7. Sediment Monitoring

7.1 Overview

7.2 Collection of Sediment Samples

• 7.2.1 Field Measurements and Observations
• 7.2.2 Criteria for Selection of a Sample Collection Device
• 7.2.3 Collection Device Penetration Depth
• 7.2.4 Sample Volume
• 7.2.5 Grab Sampler Operation
• 7.2.6 Sub-sampling of Sediment Grab Samples
• 7.2.7 Criteria of Acceptability of Samples
• 7.2.8 Replicate Samples
• 7.2.9 Sediment Variables that May be Measured in the Field

7.3 Sample Handling and Analysis

• 7.3.1 Procedures for Handling of Sediment Samples
• 7.3.2 Compositing Sediment Grab Samples
• 7.3.3 Sample Containers
• 7.3.4 Transportation and Storage of Sediment Samples
• 7.3.5 Laboratory Test Sample Preparation
• 7.3.6 Prevention of Sediment Sample Contamination

7.4 Sediment Variables

• 7.4.1 Determination of Particle Size Distribution
• 7.4.2 Determination of Total Organic Carbon Content
• 7.4.3 Determination of Total Metal Concentrations

7.5 Sediment Toxicity Testing

7.6 Confounding Factors

7.7 References

7. Sediment Monitoring

7.1 Overview

As part of each benthic invertebrate community survey, mines collect sediment samples for analysis of total organic carbon content and particle size distribution, if it is possible to sample sediment (Metal Mining Effluent Regulations [MMER], Schedule 5, section [s.] 16a)(iii)). Sediment samples are collected at the same sampling stations and at the same time as benthic invertebrate samples.

More sampling stations within each area may help to better understand potential contaminant concentrations in the exposure area. Each study design for benthic invertebrate community surveys should identify the sediment sample collection and laboratory analysis methods to be used (field and laboratory methodologies selected). The results of these analyses, including calculation of the mean, median, standard deviation, standard error, minimum and maximum values for each sampling area, are included in the interpretative report. The results of analyses of particle size distribution and total organic carbon are used to determine if there are habitat differences between the exposure and reference areas, in order to aid in the interpretation of the results of benthic invertebrate community surveys. The overall purpose of sediment monitoring is to answer the question, “Are there habitat differences that may contribute to effects in the benthic invertebrate community?”

For monitoring programs where the sampling of benthic invertebrates is conducted in an erosional habitat, sediment sampling may not be possible as a standard supporting environmental variable; in these cases the sediment monitoring data would not be reported. Some methods for retrieving sediments from erosional zones require elaborate equipment or two field visits, one for the placement and one for the collection of sediment traps. However, site-specific conditions may warrant the consideration of sediment sampling in some erosional habitats, as useful information regarding exposure can be obtained with these methods. These approaches could be considered during the study design exercises for magnitude and geographic extent or investigation of cause as an additional supporting variable or tool for determining effects.

When a benthic invertebrate community survey is conducted as part of magnitude and geographic extent or investigation of cause, it is recommended that each sediment sample collected also undergo chemical analysis, (e.g., metals). The study design for that study should identify the parameters for which sediment samples will be analyzed and the laboratory methods to be used, and the results of analyses should be reported in the interpretative report.

When investigation of cause (IOC) is conducted to identify the causes of the effect on the benthic invertebrate community, detailed studies of sediment may be appropriate as a tool to help determine the cause of effect. Chapter 12 contains extensive technical guidance on the conduct of detailed sediment studies (e.g., sediment mass transport, depositional rate, coring, chemistry, sediment toxicity testing, sediment quality triad, pore water analysis, toxicity identification evaluation and toxicity reduction evaluation) recommended as part of IOC.

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7.2 Collection of Sediment Samples

This section provides guidance on the collection, handling, storage and transportation of sediment samples, and on field measurements and observations. This guidance applies to sediment samples collected in all phases of the environmental effects monitoring (EEM) program.

