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The collection of deep water samples requires that at least one member of the sampling group be very familiar with boat operation and safety. If the sampling trip involves the use of a boat, then the weather forecast or marine conditions should be obtained prior to departure from home. If conditions are poor, then the sampling trip should be postponed.
Samples collected for bacterial analysis are simply grab water samples collected at designated sites. They can be collected at near shore or deep water stations. Deep samples require the use of a Van Dorn bottle for the sample collection (Figure 1). All bacteriological samples must be cooled (to 4°C) as soon as they are collected and must be shipped to the lab as soon as possible. The following are the protocols for collecting shore samples, deep water surface samples, and depth profile samples.
To avoid contamination from suspended sediments, the sample collector must wade out past the point where wave action affects the lake bottom. In most cases, this distance is not far from shore. The sampler should not exceed a depth where there exists a reasonable possibility that water might unexpectedly enter the gum-boot or hip-wader. This is particularly important during colder weather when getting wet poses a health risk (such as hypothermia).
PROTOCOL
(shore samples)
(a) Obtain labeled, sterilized, 250 mL bottles and wade into the lake at the most accessible point.
(b) Once you reach a sufficient depth (where bottom material will not interfere with the sample), stop and orient yourself towards the center of the lake. Often samplers wait standing in place for 2 - 3 minutes to ensure sediment disturbed by wading can settle.
(c) Remove the lid and hold it aside without allowing the inner surface to touch anything.
(d) With your other hand, grasp the bottle well below the neck. Lean out towards the center of the lake and in one continuous motion plunge the bottle beneath the surface to a standardized depth (two frequently used depth for surface samples are 0.1 m and 0.5 m) and slowly force it through the water until it is entirely full. This motion creates a current over the mouth of the bottle such that water entering the bottle has not come in contact with your hand.
(e) Pour out enough water to provide 2.5 - 5 cm air space above the sample. Replace the cap immediately.
(f) Return to shore and pack the sample in a cooler containing a sufficient quantity of ice packs (twice the volume of ice to samples in summer months or equal volume when sampling in winter months).
PROTOCOL
(for collecting surface samples at deep stations)
(a) The person at the bow (front) should always collect the samples. This is because the bow is the anchor point and even in a slow current the boat will drift so that the bow is upstream. This precaution reduces the potential of contamination from the boat or motor. The person in the stern (rear) can be responsible for holding the boat's position (when not anchored), taking the field measurements, and making field notes, and helping to stabilize boat when bow person is working near gunwales.
(b) Obtain a labeled, sterilized, 250 mL sample bottle and remove the lid without touching the inner surface of either the bottle or lid.
(c) Reach out an arm length from the boat to take the sample. Ensure that the person in the stern is providing counterbalance (working over the opposite side of the boat).
(d) Plunge the bottle under the surface and move it slowly towards the current (the direction the boat is facing) until the bottle is entirely full.
(e) Pour out enough water to provide a 2.5-5 cm air space above the sample.
(f) Recap the bottle and place in a cooler immediately.
Deep samples are usually collected by Van Dorn sampler. Note that Van Dorn samplers are available in both horizontal and vertical configurations. The advantage of the vertical configuration is that the water within the open bottle is flushed out as the bottle is lowered so that one can be guaranteed that the water was collected from the indicated depth. The advantage of the horizontal configuration is that a very narrow depth range is sampled. Vertical configurations are usually used in large lakes. Horizontal configurations are used for samples to be collected at or just above/below a very sharp thermocline, or to be collected near the lake bottom. Horizontal configurations are mandatory for very shallow lakes.
PROTOCOL
(for taking deep samples)
(a) Open the Van Dorn sampler by raising the end seals. Do not touch any of the inner surfaces of the apparatus.
(b) Set the trigger mechanism as in Figure 1.
(c) Lower the sampler to the desired depth. Ensure that the loose end of the rope is tied to the boat.
(d) Send the messenger down to "trip" the mechanism that closes the end seals.
(e) Retrieve the sampler to the surface.
(f) Flush the drain valve with some of the sample water so as to reduce the possibility that bacteria from previous sample sites will contaminate the current sample. Then, transfer the water sample from the Van Dorn bottle to sterilized, labeled, 250 mL sample bottles via the drain valve. Leave a 2.5-5 cm air space at the top of the bottle. Take care to avoid contact with the drain spout as contamination often occurs at this stage.
