Field Procedures for Benthic Invertebrates
1. Samplers
Marine sediment sampling protocols are specified in Washington State Department of Ecology’s Environmental Assessment Program (EAP) SOP for Marine Sediment Sample Collection, which generally follow PSEP (1987). An updated methodology used by EAP Washington State is included in another hyperlinked document in this collection (see parent document). A good review of marine sampling methods is available in Eleftheriou and Holme (1984). Depending on the scale of the aggregation in the benthos, the overall mean abundance per grab should remain the same regardless of sampler size. However, variance will change with the size of the sampler. Therefore, it is recommended that as a standard, well-tested 0.1m2 grab be used. Examples include the Smith-McIntyre grab, Shipek or a modified Van Veen grab. The one critical aspect of these samplers is that they have a screened top to allow smooth progress downward (water flow-through) with a minimum bow-wave.
In very shallow and/or sandy habitats it may be necessary to use a smaller sampler deployable from small inflatable boats and retrievable by hand. Although this is not ideal because of the inevitable loss of larger organisms, it may be necessary for logistical reasons.
Intertidal soft substrates may be sampled using any device which demarcates at least 0.1m2 area. The soft substrate is then removed to a standard depth of 10 cm using whatever device is appropriate. Note that in general, the lowest intertidal level available for sampling is preferred because less harsh physical conditions promote higher species richness and abundance.
Table 1 Summary criteria for recommended samplers
| Sampler Characteristics | ||
| Marine/ Estuarine – Depositional Habitats – | ||
| Petite Ponar Grab AREA : 0.02 m2 | · Habitats and substrates sampled: freshwater lakes, rivers, and reservoirs and estuaries with moderately hard sediments such as sand, silt, and mud; will not penetrate clay; somewhat less efficient in soft sediments and coarse gravel.· Effectiveness: limited depth penetration; not useful in clay because of light weight, half recommended sample area size requires twice as many grabs.· Advantages: good penetration for such a small grab; can be operated by hand.· Limitations: jaws can be blocked by stones, sticks and other debris causing loss of part of the sample; not efficient in swiftly flowing water; smaller size makes it inefficient for sampling larger benthic organisms; not entirely adequate for deep burrowing organism in soft sediments | |
| Standard Ponar Grab AREA : 0.05 m2 | · Habitats and substrates sampled: freshwater lakes, rivers, estuaries, and reservoirs with hard and soft sediments such as clay, hard pan, sand, gravel, and muck; somewhat less efficient in softer sediments.· Effectiveness: not entirely adequate for deep burrowing organisms in soft sediments; very efficient for coarse sediments; collects both qualitative and quantitative samples· Advantages: better penetration than other grabs; side plates and top screens reduce washout, shock waves and substrate disturbance; best quantitative grab sampler for freshwater use.· Limitations: a heavy grab that requires use of a boat with winch and cable; stones, pebbles, and other debris can hold jaws open causing loss of sample. | |
| Smith-McIntyre Grab AREA : 0.1 m2 | Habitats and substrates sampled: marine and estuaries; adaptable to large rivers, lakes, and reservoirs with sand, gravel, clay, and similar substrates.Effectiveness: has been widely used for sampling in marine and estuarine habitats, penetration – 17 mm in silt and clay, 5 dm in coarse sand..Advantages: provides reasonably quantitative samples; trigger plates help penetrate the substrate, top screens and stabilizing frame reduces washout and ensures upright landing.Limitations: very heavy, needs boat and power winch; spring-loaded jaws could be hazardous; inefficient for collecting deep burrowing organisms; jaws can be blocked by debris. | |
| Van Veen Grab AREA : 0.1 or 0.2 m2 | · Habitats and substrates sampled: marine and estuaries with sand, gravel, mud, clay, and similar substrates; could be adapted to freshwater.· Effectiveness: penetrates to a depth of 5 to 7 cm.· Advantages: jaws close better than the Petersen grab; samples most types of sediments; comes in a range of sizes.· Limitations: a very heavy grab that requires a large boat and power winch; jaws may become blocked by debris such as rocks and sticks; not useful for deep burrowing organisms. | |
| Quadrat to depth of 10 cm AREA : 0.1 m2 | · Habitats and substrates sampled: intertidal marine/estuarine soft substrates to fine gravel.· Effectiveness: very effective with care of field personnel.· Advantages: inexpensive, allows sampling of substrates not accessible with grab samples · Limitations: may not sample deep burrowing organisms in sandy sediments. | |
Source: Eleftherious and Holme (1984), Klemm et al. (1990), Scrimgeour et al. (1993)
The grab will be attached to the vessel’s cable and winch system and lowered to 2-3 meter above the sediment surface. The vessel will be manoeuvred into position above the target location. The grab will then be lowered to the bottom where it will trigger and close upon contact with the sediment surface, and a sample will be collected. The grab will then be raised back up to the vessel and landed on a grab stand.