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7.2.1 Field Measurements and Observations

Field measurements and observations are critical to any sediment collection study. It is recommended that the following information (Mudroch and MacKnight 1991) be recorded at the time each sediment sample is collected from a sampling station:

• sample number, replicate number, station number, site identification (e.g., name)
• time and date of the collection of the sample
• ambient weather conditions, including wind speed and direction, wave action, current, tide, vessel traffic, temperature of both the air and water, thickness of ice if present
• sampling area location (e.g., positioning information) and location of any replicate samples
• type of platform/vessel used for sampling (e.g., size, power, type of engine)
• type of sediment collection device and any modifications made during sampling
• the water depth at each sampling station and the sediment sampling depth
• name of personnel collecting the samples
• details pertaining to unusual or unpredicted events that might have occurred during the operation of the sampler (e.g., possible sample contamination, equipment failure, unusual appearance of sediment integrity, control of vertical descent of the sampler)
• description of the sediment, including texture and consistency, colour, odour, presence of biota, estimate of quantity of recovered sediment by a grab sampler, or length and appearance of recovered cores (photographs provide a good permanent record of a retrieved sample)
• deviations from standard operating procedures

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7.2.2 Criteria for Selection of a Sample Collection Device

There are numerous methods and procedures reported in the literature that describe how to collect various types of sediment samples and to help determine the most appropriate sampling devices for different types of environments, including freshwater, marine or estuarine environments (for reviews see Baudo et al. 1990; Mudroch and MacKnight 1991; Environment Canada 1994; ASTM 1992; Burton 1992). Environment Canada (1994) Baudo et al. (1990) and Håkanson and Jansson (1983) suggest several factors that should be considered for the selection of sediment samplers and sampling location. The ideal sediment sampler should for the most part:

• permit free water passage during descent, to avoid a pressure wave
• have a sharp-edged cutting surface, a small-edge angle, smooth inside surface, and small wall thickness to minimize disturbance
• close tightly for the ascent
• allow sub-sampling
• have the capability of adjusting weight for penetration of different substrates
• be able to retrieve a volume of sediment large enough to meet the analytical test requirements
• effectively and consistently retrieve sediments from various water depths
• effectively and consistently retrieve sediments from the desired sampling depth
• not contaminate or influence the nature of the sediment
• require a minimum of supportive equipment
• be easy and safe to operate and not require extensive training of personnel
• be easily transported to and assembled at the sampling site

Most sediment samplers are designed to consistently isolate and retrieve a volume of sediment to a required depth below the sediment surface with minimum disruption to the integrity of the sample and no contamination of the sample. Maintaining the integrity of the collected sediments is of primary concern in most studies, since disrupting the structure of the sediment may change the physico-chemical and biological characteristics, which in turn could influence the partitioning, complexation, speciation and bioavailability of the toxicants. Sometimes it is also important to maintain the profile if sectioning is required at different depths. These issues become even more important during IOC monitoring studies when sediment may be collected for toxicity tests or more complex analytical methods. In general, it is recognized that it is difficult to collect a sediment sample with most sampling devices without some degree of disruption.

There are three main types of sediment samplers: grab, core and dredge samplers. For the first biological monitoring studies and studies conducted to confirm presence or absence of effects, grab samplers are recommended. Grab samplers are used to collect surficial sediments for the determination and assessment of the horizontal distribution of sediment characteristics. Details on this topic can be found in de Groot and Zschuppe (1981), Baudo et al. (1990), ASTM (1992), Burton (1992) and Sly and Christie (1992).

Core samplers collect a column of sediment to examine the historical or vertical distribution of the physical and chemical characteristics of the sediment (Environment Canada 1994). Corers are preferred in cases where the integrity of the profile is essential, as they are the least disruptive. For these reasons, corers should be considered for magnitude and geographical extent and IOC studies. For additional information on core samplers, refer to Environment Canada (1994) and Chapter 12.