(g) Recap the bottle and place in the cooler immediately.
Zooplankton, or planktonic animals, are free floating and suspended in open or pelagic waters. They are generally collected with a conical net (Figure 2) that has a specific mesh size (ranging from as small as 64 µm to as large as 256 µm). Small mesh openings will clog more readily than larger ones, but small organisms will pass readily through larger openings. The mesh size required for a particular lake will depend on the productivity of the lake and the purpose of the study. The size of the mesh and the diameter of the net opening used for each BC Environment project will be clearly outlined in the project design. The preferred net mesh, when appropriate, is 64 µm with a net mouth diameter of 20 cm.
The net is lowered to a particular depth and pulled directly up through the water column (known as a vertical tow). Alternatives to the vertical tow are the horizontal and oblique tows in which various strata of the lake are sampled individually (a horizontal tow) or as a composite (oblique tow). These are elaborate techniques that require specialized equipment rigged to the boat and a tow net that has remote open and close capabilities. Unless there is specific need for data from horizontal and oblique tows they are not used. Therefore, the vertical tow is the only protocol that will be mentioned here.
PROTOCOL
(vertical tow)
(a) Ensure rope is securely fastened at the plankton net opening and that the dead end is tied to the boat.
(b) At the designated site, lower the net to depth outlined in the project design.
Note: The actual distance that the net travels through the water must be recorded and the total volume of water that passes through the net must be calculated (see Figure 3).
(c) In smaller lakes, haul the net hand over hand with a steady, unhurried motion at a rate of 0.5 m/s. In large lakes, when long net hauls are conducted, use a davit, meter wheel, and winch. The maximum tow speed used should be 1 m/s.
(d) Once the net is at the surface, wash the net by raising and lowering the net body below the net mouth in the water. Then squirt de-ionized water against the outside of the netting and from the top downward. This washes any adhered plankton down into the cod-end (removable `cup' at the end of the net).
(e) Disconnect the cod-end and carefully decant the water and plankton into a pre-labeled bottle. Rinse the cod-end several times, pouring each rinsate into the bottle (this ensures that all plankton are collected).
(f) Wash the net by rinsing (pulling it through the water without the cod-end). This is an absolute necessity before proceeding to the next sample site (particularly between lakes).
(g) Preserve the sample with 70% ethanol (70mL of 100% ethanol for each 30 mL of sample volume) and place in the cooler for shipping.
Figure 3. Tow volume calculation
Phytoplankton collection in the open water of a lake consists of surface and/or deep grab sampling. A Van Dorn bottle is the most common tool for collecting the deep samples.
PROTOCOL
(for collecting surface phytoplankton samples)
(a) Anchor the boat at the designated sample site (alternatively, if the water is too deep or a buoy is not present, the person in the stern will have to maintain position while the person in the bow takes the samples).
(b) Obtain a labeled, 1 litre sample bottle and remove the lid without touching the inner surface of either the bottle or lid.
(c) Reach out an arm length from the boat to take the sample. Ensure that the person in the stern is providing counterbalance (working over the opposite side of the boat).
(d) Plunge the bottle under the surface and move it slowly towards the current (the direction the boat is facing) until the bottle is entirely full. Standard sampling depths for "surface" water samples are 0.1 m and 0.5 m.
(e) Preserve the sample with 1-3 mL of Lugol's solution (1-3 mL per 1 litre of sample). Different water chemistry and density of algal material require different concentration of preservative. A general guideline is that there be sufficient Lugol's to turn the sample the colour of weak tea.
Note: Plankton nets for phytoplankton samples are not recommended for quantitative samples. They are size selective and very non-quantitative. They can be used however for identification of the species present as the higher density assists in the taxonomic work, particularly of rarer species.
PROTOCOL
(for collecting deep water samples)
(a) Open the Van Dorn sampler by raising the end seals.
(b) Set the trigger mechanism as in Figure 1.
(c) Lower the sampler to the desired depth (epilimnion, hypolimnion or thermocline - the location of which should have been determined by prior DO/temperature profile data collection - see Ambient Freshwater Sampling chapter). Ensure that the loose end of the rope is tied to the boat.