Sediment samples may also be collected using a double 0.1-m2 stainless-steel modified van Veen grab sampler, which allows sampling of sediment conventionals simultaneously with benthic infaunal samples. If a double van Veen grab is being used, one side will be used for determination of various physical/environmental characteristics, including sample penetration depth, sediment temperature, salinity of the overlying water, and sediment texture, color, and odor. The other side will be used for benthos collection.
2. Criteria for acceptability of grab samples
The collected sediment sample will be visually inspected. Any grab sample lacking fine-grained particles in the sediment (i.e., composed of all cobble, shell hash, or wood, etc.) or for which the jaws of the grab do not close completely will be rejected. Any grab sample that is rejected for any reason should be dumped overboard after the vessel has been repositioned away from the target location. Ideally, a good sample, in order of priority, is:
- Completely contained in the grab sampler (i.e., minimal sample has been lost through the bottom or top of the grab);
- Is representative of the square area sampled, with the two sides of the sample being similar in volume and thus evenly sampled (i.e., landed upright on the bottom);
- Is >70% full and is equivalent/near equivalent in volume to other replicates of the same station, and other samples in the same. An alternative approach for a marine 0.1 m2 grab sample is a criterion penetration of >/= 10 cm depth and > 4 litres of sediment would be considered an acceptable sample (Gray et al. 1992).
Samples not meeting all of the above criteria may be acceptable in circumstances where substrate penetration is difficult. The maximum number of attempts for any one location should be pre-determined. If this number is reached, the station/sample may be abandoned.
Changing the weighting of the grab is the best tool for trouble-shooting penetration issues. The addition of more weight may help with penetration to a certain point, but if the substrate is exceptionally difficult (hard packed sand, lots of rock), it is unlikely the addition of weight will make a difference.
Conversely, if the substrate is soft, it is possible for the grab to be over-weighted. In this circumstance, the soft sediment may overflow through the top flaps of the grab. This is undesirable as it a sample loss. In this circumstance, weight should be removed from the grab. Also, the height off the bottom from which the grab is allowed to free-fall may be adjusted depending on the substrate type.
Once a sample is deemed acceptable, a picture of the sample (in the grab with the flaps lifted) is taken, and then the sample placed in a tote for screening. It is important that the sample be rinsed out in its entirety into the tote. This sample must be labeled, at minimum, with a label inside in the sample tote, in the sample itself. It is also useful to label the outside of the tote.
3. Replication
The allocation and distribution of replicate stations is dependent on the site-specific conditions, questions being addressed and the sampling pattern used. For a complete example of a power analysis with invertebrate community data please refer to Lowell (1997). An example of determining critical effect size from the variability inherent in reference areas is presented therein.
A simpler way to examine adequacy of replication is to assess the abundance and degree of aggregation of organisms in relation to the desired level of precision for replicate station estimates. For a given replicate station, the number of field sub-samples needs to be sufficient to give a mean and variance which provides confidence that a representative number of animals has been captured (for review see Burd et al. 1990). The more aggregated a community, the higher the variance of mean abundance for each replicate station. Elliott (1977) and Holme and McIntyre (1984) suggested the same simple method of determining the number of field sub-samples required to obtain a predefined level of precision. Elliott (1977) suggests that toleration of an index of precision (D) of 20% (i.e., that the standard error is equal to 20% of the mean) is acceptable for most bottom samples. The number of field sub-samples can then be calculated as follows:
n = s2 / D2 X
where,
X = the sample mean
n = the number of field sub-samples
s2 = the sample variance
D = the index of precision (i.e., 0.20)
Based on extensive experience with this process in the Pacific Northwest, standard procedure for subtidal coastal marine areas for which biota distributions are not well known is to collect 5 replicate samples at each location. A minimum of 3 replicates should be collected and processed in the laboratory to determine sampling precision (SE/Mn should be </= 20%). If this is adequate, the remaining 2 replicates could be archived unprocessed, for possible use in future studies or analyses. In summary, a minimum of 0.3m2 of grab sample surface area should be processed in its entirety for benthos.