Dredges are used primarily for the collection of benthos, since they are usually equipped with net sides designed to filter out fine-grained sediments and retain coarse sediments and fauna. It is virtually impossible to accurately measure the surface area covered by the dredge sampler, or judge the depth to which the sediment sample has been collected. In addition, sediment integrity is disrupted, pore water excluded, and fine-grained sediments lost during ascent using dredge samplers. For these reasons, only grab (first biological monitoring studies and studies aim to confirm presence or absence of effects) and core samplers (magnitude and geographic extent and IOC) are being recommended for the collection of sediments.

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7.2.3 Collection Device Penetration Depth

The desired depth of sediment penetration is a decision that depends upon the type of sampling device, the nature of the sediment, and the volume of sediment required. The actual depth of penetration depends primarily on the type of sampling device and the nature of the sediment.Generally, the most recently introduced contaminants of concern and most infaunal organisms are found in the upper 2 cm. Epifaunal organisms also have access to this horizon (Burton 1992). Therefore, a preferred penetration depth of 10-15 cm and a minimum penetration depth of 6-8 cm are recommended to ensure minimum disturbance of the upper layer during sampling. This depth is also appropriate for monitoring studies where historical contamination is not a priority (upper 0-5 cm of sediment).

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7.2.4 Sample Volume

The recommended minimum volume or weight of sediment needed for each end use should be determined on a case-by-case basis and is available in Table 3 of Environment Canada (1994). Before commencing a sampling program, the type and number of analyses and tests should be determined, and the required volume or weight of sediment per sample calculated. Each physico-chemical test requires a specific amount of sediment. After the sample size is determined, it is important to compare the sample size required with the capacity of the sampler to deliver the desired amount of sediment, and reassess the number of replicate samples per station. The volume or weight requirements might dictate further sample handling such as sub-sampling, compositing, or sample splitting.

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7.2.5 Grab Sampler Operation

When collecting bottom sediments with grab samplers, the speed of descent of the sampling device should be controlled and the sampler should not be permitted to “free fall.” To minimize twisting during the descent, a ball bearing swivel should be used to attach the sampler to the cable. The sampler should contact the substrate or be positioned just above it and only its weight or piston mechanism should be used to force it into the sediment. The winching system should be in place to control both the ascent and descent of the sampling device, especially in deep water. After the sample is contained, the sampling device should be lifted slowly off the bottom, then steadily raised to the surface at about 30 cm/s. When the sampler is brought to the surface, the outside of the sampler should be carefully rinsed with water from the sampling station to remove material that could potentially contaminate the sample during transfer. The sampler should be inspected to ensure that it has closed properly. The standard operating procedures specific for each grab sampler should be followed in order to ensure proper operation of the sampler.

Regardless of the type of samplers used, standard operating procedures for each device should be immediately accessible, and all personnel involved with the collection of samples should be familiar with these procedures. The sampling vessel or platform should be stationary, and sufficiently stable to permit inspection and handling of the retrieved sample. Field notes should accompany each sample that is collected. The sampling device should be cleaned thoroughly between sampling stations and between within-station samples by dipping the sampler into and out of the water at a rapid speed to wash the sediment off. Alternatively, a hose can be used to wash the sediment off of the sampler with water from the sampling station. The sampler should be rinsed with water from the next sampling station before collecting a sample.

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7.2.6 Sub-sampling of Sediment Grab Samples

If sediment grab samples are to be sub-sampled, access to the surface of the sample without a loss of water or fine-grained sediment is a prerequisite for selection of the sampler.

The non-turbid overlying water, if present, should be gently siphoned off before the sediment is sub-sampled, using a flat, clean scoop (e.g. Teflon® or a similarly inert, non-contaminating, non-reactive material) or a suitable hand-coring device. Ideally, each sub-sample should be placed into a clean, separate, pre-labelled container. The labelled sample container should be sealed and the air excluded.