(d) Send the messenger down to "trip" the mechanism that closes the end seals.
(e) Retrieve the sampler to the surface.
(f) Transfer the water sample from the Van Dorn bottle to labeled, 1 liter sample containers via the drain valve.
(g) Preserve the sample with Lugol's solution (1 to 3 mL per 1 liter of sample).
(h) Recap the bottle and place in the cooler.
Benthic invertebrates in lakes or large slow moving rivers are generally collected in the same fashion as sediment samples (see Lake and Stream Bottom Sediment Sampling document). The processing of the sample once it has been collected is where the techniques differ. The type of grab sampler to be used at a particular site will depend on the site conditions and the purpose of the study. The equipment to be used will be dictated by the project design and must be outlined in the field log book and pre-sampling checklist. Figure 4 presents some of the grab samplers that are available, while Figure 5 presents the core sampler most commonly used.
Figure 5. Kajak-Brinkhurst Sediment Core Sampler
PROTOCOL
(boat sampling with a grab sampler)
(a) Ensure that the rope is securely fastened to the sampler and that the loose end of the rope is tied to the boat.
(b) Set the grab sampling device with the jaws cocked open (see
Figure 4). Great care should be taken while dealing with the device while it is set as
accidental closure can cause serious injuries.
(c) Lower the sampler until it is resting on the sediment (its own weight is adequate to penetrate soft sediments). At this point, the slackening of the line activates the mechanism to close the jaws of the Ponar and Petersen grabs.
(d) For the Ekman grab, send the messenger down to `trip' the release mechanism.
(e) Retrieve the sampler slowly to minimize the effect of turbulence (that might result in loss/disturbance of surface sediments).
(f) Place a container (i.e., a shallow pan) beneath the sampler just as it breaks the surface of the water.
Note: If the jaws were not closed completely, the sample must be discarded. Discard the sample into a bucket if the second collection attempt is made from the same general area. Dump the unwanted sample only after the "real" sample has been successfully collected.
(g) Place a sieve between the sampler and the pan and gently open the jaws and allow the sediments to empty into the sieve. The size of the sieve mesh will depend on the purpose of the study, but a common mesh size is 0.20 mm (200 µm). This size represents the practical lower limit for general study of benthic organisms. It is not as crucial to have small mesh size when the only analysis to be conducted is biomass.
(h) Immediately record (in the field log book) observations regarding the appearance of the sediment (i.e., texture, colour, odour, presence of biota, presence of detritus).
(i) Rinse the sieve with de-ionized or on-site water to remove as much sediment as possible.
(j) Transfer the organisms to a pre-labeled sample bottle and preserve with 70% ethanol. Formalin may be used as a fixative for initial preservation but should be subsequently washed and transferred to 70% ethanol.
PROTOCOL
(boat sampling with a core sampler)
(a) Open the valve and set the trigger mechanism (as per Figure 5). Ensure the rope is securely fastened to the corer and attach the loose end of the rope to the boat.
(b) Most corers are designed to be simply lowered into the sediments and fill the core tube by their own weight and need not be dropped from any hight. Consideration of the type of corer used and the nature of the sediment being sampled will need to be taken into account.
(c) Send the messenger down to release the trigger mechanism.
(d) Carefully retrieve the sampler and place a stopper into the bottom opening before removing from the water to prevent loss of the sample.
(e) Remove the core tube or liner from the corer and stopper the upper end.
Note: Once on shore, the sample can be treated as a bulk sample or it can be sectioned and the organisms separated from the sediment in strata.
(f) For bulk samples all the sediment may be sieved as per the grab samples above. Otherwise, the sediment should be sectioned in regular intervals as it is extruded (record the thickness of each stratum the length of entire core). Each stratum may be sieved and its contents placed in pre-labeled sample bottles or, the unsieved sediments can be placed directly into pre-labeled sample bottles.
(g) Preserve the samples with 70% ethanol.
Note: It is preferable to section the core as soon as possible after it is retrieved. As the sediment warms, it tends to expand in the core tube. With warming, decomposition gases are liberated at a much faster rate and if they bubble through the core, they will disturb the stratigraphy.
