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Survey and Study Techniques

Surveys

Introduction

Mussel search strategies can be qualitative or quantitative. Qualitative strategies are useful for determining presence/absence of common species and qualitative diversity. More detailed qualitative strategies allow for estimates of species richness, presence/absence of rare species, or distribution. Quantitative strategies are useful when estimating specific parameters such as density, relative abundance, age distribution, recruitment, and mortality. The ideal search strategy encompasses both strategies. Published information on survey methodologies can be found in Dunn (2000), Hornbach and Deneka (1996), Miller and Payne (1988; 1993), Payne et al. (1997), Strayer et al. (1997), Vaughn et al. (1997), and Young, et al. (2001).

It is best to know one's objectives first, including the benefits and costs associated with each method. No matter which method is chosen, there are several approaches available to the collector based upon the type of collecting gear to be used.

Before collecting any live specimens a review of current local, state and federal regulations is advised. For contact information for the metro area state and federal offices, go to the Collecting Permits page.

Hand-picking typically involves walking or wading along the shoreline of a lake or stream and visually searching or feeling (when visibility is low) for specimens. Specimens are often exposed in desiccated pools or as beach wrack, particularly during dry seasons. Surveys in these areas can be undertaken by visually searching along shorelines in dried up ponds. Live aestivating mussels are often located in this manner. Spent shells, sometimes found in shell heaps known as middens that are left behind by foraging raccoons and muskrats, are also easy to find using nothing more than a good pair of eyes. However, these specimens will not contain unbiased samples of the mussel community (Walters, 1994c).

Specimens are also collected by hand from shoreline debris or various benthic substrates. When in flowing water, it is best to walk upstream so as to flush sediment behind the viewer. A useful tool for hand-picking is the look box, a clear glass or plastic-bottom cone-shaped box placed on the surface of a shallow river or lake and viewed through the top to find specimens on the substrate. Such a tool is often utilized when hand-picking in shallow depths where visibility is high. A makeshift look box can easily be assembled by removing the bottom from a five gallon bucket and attaching in its place a circular piece of clear plastic with epoxy. Little or no equipment is necessary and large areas may be covered quickly while hand-picking. However, this technique may overlook smaller, less familiar specimens and the risk of stepping on fragile organisms in the substrate is increased.

Net collecting, a modified form of hand-picking, involves sweeping a net across a lake or stream bottom to gather mussels and shells at the surface of the substrate. The net is attached to the end of a pole and is typically made of a strong wire bent into a ring. A sack-net of loose burlap or mesh copper screen attaches to the ring. After several sweeps across the substrate, the net is rinsed to remove mud and fine sand. Coarse material is removed and examined for mollusks, and finer material may be retained in glass jars and sorted at a later time. Samples garnered by net collecting tend to be more random, an advantage over hand-picking when arbitrary population size estimates are required. One disadvantage to this method is that firmly entrenched living specimens will often not be retrieved by the passage of the net, so this form of collecting must be limited to sand, mud, or grassy substrates

Sieving requires a mesh screen of a chosen size supported by a wooden frame. A given volume of substrate is scooped up with the sieve, and sand and mud are sifted in the water for the desired specimens. These specimens are then sorted into a metal dissecting pan, which has a white-colored bottom that makes identification of small individuals easier. Sieving, unlike hand-picking and net collecting, provides the means to collect burrowing animals. However, compared to the other methods, sieving is more time consuming and labor intensive.

Trawling involves dragging a mesh trawl net behind a boat, or dropping the trawl net some distance away and hauling it to shore with a rope. The trawl net is dragged along the bottom where it picks up benthic organisms. Trawling works particularly well for large mussels and can estimate density per unit area. But the time-consuming nature of this technique, as well as the fact that it usually requires a boat, puts trawling at a disadvantage over other methods.

A benthic grab sampler (Ekman or PONAR) is a metal sampling device that closes around a substrate sample on a lake bottom and scoops a portion of the substrate into metal jaws. A metal screen across the top of some grab samplers prevents the sample from escaping. The sample is then retrieved to a boat or bridge crossing and emptied, exposing specimens and sediments. A grab sampler is particularly useful for deep water species, substrate samples, or collecting juveniles and can be used in most substrates except packed clays. However, this method is of little use in shallow areas and tends to be more costly and time-consuming. A mussel rake is a long-handled garden rake with blunted tines. A wire mesh basket is attached behind the tines. As the rake pulls through the substrate, mussels are gathered into the basket. The basket is then stirred in the water to remove loose sediments and the specimens retrieved. Mussel raking is particularly useful for attached mollusks or those lodged in the substrate. A mussel rake makes a welcome addition to ordinary hand-picking. Unfortunately, specimens smaller in diameter than the spaces between the tines of the rake tend to be missed and specimens may be damaged during the collecting process.

Snorkeling utilizes a mask and snorkel to search the substrate for mussels while swimming. A wet-suit is highly recommended for snorkeling during cooler months or for extended periods in the water. Advantages to snorkeling include increased spatial area and depth of surveys and less disturbance to natural communities. Disadvantages include greater equipment costs and more time- consuming surveys.