Where smaller samplers are being used (see section 1 above), a minimum number of replicates should be processed in the laboratory to make up to 0.3m2, with collection of at least 2 extra replicates for safety. Pooling of replicates is not recommended, since it could later be important to determine intra- and inter-sampler variability in different habitat types.
4. Equipment
The optimum field screening system consists of an aluminum stand with stacked trays and a seawater pump (battery operated when used on board a ship) with an intake and outflow hose. The washing stand should include a chute for directing the wash-water screened mud/silt off the deck of the boat (or side of the dock). This stand should be secured to the side of the boat or the edge of the dock during screening, with the chute directed overboard/over the edge of the dock.
Generally one 1.0mm screen is placed on the top of the screen and is the only screen used. If additional fractions are being collected, is it is important to place the plastic flaps on the screen to direct the sample into the tray underneath.
Seawater pumps have water flow from 1.5-2 gallons per minute (gpm) (old pumps) or 3-4 gph (new pumps). Flow on the new pumps should be adjusted with the ball valve to ~2 gallons per minute so it is a gentle flow and does not cause damage to the organisms. Spray nozzles should not be used for this reason. The intake hose has an attachment on the end with a 250 mm screen that prevents planktonic organisms or other debris from entering the sample. This “hose-end” should also have a check valve that prevents backflow of water.
5. Field Processing of samples
Extreme care should be taken during washing of samples to avoid breakage of specimens, which can greatly reduce taxonomic efficiency and cost-effectiveness. Methods have been described to reduce breakage, particularly in marine samples (Gray et al. 1992), and are summarized below.
Screening should ideally take place immediately after collection, as specimens can start to die and decompose within ~6 hours after collection, depending on the air temperature. If samples are collected on a warm day, it is imperative these are screened and preserved as quickly as possible (screening should commence within ~1 hour). However, in colder weather (water temperature or below), the samples may be screened within 4 hours of collection without detriment. However, it is best not to have the samples sitting after collection in either case to minimize potential damage to the sample integrity.
Field sieving should also be done before preservation as many organisms become fragile and brittle after preservation and handling damage should be minimized. Furthermore, storing large volumes of fine sediment with preservative is less efficient than sieving and then preserving. Not only do organisms at the bottom and in the middle of a densely packed sediment not get sufficiently preserved, considerably more toxic preservative (i.e., formalin) is needed. Formalin must also be disposed of in the appropriate manner, often at additional cost, and in the interests of environmental conservation, the quantity of formalin used should be minimized wherever possible.
For marine organisms, samples should be sieved with seawater rather than freshwater, since the osmotic shock of freshwater may cause cell bursting and gross distortion of the animals and where appropriate, field water used to sieve should be screened for ambient organisms with a mesh smaller than the required minimum screen size used for the study.
6. Seive sizes
In marine systems, macrobenthos are typically those retained by sieves with 500-1000 µm mesh (Reish 1959, Thiel 1975, Pearson 1975, Holme and McIntyre 1984, Gray et al. 1992). It is estimated that a 1.0 mm sieve will retain about 95% of the biomass of marine macrofauna (Reish 1959) while reducing the numbers of juvenile taxa and meiofauna present in samples which respond functionally differently to environmental perturbation than adult macrofauna (Schwinghamer 1981, 1983, Warwick 1986).
For SSAMEx it is recommended that a maximum mesh size of 1mm be used for processing of benthic macro- infauna. If a smaller screen is required for site-specific purposes, the grab samples should be washed in the field through the minimum screen size, then rewashed prior to processing in the lab through a stacked set of sieves to a maximum size of 1 mm. The results for the 1mm screen and any smaller mesh sizes should be reported separately to allow inter-comparison of the standard 1 mm size assemblage throughout the Salish Sea.