Aquatic plants are collected for one of three purposes: first, biomass studies for the determination of how much plant material is present on an area or lake basis; second, tissue analyses for the detection of metals, pesticides and plant products; or third, taxonomic analyses as part of ecological or impact studies (permanent records of community structure over time) and to serve as reference specimens for the above two analyses. These reference specimens are a valuable scientific record and their collection and subsequent handling should be done with care so that much of the time and expense that has gone into their collection is not wasted. Reference specimens are stored in herbaria which provide permanent records of what was found and allow identifications to be re-verified if necessary. Even when the major thrust of the work is for biomass studies or tissue analyses, representative specimens must still be collected and saved as vouchers of the species that were analyzed. With respect to sampling for taxonomic purposes, Warrington (1994) documents techniques regarding the processing of individual taxa within each of the three groups of aquatic plants (floating, submergent, and emergent). This manual presents a general overview of the protocols for collecting aquatic plants for taxonomic purposes.
Generally, the whole plant should be collected. Some groups cannot be identified to species without mature fruits or flowers. Large plants are pressed and mounted on a 30 by 40 cm white card that is dimensionally stable when wetted. Small plants like the duckweeds do not make satisfactory pressed and dried specimens. In this case, small screw cap vials make good collecting and preservation containers.
PROTOCOL
(small floating species collection)
(a) Scoop a few individual plants into a pre-labeled vial.
(b) Preserve the specimens in a solution of 70% ethanol, 25% water and 5% formalin.
(c) These specimens must be submitted to a herbarium where they will be identified to species and stored for future reference.
PROTOCOL
(collection of large emergent or rigid plants for taxonomic purposes)
(a) Field notes should be written directly on the card stock before you place the plants onto the card (you will not be able to write on the card once it is wet). The notes should be written in pencil in the lower right hand corner of the card where it will later be covered by the permanent label.
(b) Collect an entire specimen and store it until conditions are appropriate to mount.
Note: Do not leave them out in the sun even briefly as they will wither very quickly and become useless as specimens. Emergent plants should not be submerged, but kept in a bag with a little water in the bottom to maintain high humidity. It is best to keep each species in its own bag and all the bags from one lake or site together in one large (garbage) bag.
(c) To mount each specimen, lay the plant on the card stock with the roots in the bottom left corner (fold over the top if it is too tall to fit).
Note: Do not cover the label area in the bottom right corner.
(d) Spread out leaves and flowers, turn some over so the bottoms can be seen, and try to make a neat and tidy specimen that covers the whole sheet.
Note: For small plants fill the sheet with more specimens from the same clump or clone, to show as much variability as possible.
(e) For fruiting plants where the seeds may be shed on drying, collect the seeds into small paper or cellophane pouches and attach these pouches to the finished herbarium sheet.
(f) After mounting the aquatic plant on the card stock, place a piece of heavy blotting paper on top of the specimen to help dry the plant quickly.
(g) Wrap the card stock, plant, and blotter in a newsprint folder. The newsprint should be 30 by 90 cm in size and folded in half to form a folder in which the mounted plant is placed.
(h) Once you have accumulated several of these wrapped packages (each containing one card stock/specimen), place them into a plant press with a piece of corrugated cardboard separating each package.
Note: The corrugations should run in the same direction so that air flow through the press is facilitated.
(i) Drying should take place within several days to prevent fungal growth and rotting and to preserve colours and shapes as much as possible. If you will be in a laboratory or herbarium the same day, the plant presses may be dried in a proper plant drier or a forced draft oven at 40°C. In the field, use motel hot air registers, baseboard heaters, or hair dryers to move warm air through the corrugated cardboard. If the weather is dry, secure the plant press on the roof of the truck and allow air to blow through the corrugated cardboard as you drive from site to site. As the plants in the presses dry it will be necessary to retighten the presses periodically, at least daily, to maintain the pressure and hold the plants flat.
(j) These specimens must be submitted to a herbarium where they will be identified to species and stored for future reference.
PROTOCOL
(collection of large submergent or
non-rigid plants for taxonomic purposes)
(a) Field notes should be written directly on the card stock before you place the plants onto the card (you will not be able to write on the card once it is wet). The notes should be written in pencil in the lower right hand corner of the card, where it will later be covered by the permanent label.
(b) Collect an entire specimen and store it until conditions are appropriate to mount.
