-downstream, tributary mouths, sensitive habitats, dilution gradient, beneficial uses, “typical” habitats

Beneficial uses “Beneficial” component (e.g., water supply, fishery, swimming) at critical areas

Flow, mixing, depth, particle size distribution, land use patterns, runoff coefficients, season, life-cycle, behavior, patch dynamics, pollutant partitioning (fate)

Pollutant partitioning (fate), mixing, loading characteristics, toxicity target species and endpoints, habitat complexity

Above factors

watershed should, ideally, have baseline characteristics identical to those of the test system when the pollutant problem (e.g., stormwater) being assessed is removed. However, since no two ecosystems are identical, this reference should be considered as a general benchmark from which to determine relative effect.

The next chapter presents much detail and information on evaluating samples and conditions (flow, rainfall, soil, aesthetics, habitat, water, sediment, microorganisms, benthos, zooplankton, fish, and toxicity), heavily supported with case study examples. Chapter 7 discusses statistical evaluations of the data, and Chapter 8 discusses data interpretation.

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chapter 5c sampling effort and collection methods:...

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