In the event that the collection device does not allow access to the surface, the following procedures should be followed. Upon retrieval of the sample, the contents should be carefully deposited into a clean, inert container that is the same shape as the sampler. The sampler is placed into the container and the jaws opened slowly to allow the sample to be deposited into the container with as little disturbance as possible. Once the sample is in the container, sub-samples can be collected from the sample with a hand corer or scoop. The edges of the sample where the sediments may be disturbed during removal from the sampler should be excluded during sub-sampling.

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7.2.7 Criteria of Acceptability of Samples

All samples should be visually inspected to ensure that:

• the desired depth of penetration has been achieved
• there is no evidence of incomplete closure of the grab sampler, or that the grab sampler was inserted on an angle or tilted upon retrieval (i.e., loss of sediment)

If the collected sample fails any of the criteria listed above, then the sample should be rejected and another sample collected at the site. The location of consecutive attempts should be as close to the original attempt as possible while avoiding any overlap and, where the direction of the current is known, consecutive attempts should be located in the opposite direction of the current, or “upstream.” Rejected sediment samples should be discarded in a manner that will not affect subsequent samples at that station or other possible sampling locations.

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7.2.8 Replicate Samples

A single sediment sample from a sampling station will impart little information on the variability. Environment Canada (1994) therefore recommends the following for the minimum number of replicate samples:

• When replicate samples from a sampling station are recommended, the collection of a minimum number of five replicate samples within a sampling station is recommended unless determined otherwise from preliminary sampling and analysis.
• The collection of replicate samples is needed as part of the QA/QC of any good sampling program and should comply with the data quality objectives.
• The number of replicate samples should be higher at stations located close to a source of contamination (Skei 1992).

Collecting separate replicate samples at each sampling station allows for quantitative statistical comparison within and among different stations (Holland et al. 1993). The collection of separate samples within a sampling area can impart valuable information on the heterogeneity of the sediments. Separate sub-samples from the same grab can be used to measure the variation within a sample but not necessarily within the sampling station.

The number of replicates needed per sampling station is a function of the need for sensitivity or statistical power. Typically, the smallest deviation from the null hypotheses that is considered scientifically or environmentally important to detect should be decided a priori, together with the power of the test that is desired for the specific alternative (Green 1989).

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7.2.9 Sediment Variables that May be Measured in the Field

In addition to the required determination of total organic carbon and particle size distribution, it is recommended that the following sediment variables be measured in the field, particularly during magnitude and geographic extent and investigation of cause:

• temperature and pH of the sediment at the sediment-water interface
• a measure of the redox potential of the sediments to determine if the sediments are oxic or anoxic, or to determine the depth of the interface between these conditions in the sediments. Dissolved oxygen is recommended for freshwater sediment and redox potential (Eh) is recommended for marine sediments.

These measurements could be useful for the interpretation of the analytical results.

7.3 Sample Handling and Analysis

7.3.1 Procedures for Handling of Sediment Samples

Any time that sediment samples are handled, it is recommended that the following procedures be observed:

• Sediment might contain a mixture of hazardous substances, so it is prudent to avoid skin contact with sediments by wearing protective clothing and equipment (e.g., gloves, boots, lab coats or aprons, safety glasses, and respirator) during sampling, sample handling, and the preparing of test substances.
  
• Handling of samples should be performed in a well-ventilated area (e.g., outside, in a fume hood, or in an enclosed glove box) to minimize the inhalation of sediment gases.
  
• Work surfaces should be covered with Teflon^® sheets, high-density polyethylene trays, or other impervious or disposable, similarly inert material.
  
• A spill control protocol should be in place in the laboratory or sampling vessel, and participants in the project should be familiar with all standard operating procedures and recommendations.

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7.3.2 Compositing Sediment Grab Samples

If the objective of the study dictates compositing sub-samples from separate grabs within a sampling station, the sub-samples may be placed into one clean sample container and, when full, sealed without trapped air. Compositing of sediment samples or sub-samples may also be performed in the laboratory.