Note: Do not leave them out in the sun even briefly as they will wither very quickly and become useless as specimens. Keep them in a bag or bucket of water at all times until you are ready to press them. It is best to keep each species in its own bag and all the bags from one lake or site together in one large bag.
(c) Since these plants are flaccid and clump together when removed from water, they need to be floated onto the card stock and arranged neatly to keep them from becoming a useless mat. Start by slowly lifting the bottom of the card stock out of the water at the root end and arranging and spreading the plant as you continue. Once a portion is out of the water, it will stay in place. Some, but not all, plants will allow a limited amount of rearranging once they are out of the water. Hold the card stock with one corner down and let most of the excess water drain off. Floating is best done in 36 by 50 cm photographic trays or directly in the lake from which the specimen was collected (it will be difficult in lakes if there is any wind or waves). A tray on a picnic table at a site with running water is an ideal mounting and pressing site.
Note: Do not take plants to a different lake to float them onto the card stock; you risk spreading weeds from lake to lake.
(d) Continue as in (e) through (j) on the previous page.
Plant tissues may be collected for analyses of metals, pesticides, nutrients, plant products, for dry weight/wet weight ratios, or for other laboratory analyses. In all cases an entire, intact, voucher specimen of each species should be collected and filed in a herbarium as a record of what was analyzed. The procedures outlined above under the section headed Taxonomy (4.1.5.1) should be followed.
For analyses of chemicals, it may be necessary to analyze distinct portions of the plant separately to determine where the material is localized in the plant (the portion to be collected will be outlined in the project design).
PROTOCOL
(collecting samples for tissue analysis)
(a) Collect entire specimens and keep them submerged and covered until they can be processed (plants should never be allowed to desiccate).
(b) Place each specimen in individual air tight bags (e.g., Zip Lock) or tissue cups or glass bottles (for analysis of trace organics). Ensure each is fully labeled. Ensure the quantity of tissue and the type of container are appropriate for the analysis that will be conducted (see Appendix 2 of this chapter).
(c) Place the sample container in the required cooler for shipping (some analyses need to be frozen, therefore, these samples will be placed in a cooler containing a sufficient quantity of dry ice). Always wear gloves when handling dry ice.
Biomass studies, mass of plants per unit area, are not routine and are usually conducted as part of a research project. They are not performed as part of standard sampling programs. They often require the use of SCUBA to map out and harvest the designated area. Refer to Warrington (1994) for further information regarding biomass protocols.
For fish collection and processing protocols, refer to Fish Collection Standards and Methods (RIC 1997), which has replaced the Fish Collection, Preservation, Measurement and Enumeration Manual (Triton Environmental Consultants, 1994) as the standard manual. The Field Key to Freshwater Fishes of British Columbia (RIC 1994), is used to identify fishes in the field.
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In some instances, the protocols for collecting biological specimens in rivers are similar to those used for sampling in lakes. But in the case of benthic fauna and fish, the collection techniques are considerably different. The following are the protocols for the collection of biological samples from flowing waters.
As is the case for lakes, bacteria are collected as grab samples and immediately cooled to 4°C to minimize metabolic activity (growth/reproduction) until they can be analyzed by the laboratory.
Wherever practical, samples should be collected at mid-stream rather than near-shore. Samples collected from mid-stream reduce the possibilities of contamination (i.e., shore effects - back eddies, seepage from near shore soils, etc.). The most important issue to consider when deciding where the sample should be collected from is SAFETY. If the flow is sufficiently slow that the collector can wade into the stream without risk, then the sample can be collected at a depth that does not pose a threat (discretion is the key - never wade into water that appears deep or fast flowing). When conditions dictate that the sample be taken from the stream bank, deviations from the standard protocol should be accurately documented in the field log book and transferred to the database as soon as possible.
PROTOCOL
(for wading into flow)
(a) Obtain a labeled, sterile 250 mL bottle and wade into the river downstream from the point at which you will collect the sample. Wade upstream to the sample site. This ensures that you will not disturb sediments upstream from the sample point.
(b) Stand perpendicular to the flow and face upstream.
(c) Remove the lid and hold it aside without allowing anything to touch the inner surface.
(d) With your other hand, grasp the bottle well below the neck. Plunge it beneath the surface with the opening facing directly down, then immediately orient the bottle into the current.