SCUBA sampling involves diving in water too deep to be sampled by other methods and blindly feeling around the sediment surface for specimens. A one meter quadrat is often used to maximize sample randomness. Another method divers use involves hammering a one-half meter long steel stake into the substrate at a chosen depth and attaching a five to ten meter rope marked with electrical tape at one meter interval sections. The diver then swims slowly in circles around the stake steadily letting out more rope at one meter increments following each complete revolution until a circular substrate area is completely surveyed to a desired radius. The diver gently drags both hands back and forth across the top few centimeters of sediment, feeling for mussels. Fingerless dive gloves work best for this type of searching. Avoid swimming too quickly to prevent the sediment from stirring up, making the already poor visibility even poorer. Slow, regular swimming at full extension of the rope works best. SCUBA sampling allows little bias from sample conditions and an estimate or specimen density per unit area can be obtained (density = no. of specimens/radius>2). The cost for SCUBA gear and mandatory training may make this technique prohibitively expensive. Also, SCUBA sampling is not practical for investigating large areas at one time.

Other methodologies exist (such as side-scan sonar), but the ones listed are the most economical, require little or no training, and are the most familiar to the casual collector.

Study Techniques

Preservation/Fixation

Live mussels may be relaxed prior to preservation by placing specimens into glass or plastic containers with enough water to bring the water level to 2-3 centimeters above the largest specimen. Menthol crystals (dissolved in water) slowly applied until a very light film is formed on the water surface serve as a relaxing agent. The container must be sealed for 24 hours (6-12 hours for small specimens). After the allotted time has passed, mussels should begin to gape open with the foot extruding. Next add more menthol (about half as much as was originally added) and reseal the container, periodically inspecting specimens for activity by gently probing the leading edge of the foot or mantle. Small cork stoppers are particularly useful for pegging open the valves to prevent subsequent closure. When contractile response is slow or non-existent, the animal is relaxed. Siphon off the remaining relaxing fluid and add enough fixative (see below for suggested fixatives) to cover all specimens. Particularly large specimens may need to be injected with fixative at various points using a syringe.

Chemicals

For most specimens, 10% formalin (formaldehyde gas bubbled in water) works well for fixation and 70% ethyl alcohol (EtOH) works best for preservation. Other techniques may be used out of necessity or need. The most inexpensive and effective fixative and preservative for preserving large numbers of specimens is formalin at 5% (1:19 with water) for fixing and 10% (1:9 with water) for preserving. Formalin hardens tissues by denaturing the proteins and stabilizing them against further chemical change. Because formalin is an acid, it must be buffered before being applied to shelled mollusk as the acid destroys the shell. Buffered formalin is available commercially or, when unavailable, crushed marble chips (one-half pound per gallon) can be added as a buffer. Because formalin is noxious and irritating to eyes and skin, it is wise to work outside or in a well-ventilated area using latex gloves. Advantages to using formalin are low cost, low quantity per usage, wide range of applications, and non-flammable nature. Disadvantages include its toxicity, acidity, and incompatibility with molecular study techniques.

Denatured ethyl alcohol (EtOH) makes a good all-around preservative and can also be used as a fixative. The standard preserving solution is 70%. Specimens may be fixed by running them through an alcohol series (30% for 24 hours, 50% for another day, then 70% a day later) before preserving to prevent shrinkage and excessive water loss into solution. Advantages to using ethyl alcohol are well-preserved specimens, lack of toxicity, and applicability to molecular study techniques. Disadvantages include high cost, flammability, and difficulty in obtaining.

Isopropyl alcohol (rubbing alcohol) may be substituted for ethyl alcohol at 40-50% as a preservative. It can also be used as a fixative at 30% for 24 hours then switched to 40 or 50%. An advantage to using isopropyl alcohol is its ease in obtaining and lower cost. It can be acquired at most drugstores. Disadvantages include high cost, flammability, and slightly poorer quality specimen preservation when compared to formalin or ethyl alcohol.

Recently, Lellis et al. (2000) advocated the use of 500 ppm of buffered MS- 222 (tricaine methane-sulfonate), available from most chemical supply companies as a relaxing agent. Advantages include rapid relaxation time (30-60 minutes) and the benefit of full recovery, if necessary. Disadvantages include decreased availability and high cost.

Data Labels

It is important to label specimens immediately upon collection. Important information, if kept separate or memorized, can easily be misplaced, forgotten, or assigned incorrectly later. When labeling, always use pencil or India ink on acid-free paper, if possible, as these lend better to permanence. Label information includes four general categories: taxonomy, locality, collection, and habitat.

Taxonomy includes, but is not limited to, family (Margaritiferidae or Unionidae for freshwater mussels; Corbiculidae for Asian clams; Dreissenidae for zebra or quagga mussels; and Sphaeriidae for fingernail clams), scientific name, authority and date. Locality information includes, but is not limited to, map locality (including UTM or latitude/longitude), specific locality, water body, distance and direction from nearest city or town, county, and state. Collection information includes, but is not limited to, collectors, identifiers and collection date. Habitat information includes flow direction and rate, depth, substrate, water chemistry, temperature, and surrounding land use.

See the bibliography for additional sources of information.

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