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7.3.3 Sample Containers

Environment Canada (1994) provides information concerning the storage and transportation of field-collected sediment samples.

Whole-sediment samples may be transferred directly from a sampler into a clean, large-volume (e.g., > 1 L) container. If smaller volumes of sediment are collected or sub-sampled, containers with wide mouths and Teflon^®-lined lids are recommended for volumes ranging from 250 to 1000 ml.

If samples are to be stored at 4°C, sample containers should be filled to the rim and air excluded during capping. If samples are to be frozen for storage, glass containers should not be filled completely. A space of approximately 2.5 cm should be left to accommodate expansion of the sample when frozen; however this will depend on the size of the container and the percent moisture of the sample. The headspace in the container should be purged with nitrogen before capping tightly. Clear glass containers may be wrapped with an opaque material (e.g., clean aluminium foil) to eliminate light and reduce accidental breakage.

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7.3.4 Transportation and Storage of Sediment Samples

The recommended procedures and conditions for the transportation and storage of sediment samples are as follows (Environment Canada 1994):

• The transport container should be refrigerated to 4 ± 2°C or contain ice or frozen gel packs that will keep the field samples below 7°C during transport to the laboratory.
  
• If field-collected samples are warm (e.g., > 6°C), they should be cooled to between 1 and 6°C with ice prior to placement in the transport container.
  
• Samples should not freeze during transport.
  
• Ideally, a maximum/minimum thermometer or a continuous temperature recorder should be placed inside the transport container and the container sealed. Deviations in temperature should be reported.
  
• Light should be excluded from the transport container.
  
• All field-collected samples that require further processing before storage should be transported to the laboratory within 72 h, preferably within 24 h, of collection.

Where these conditions cannot be met due to operational constraints, the storage method and conditions adopted should strive to compromise the integrity of the sample as little as possible (Mudroch and MacKnight 1991).

Each sample container should be properly labelled and stabilized in an upright position in the transport container. Labelling of each sample container should include, at a minimum, the site, station location or identification, the sample type, the method of collection, the name of the collector, and the date and time of collection.

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7.3.5 Laboratory Test Sample Preparation

Sediment samples should be prepared in a well-ventilated area (e.g., fume hood) and the appropriate health and safety precautions should be followed. For first and second biological monitoring and magnitude and geographic extent, the preparation of samples under anoxic conditions is not a concern. However, for investigation of cause techniques such as toxicity testing, preparation of anoxic test sediments should be performed in a glove box in the presence of a controlled flow of an inert gas, if it is desirable to maintain these anoxic conditions. Below are some details on sediment preparation techniques used to allocate sediment to test containers:

Homogenizing: Mixing by hand or mechanical means may be used to achieve homogeneity of colour, texture and moisture; however, the efficacy of the method should be demonstrated, a priori, and the mixing time standardized to ensure consistency and minimize alterations in the size distribution of sediment particles.

Mixing of sediments should take place in the sample/storage container.

Partitioning: Coning or caking and quartering are the recommended techniques for partitioning the sediment for distribution among test containers. If a sediment splitter is used, its efficacy should be demonstrated and documented and it should be made of an appropriately inert material.

Drying: The recommended methods for drying sediment are oven-drying sediment sub-samples (1-5 g of wet sediment) at low temperatures (40-60°C) until a constant weight is reached or freeze-drying sediment subsamples.

Crushing/Grinding: Commercially available ball and pebble mills are recommended for fine-grinding small volumes of sediment (Mudroch and MacKnight 1991); however, it should be noted that grinding could change the chemistry of the material. Crushing can usually be achieved with a mortar and pestle.

Dewatering: Centrifugation with subsequent decanting of the supernatant is the recommended method for dewatering sediment samples. The centrifugation speed depends on the sample size and particle size (e.g., sediment weight or volume).