(e) Once the bottle is full, remove it from the water by forcing it forward (into the current) and upwards. Pour out some water to ensure there is an air space.
(f) Replace the cap immediately.
(g) Return to shore and place the sample in a cooler with sufficient ice packs (twice the volume of ice as the sample in summer, same volume in winter).
PROTOCOL
(for sampling from the stream bank)
(when the current is too strong, water is too deep, or ice is too thin)
(a) Secure yourself to a solid object on shore (with a safety harness and line if necessary).
(b) Remove lid from a labeled, sterile 250 mL bottle and place into a clean resealable bag (e.g., Zip Lock).
(c) Hold the bottle well below the neck.
(d) Reach out (arm length only) and plunge the bottle under the water and immediately orient it into the current.
(e) When the bottle is full, pull it up through the water while forcing into the current. Pour some water out so as to leave an air space.
(f) Immediately recap the bottle.
(g) Place the sample in a cooler with sufficient ice packs (twice the volume of ice as the sample in summer, same volume in winter).
Note: If conditions are such that sampling from a bridge is an option, refer to the chapter, Ambient Freshwater and Effluent Sampling, 4.2.2, for the protocol.
Periphyton are defined as attached species of microflora (algae) and are primarily found on hard, immobile surfaces such as large stones (cobbles, boulders) and on aquatic plants. Analysis of periphyton communities, particularly for the purpose of biomass calculations, involves collecting the algae from a known surface area. To do so, the following pieces of equipment are required:
· A piece of equipment to outline a known surface area and contain the removed biomass - for example, a toilet bowl plunger that has had the handle insertion hole cut larger. The diameter of the hole is known, therefore the surface area of the exposed rock below the hole is known. A rim of neoprene is glued around the hole to prevent water and algae from escaping when collecting the sample. It is also possible to use a template if the algal mass is gelatinous or coherent enough to be picked off the growing surface. A template is a piece of flexible rubber or plastic with a square or round hole cut in it to define the area to be scraped.
· A water bottle (squirt bottle) containing de-ionized water
· A tooth brush (to scrape the algae free from the rock)
· A turkey baster (to transfer the sample from the scraping cup to the sample bottle
· Pre-labeled sample bottles
· Preservative (Lugols)
PROTOCOL
(natural substrate)
(a) Choose a location of fairly uniform substrate over wetted width. Try to work in upstream but random direction. Select rocks that are relatively flat and large enough to accommodate at least three scraping cups. Unless the project design states that rocks with the most dense growth are to be sampled, the sample collector should select the rocks randomly. To do so, the collector should not specifically look for an area from which to sample, but instead should wade into the water and stop without regard for apparent algal concentration and then grab the first rock that is suitable for scraping. Return to shore with the selected rock and repeat the process until 3 or 4 rocks are collected. Don't attempt to lift rocks that are too heavy and don't enter water that may pose a threat to your safety. Watch your footing while returning to shore with the rock.
(b) Hold the scraping cup firmly over a selected patch on a rock. Squirt some de-ionized water into the cup. With the tooth brush, scrape the algae (within the cup) off the rock. Each new algal patch that is scraped can be referred to as a disc.
(c) Transfer the water-algae mixture to a pre-labeled bottle (a turkey baster with the tip cut off works very well as the transfer tool). Do not remove the cup yet.
(d) Squirt some fresh de-ionized water into the cup, re-scrape and collect the rinsate. At this point you have collected one disc. Collect and composite more discs until you have transferred a sufficient quantity of algae to the sample bottle [see (e)].
(e) If algal density is low or patchy, collect at least five discs from each rock. If algal density is high or homogeneous, then take fewer discs per rock, but ensure that you use at least five rocks to characterize the periphyton at the site. Always record the total number of discs collected.
(f) If the sample is to be analyzed for biomass, place the bottle in a black garbage bag and then immediately into a cooler that contains a sufficient quantity of ice packs (twice the volume of ice as the total sample when sampling in summer, equal volumes when sampling in winter).
(g) For measurement of biomass, the samples are prepared for chlorophyll analysis by filtering the sample onto a membrane filter (0.45 or 1.0 _ pore size), then folding this filter and placing it inside a larger (8 to 10 cm.) Whatman type paper filter labeled (date, site, area scraped), then clipped with a plastic paperclip and placed into a jar with desiccant and placed in the freezer or in a cold cooler for shipment to the lab.