7. Washing the Sample
Samples are washed in portions to minimize the opportunity for animals to become fragmented on the screen. The washer holds the tote containing the sample on the edge of the washing stand, and puts the hose into the sample itself, breaking up the sample inside the tote. Thus most of the washing takes place in the tote itself, with the overflow containing debris that is retained on the screen.
The unwashed sample should NOT be placed directly on the screen. If a large volume of unscreened debris on the screen requires washing, this not only creates damage to organisms, but also is often much slower, and results in samples that are not as well-screened.
Large, heavy debris such as rocks can be removed and preserved separately (in a bucket, jar or bag) to prevent damage to organisms. It is recommended to preserve all of the rocks in the sample, as these may have epifaunal organisms. However, if a sample contains a large volume of smooth, clean, rocks, and each rock is checked carefully for epifauna, these may be discarded.
Any visible fragile organisms (e.g., brittle stars, nemertean worms, some tubeworms, scaleworms) may be removed and preserved separately during the washing process to prevent damage. These organisms can be placed directly in 10% formalin in a small vial. These “picking” vials must have an internal label matching the internal label in the sample debris, as well as the external label on the jar. The whole vial can be placed inside the larger sample jar upon preservation. The retention of fragile organisms in a picking vial is optional, as it may slow the washing process down. However, it can be helpful in the identification of organisms prone to fragmentation, and should be done as long as there is confidence in the labelling process.
Once the portion on the screen has been washed thoroughly the sediment is washed off the screen down into a corner of the pan for easy transfer into the sample jar. The washed sample fractions are then placed into an appropriately labeled container. The sample jar should have internal and external labels.
8. Post-washing Sample Processing
Organisms retained on the screen will be transferred to plastic zipper-type freezer bags or Nalgene leak-proof jars. Furthermore, non-breakable sample jars should be sealed with parafilm, doubled bagged for transport back to the laboratory facilities. Sample containers will be labelled internally and externally, then sealed in plastic 5-gallon buckets also labeled externally with sample numbers, date, and a hazardous materials (i.e., formaldehyde) warning label.
Jars should not be filled to capacity with sample debris, as this leaves little room for the preservative to mix into and penetrate the sample. Sample jars should be 50-75% full. If the sample is high in organic debris (e.g. wood fibre), the volume of debris should occupy no more than 50% of the jar. If the sample is more gravelly, the debris volume can maximally occupy 75% of the jar.
After the entire sample is washed, 37% buffered formaldehyde is added to the seawater such that its final concentration is 10% formalin (=3.7% formaldehyde; 10x dilution). The preservative is mixed in by gently rolling the sample jars after ensuring the lid is sealed tightly. Calcium carbonate chips are added along with the collection labels. Adhesive waterproof labels are placed on the exterior of the sample containers. Any samples taking up more than one container are labeled in series (ex “1 of 3”, “2 of 3”, etc.). Any extra vials (e.g. containing fragile organisms) are placed inside the sample jar. The lid is sealed with electrical tape for transport.
Samples can be held in buffered formalin for two to four days following collection, and then gently washed in portions on a 500-µm screen to remove the formalin and any residual fine debris. Use of a smaller sieve than that used in the field facilitated retention of smaller organisms and fragments. All material retained on the sieve was returned to the original sample container, and preserved in 70% (v/v) ethanol to prevent erosion of calcium carbonate structures. All original internal and external labels remained with the re-screened sample material. Additional borax can be added directly to the sample if the pH drops below 7.
9. Staining
Various stains such as rose bengal or phloxine B can be used at a concentration of 100 mg/l to make the organisms easier to see. A cautionary note is required, however, as the efficiency of many of these techniques differ between taxa. If these techniques are used, then the QA/QC performed on these samples should be designed to detect any differential sorting efficiency of taxa caused by the staining procedures. Rose Bengal is the most commonly used for marine invertebrates, since it targets protein, turning organisms pink or red and making them easier to separate from the sample sediment matrix.
Megafauna (large, rare organisms) should be removed during field washing. This includes in particular visible holothuroids (sea cucumbers) which are cut or punctured to allow the alcohol to penetrate and to facilitate future biomass determinations. Other large echinoderms, predaceous polychaetes and bivalves should be removed at this point and placed in a separate container to keep debris from entering the incisions.
REFERENCES
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PSEP 97a
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