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7.3.6 Prevention of Sediment Sample Contamination

When sediment samples are to be collected for chemical analysis, the procedures for the collection, handling, transportation and storage of samples are much the same as those outlined above. However, in such cases it is important that appropriate measures be taken to ensure that sediment samples are not contaminated.

When sediment samples are to be collected for chemical analysis, sample collection devices should not be made of copper, zinc, brass or galvanized material, since collection devices made of unprotected metallic material can potentially affect the concentrations of metals in sediment samples. If this is not possible, when sub-sampling, the sediment that is in direct contact with the sides of the sampler should be excluded. The sub-sampled sediment should be transferred to clean containers made of inert material that will neither contaminate nor influence the characteristics of the sediment sample. The container should be tightly sealed and air should be excluded.

All sample containers should be pre-treated prior to receiving a field sample (Environment Canada 1983, 1989). New glass and most plastics should be pre-treated to remove residues, and/or leachable compounds, and to minimize potential sites of adsorption. Pre-treatment includes the following sequence of activities (adapted from Environment Canada [1989]):

• Scrub with phosphate-free detergent and hot water.
• Rinse with high-pressure hot water.
• Subject to a 72-h acid bath with 8 M HNO₃ (50 ml of HNO₃ per litre of water).
• Rinse four (4) times with hot water.
• Rinse three (3) times with DDW (double distilled water).
• Wash bottle caps (Teflon^® or Teflon^®-lined) with detergent and hot water, and rinse with DDW.

The acid bath will leach trace metals (e.g., Cu, Fe, Mo, Ni, Zn) from plastics. The triple rinse with distilled water is necessary because the acid treatment can activate adsorption sites on polymers which are then capable of binding trace metals in the field sample.

7.4 Sediment Variables

7.4.1 Determination of Particle Size Distribution

Where it is possible, the determination of sediment particle size distribution should be conducted each time that a benthic invertebrate community survey is conducted. Particle size should be determined for a minimum of one sample from each benthic sampling station.

Particle size determination is important in the interpretation of the results of chemical or biological analyses. Most importantly, from the point of view of using this data to aid in the interpretation of the results of benthic invertebrate community surveys, particle size has a significant impact on the structure of benthic invertebrate communities. It may also provide insight into the origin of sedimentary materials and about the dynamic conditions of sediment transport and deposition. From particle size analysis, specific surface, expressed as m²/g, can be determined, and with this, the adsorptive capacity of metals and organic substances can be assessed.

Many different classifications of particle sizes exist; however, the following breakdown based on the Wentworth (1922) classification is recommended for the interpretation of EEM data.

Procedures for methods of sediment particle size analysis can be found in ASTM (2003). Particle size analysis or grain size analysis is generally performed in two parts: sieve analysis and hydrometer analysis. The sieve analysis classifies particles greater than 0.06–0.075 mm in size (actual minimum size depends on the sieve set used). This is done by wet-sieving the sample through a set of at least four sieves, ranging in size from 0.06 mm to 16 mm. The material retained on the sieves is dried and weighed. Particles passing through the 0.06-mm sieve are collected and transferred to a 2-L container, together with the wash water. A hydrometer is used to determine the quantity of particles in this fraction from 0.06 mm down to 0.0014 mm. The data from these two tests are then tabulated and calculated to produce a particle size distribution curve. This curve graphically defines the percentage of material in the different fractions based on the total sample weight.

It is also possible to determine particle size distribution using laser diffraction, and this method is increasingly available. This method is more efficient and provides higher resolution results than the above methods. A laser diffraction instrument uses light from a low-power helium-neon laser (the analyzer beam). Particles from sediment samples enter the beam via a dispersion tank that pumps the material, carried in water, through a sample cell. The light scattered by the particles is incident onto the receiver lens, which focuses the scattered light onto a diode composed of numerous concentric rings. Through a process of constrained least squares fitting of theoretical scattering predictions to the observed data, the computer calculates a volume size distribution that would give rise to the observed scattering characteristics. No a priori information about the form of the size distribution is assumed, allowing for the characterization of multi-modal distributions.