Note: In washing and transferring the sample, keep the volume of water used to a minimum. Filtering periphyton samples can be very difficult due to suspended sediments that generally accompany the periphyton.
It may be necessary to use a larger pore size filter than for phytoplankton (typically 0.45 µ). Periphyton cells are typically much larger than phytoplankton. An analysis of the filtrate from the periphyton sample will indicate if significant biomass is passing through the larger filter size chosen.
(h) If the samples are to be analyzed for taxonomy (only two or three discs are required), then the sample must be preserved with Lugol's solution. Use at least 1 mL of Lugol's per 250 mL of sample (with heavy biomass, more preservative is needed). No filtration is required. Tip once or twice to mix preservative and sample water.
(i) Different conditions or circumstances may require different methods than described here. Other methods are acceptable as long as they are described and documented and approved by Ministry authority. For example, where heavy periphyton growth occurs, a method for sampling heavy growth could use a template disc to define a sampling area and scrape the area around it, then remove the disc of periphyton with a scalpel. This method reduces the volume of wash water that must be used and potential clogging of filters.
Samplers used for the collection of invertebrates in the shallow portions (<one metre deep) of rivers or streams fall into two categories: those sampling natural substrates, and those using artificial substrates. Two of the more common samplers used for natural substrates are the Surber and Hess samplers (Figure 8). The Drift net sampler can also be used to collect the emerging or drifting invertebrate stages. Kick nets are also used for inventory type surveys,
Natural Substrate. The advantage of these types of sampler is that the sample reflects the natural community of the stream. The disadvantages are that such samples have a very high variability and many samples need to be taken to characterize the benthic community. The large number of samples necessary generally means relatively high cost.
The recommended mesh size is 210 µm (microns) for most samples. In some specialized studies a different mesh size may be appropriate. It may be appropriate to collect the samples using a 210 µm mesh size but to do a preliminary sieving at 500µ. The sample material retained on the 210 µm mesh is stored to be processed later if the program design calls for it.
Figure 8. Common benthic macro-invertebrate samplers
The Surber sampler consists of two interlocking frames that support a capturing net. One frame outlines the area of stream bed to be sampled while the other supports the net. The sampler is intended for use in shallow (30 cm or less) flowing waters. Use A1 tent pegs (hooked) to anchor surber in fast moving water. Repetitive sampling should be timed (i.e., 5 minutes each for more uniform sampling).
PROTOCOL
(Surber sampler)
(a) Choose a wetted width location where substrate is fairly uniform. Provide a description of general habitat (e.g., is this a run, pool or riffle section, etc.). Position the sampler securely at a random location on the stream bottom parallel to water flow with the net portion down-stream. The mesh size of the net should be compatible with the goal of the program.
Note: Take care not to disturb the substrate upstream from the sampler.
(b) Carefully turn over and lightly rub all rocks and large stones that lie within the frame. This process dislodges organisms that are clinging to the stones. Examine each large stone for organisms, including larval or pupal cases, that may be clinging to the stone before discarding (downstream or to the side of the sampler). In order to maintain comparability between stations, a limit on time spent on handling and rubbing the substrate should be set (5 minutes recommended).
(c) Stir remaining gravel with your hands to a depth of 5 to 10 cm.
(d) Move the sampler upstream to a new randomly selected patch of stream bottom and repeat steps (a) through (c). Continue this process until five patches of stream bottom have been sampled, each upstream from the last. This creates a composite of the five samples. The total area sampled will depend on the size of the sampler (x 5) and should be calculated and recorded in the field log book.
(e) Return to shore and carefully invert the net into a shallow pan containing stream water. Ensure all invertebrates are rinsed from the net into the pan.
(f) Transfer the organisms into a pre-labeled plastic sample bottle and preserve with 70% ethanol (may be initially fixed with 10% formalin ). Rinse sample net after each use.
The Hess sampler is a metal cylinder with a screened opening on one side and an opposite opening with a net attached (Figure 8). The sample collector places the Hess sampler in the stream with the screen oriented into the current and the net trailing behind. The water is able to flow freely through the sampler and out through the net. With a known radius, the stream bed area that is sampled is easily calculated. This value must be recorded in the field log book.