The efficiency of laser diffraction is also a major benefit. A typical measurement takes only a few seconds, and the data are saved digitally and are instantly available for plotting and other calculations. Often, the entire distribution can be accounted for in a single measurement. Depending on the instrument used, a laser particle size analyzer can measure all sizes ranging from 0.05µm to 2000µm. For samples with a size range greater than 2000µm, sieve data can be merged with the laser results. Finally, the results using laser diffraction are very high resolution, and are easily reproducible--overcoming a major shortcoming of the hydrometer and sieve methods.

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7.4.2 Determination of Total Organic Carbon Content

As with particle size distribution, the amount of sediment total organic carbon (TOC) should be determined each time that a benthic invertebrate community survey is conducted. TOC should be determined for a minimum of one sample from each benthic sampling station.

Carbon is present in sediment in several organic forms such as humic matter; chemical, plant and animal matter; as well as inorganic carbonate forms. Organic carbon in sediment and the water column causes a decrease in dissolved oxygen by using up available oxygen, hence creating a more anoxic environment. Also, at certain pH levels, humic substances form complexes with metals, increasing metal solubility in the water column. Two methods are commonly used to analyze TOC in sediment. The elemental analyzer method, valid for samples of 0.5–25 mg, is based on the use of thermal conductivity. The oxidizing furnace method requires samples of 0.25–0.5 g and is based on the use of infrared spectrophotometry.

Elemental analyzer:Inorganic carbon is first eliminated by treatment with hydrochloric acid. TOC is then oxidized to carbon dioxide in the presence of a catalyst. The gas produced is separated by chromatography and quantified with a thermal conductivity detector.

Oxidizing furnace:Inorganic carbon is first eliminated by treatment with hydrochloric acid. TOC is then oxidized in the oxidizing furnace in the presence of manganese dioxide. The carbon dioxide formed from the organic carbon is measured directly by infrared absorption at the characteristic wavelength for carbon dioxide.

Procedures for these methods of analyzing TOC in sediment are described in US EPA (1986) and APHA (1995).

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7.4.3 Determination of Total Metal Concentrations

The determination of total metal concentrations in sediments is not required as part of the EEM program. However, mines are encouraged to determine total metal concentrations in sediments when benthic invertebrate community surveys are conducted. Information regarding metals in sediment can be important to the interpretation of the results of benthic invertebrate community surveys, and to the design of subsequent surveys. If effects at a site are suggestive of eutrophication, consideration should also be given to determination of sediment nutrient concentrations.

Sediments are an integral component of aquatic ecosystems, and hence, a frequent aspect of many environmental monitoring programs. They originate from the differential settling of both suspended terrigenous particles that have been introduced into aquatic ecosystems and precipitates that have resulted from chemical and biological processes within aquatic systems. Suspended particles entering the aquatic system may already contain contaminants. Alternately, non-contaminated particles suspended in water may accumulate soluble contaminants present in the waters of aquatic systems. Precipitation processes are also capable of scavenging contaminants. As a result, sediments can be viewed as either a reservoir or a sink for contaminants.

Contaminated sediments from point-source inputs such as mining effluents can become bioavailable and enter aquatic food-webs, therefore affecting the quality of the habitat. Measuring sediment quality helps identify which contaminants are entering the exposure area. Sediments provide a better integrator of average long-term environmental conditions than single-event water chemistry samples.

The selection of parameters for sediment chemistry analysis will be determined on a site-specific basis. Where historical data exist on sediment quality, they should be used in conjunction with effluent characterization and water quality data to help determine parameters to analyze.

Total, or bulk, sediment chemistry provides information on the loading rates of particular elements, and on depositional patterns. Techniques used for determination of metals in sediments include atomic absorption spectrophotometry (AAS), X-ray fluorescence (XRF), instrumental neutron activation analysis (INAA), inductively coupled atomic absorption spectrophotometry (ICP-AES) and ICP-mass spectrometry (ICP-MS). Because of the high concentrations of metals in sediment, particularly in mining areas, analytical techniques with higher detection limits (e.g., ICP, ICP-AES) are generally acceptable for sediment chemistry analysis.

Bulk sediment samples are digested using either aqua regia or a mixture of perchloric, nitric and hydrochloric acids for extraction of total metals. Metal concentrations can be influenced by sediment particle size and organic carbon content. Smaller particles and organic material have a higher affinity and more binding sites for metals than coarser grained material. Therefore, with all other factors being equal, total metal concentrations tend to be higher in fine organic substrates. To account for this influence, it is recommended that sediment samples be sieved through a 63-mm mesh screen, and that only the fraction less than 63 mm be analyzed.

Alternatively, metal concentrations in sediments can be normalized for particle size or organic content when comparing results between areas. Sediment metal data can be normalized to percent fines (silt + clay fractions) using the following equation (ESP 1996):

7.5 Sediment Toxicity Testing

Sediment toxicity testing is not a required element of EEM. There are many uses of sediment toxicity testing, including evaluating potential contamination in aquatic environments, verifying alterations seen in benthic invertebrate communities that may be due to toxicity of sediments and not other physical or biological factors, and possibly to interpret confounding factors (see below). Chapter 12 contains extensive guidance on sediment toxicity testing.

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7.6 Confounding Factors

Sediment toxicity testing can help interpret situations where field effects are inconclusive due to confounding factors such as historical contamination or multiple dischargers to the same watercourse. Whole-sediment toxicity tests, conducted in the lab, typically use a standard overlying water, thereby isolating the effects of the sediment. Research underway by Environment Canada (Lisa Taylor, personal communication, Ecotoxicology and Wildlife Health Division, Environment Canada) is looking at the modifying effects of water chemistry parameters on sediment toxicity and whether it can be accounted for by using water collected from the study site as the overlying water. Site waters could include upstream receiving water, downstream receiving water (i.e., effluent mixed with receiving water), full-strength effluent, and/or a “clean” reference site water. The choice of overlying water should be decided based on the objectives of the study. Examples of study objectives include isolating effects due to historically contaminated sediments from those due to current effluent, determining whether the current effluent is modifying the bioavailability of contaminants within the sediment, or verifying whether the water upstream of a discharge point influences the toxicity of sediment collected below the discharge point. If the site water is confounded due to multiple dischargers to the same watercourse, it may also be more useful to use a standard overlying water that is simulated in the laboratory to match the field conditions. Parameters likely to ameliorate toxicity, such as pH, hardness, alkalinity or dissolved organic matter, can be adjusted in laboratory water to approximate the situation in the field.

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7.7 References

[APHA] American Public Health Association. 1995. Standard methods for the examination of water and wastewater. 19th edition. Washington (DC): American Public Health Association.

[ASTM] American Society for Testing and Materials. 1992. E 1391-90, Standard guide for collection, storage, characterization and manipulation of sediments for toxicological testing. In 1992 Annual book of ASTM Standards, Vol. II.04, Section 11. Philadelphia (PA): American Society for Testing and Materials. p. 1134-1153.

[ASTM] American Society for Testing and Materials. 2003. D422-63, Standard test method for particle-size analysis of soils. In Annual book of ASTM Standards, Vol. 04.08. West Conshohocken (PA): American Society for Testing and Materials. p. 10-17.

Baudo R, Giesy JP, Muntau H, editors. 1990. Sediments: chemistry and toxicity of in-place pollutants. Chelsea (MI): Lewis Publishers, Inc.

Burton GA Jr, editor. 1992. Sediment toxicity assessment. Chelsea (MI): Lewis Publishers Inc.

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