PROTOCOL
(Hess sampler)
(a) Position the frame securely on the stream bottom. Ensure the screened opening is facing into the current and the net portion is trailing down-stream. Hold the sampler in position by applying pressure with your knees.
Note: Take care not to disturb the substrate upstream from the sampler.
(b) Reach into the cylinder and carefully turn over and lightly rub all rocks and large stones. This process dislodges organisms that are clinging to the stones and washes them into the net. Examine each stone for organisms, including larval or pupal cases, that may be clinging to it before discarding it outside of the cylinder. In order to maintain consistency between samples, a standard time should be assigned to sampling each site (5 minutes recommended).
(c) Stir remaining gravel with your hands to a depth of 5 to 10 cm.
(d) Move the sampler upstream to a new patch of stream bottom and repeat steps (a) through (c). Continue this process until five patches of stream bottom have been sampled, each upstream from the last. This creates a composite sample of the five areas.
(e) Return to shore and carefully wash the contents of the net into the cod-end then transfer to a shallow pan. Ensure all invertebrates are rinsed into the pan.
(f) Transfer the organisms into a pre-labeled sample bottle and preserve with 70% ethanol. Rinse sample net after each use.
Drift net samplers are designed to be anchored in flowing water to capture macro-invertebrates that have migrated or have been dislodged from the bottom surfaces into the current. They are limited to use in small, shallow streams. Ideally, drift nets should span the entire width of the stream that is being sampled. Several nets can be placed across the stream channel to capture all drift organisms and to measure spatial variation in the drift.
PROTOCOL
(Drift Net sampler)
(a) Anchor drift nets in water sufficiently shallow that they will extend above the water surface. Set drift net samples for the time specified by the project design (the length of time is designed to collect a representative sample, but not so long that flow through the nets becomes impaired by clogging from captured particulate matter).
(b) Transfer the organisms into pre-labeled sample bottles.
(c) Preserve with 70% ethanol and place in cooler. Initial fixation may be done with 10% formalin.
(d) Record time, area of net opening, stream discharge per unit time (see the Ambient Fresh Water and Effluent Sampling chapter, 6.7), and volume of water filtered in the field log book.
Artificial substrates. The above techniques sample the benthos directly from the natural substrate. Another technique is to use artificial substrate and to place it in the stream so it is colonized by the organisms in the stream then removed later and the community analyzed. These types of samplers are best suited for upstream/downstream studies or studies designed to test for changes over time. They do not necessarily provide a representative sample of the actual community which is living in the stream.
There are advantages and disadvantages to artificial substrates.
Advantages:
· Allow collection of samples from locations that can't be sampled because of substrate or depth
· Reduced variability
· Non-destructive sampling of a location
· Flexibility in sampling design
Disadvantages:
· Colonization rates differ from site to site
· Species in sampler may be different than stream bottom
· Long incubation/exposure times (6-10 weeks)
· Vulnerability of samplers to vandalism
The most frequently used artificial substrate sampler is the "barbecue basket" sampler. The basket sampler is fabricated by filling the basket (available in a number of variations from hardware stores) with gravel (2.5 to 7.5 cm diameter) that is then placed in the stream bottom. The substrate becomes colonized and is removed after some predetermined length of time.
PROTOCOL
(basket sampler)
(a) Place the basket sampler in the stream and anchor if necessary. Leave in place for the necessary colonization time.
(b) When the samplers are removed, take particular care not to dislodge organisms from the sampler. A general technique is to carefully place the basket sampler in a plastic bag underneath the water before it is lifted out.
(c) Record time, any site related data such as flow, temperature and pH (see the Ambient Freshwater and Effluent Sampling document), and data about the appearance and condition of the basket sampler in the field log book.
(d) In the lab, remove the animals from the sampler by carefully washing each rock into a sieve. Transfer the organisms into pre-labeled sample bottles. Preserve with 70% ethanol and place in cooler. Initial fixation may be done with 10% formalin before transfer to ethanol for longer term storage.
Due to the similarity in the techniques, refer to 4.1.5 (in lake sampling) for the protocols for collecting and processing macrophytes in rivers.
Refer to Fish Collection Standards and Methods (RIC 1997) for protocols involving fish collection and processing.
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