Fish Sampling Techniques - Enviromental Sciences - Lecture Notes, Study notes of Environmental Science

Download Fish Sampling Techniques - Enviromental Sciences - Lecture Notes and more Study notes Environmental Science in PDF only on Docsity! FISH SAMPLING TECHNIQUES Key words: CPUE, Nets, Processing, Genetic sampling Fig – 11.1 INTRODUCTION Standardized sampling and data comparison methodologies are used in a wide variety of fields such as medicine, finance, education and agriculture. Standardized sampling is necessary to compare growth, condition, and population sizes of various lacusterine fish species among years and among lakes. Standardized sampling methodologies are also extremely important in fisheries and are required to evaluate how a fish population changes over time, or is functioning compared to an ―average‖ in a state or a region. Use of standard techniques allows biologists to concentrate resources on improving fish populations instead of routine monitoring considerations. This allows the biologist to identify problem fish populations, discover populations with exceptional angling opportunities, set regulations, or apply various management strategies and monitor their effects. Different methods were developed to capture the largest number of fish of various species in a majority of waters. It can be tempting to change sampling on a lake–by–lake basis to try to capture an even larger number of fish. However, the best results will be obtained by those biologists who adhere closely to standardized procedures so their data will be comparable to state averages where fish were collected in a similar manner. Application of these techniques whenever possible, even when just determining species composition, will improve your ability to evaluate lakes, and build a robust state database for comparison purposes. Docsity.com Timing the Survey Time of survey can greatly affect sampling data (Bettross and Willis 1988, Guy and Willis 1991). Fall surveys—should occur between the last week of August and the first week of October. Spring surveys—should occur between the last week of April and mid–June. Choosing between Spring or Fall—Large largemouth bass can most easily be captured in the spring while they are staging for spawning. However, yearling largemouth bass are still offshore during this time, and can be more easily captured in the fall. The biologist should determine which life history stage is of most interest and time the sampling accordingly. Never compare Spring to Fall samples and vice versa. Initiating the Survey Obtain standardized survey equipment—Survey equipment will consist of an electrofishing boat, standardized gill net(s) and standardized fyke net(s). Get map of the lake. Measure or obtain the shoreline perimeter— can be obtained easily from a map of the lake with a scale. Randomly select a starting point on the lake. Decide if it is feasible to electrofish the entire shoreline during the time allotted for the survey.  Entire shoreline can be sampled during the survey: This is possible most often in small– and medium–sized lakes. Start from the randomly chosen starting point and move around the shore. Shock for 600 seconds, work up fish, shock again for 600 seconds, work up fish, and continue this procedure until the entire lake is covered. For the last section, cover the amount of distance to reach the starting point (e.g., 278 sec, 342 sec. etc.) and stop. Do not re–shock part of the first section again to get 600 seconds. For setting gill and fyke nets, randomly choose sites. On small lakes it is possible to have a substantial impact on the existing fish populations if enough gill net sets are placed to detect a certain percent change. The biologist should use judgement to decide when to stop setting gill nets if the population may be substantially impacted, with the understanding that change may not be detectable from the few gill net sets.  Entire shoreline cannot be sampled during the survey: This is likely in larger lakes. Use the following procedure: • Mark sampling points on map of lake—from that starting point, put a mark every 400 meters (1300 feet) along the shoreline perimeter on the map6. These will be the ―sampling points‖ where you will start your electrofishing surveys and place nets. For a rough, but easy field estimate, take a piece of string, lay it on the map scale and mark it off at 400 m increments. Lay this string around the perimeter of the lake on the map and mark points on the map. Docsity.com several years before and several years afterwards, to measure trends in both ―treatment‖ and ―control‖ lakes are necessary to statistically validate that the change was related to the management action. This is most definitely the preferred situation if money and manpower are available. • Stratified Sampling: Normally you should not stratify unless there are clearly major differences between CPUE in large sections of the lake. Some of the computational drawbacks will outweigh the advantages. However, to reduce the variance and increase the ability to detect changes in CPUE, we can stratify the lake if it exhibits great differences in major habitat types. Larger lake and those with wide variations in habitat such as cliffs, rocky rip–rap, and weedy coves are good candidates. Some guidelines for stratifying: • Determine what fish specie(s) are of greatest interest or those which are the principal players. • Determine how to stratify based on habitat where CPUE of the ―principal player(s)‖ would probably be highest (e.g., weedy coves, largemouth bass; rock rubble, smallmouth bass, etc.). • Designate strata locations on the map—for example 1/3 of shoreline is highlighted as cliff (where biologist feels that largemouth bass CPUE would be low) and 2/3 of shoreline is highlighted as weedy habitat (where biologist feels that largemouth bass CPUE would be high). • Select needed sample size. These sample sizes are designed for simple random sampling and should, therefore, be more than adequate for stratified sampling. • Use one of two types of allocation methods to assign sampling sections to strata. o If the regional biologists can make an educated guess about the degree catch rates will be higher in one strata versus the other, use non-uniform probability allocation based on the degree catch rates might be different. o If you have no idea how much the catch rates will vary from one strata to another, proportionally allocate samples to strata based on size or ―weight‖ of strata. • Special considerations for net sampling—for net sampling, exclude those randomly chosen sampling points where it is impossible to set nets (i.e., no sheer cliff faces, boat launches, areas where turbines are, etc.). Then randomly select other sampling points to make up for those excluded. Standardizing Techniques on the Lakes: • Gill Nets Docsity.com Fig – 11.2 o Gill nets should be set in the evening before electrofishing starts and retrieved the next morning; o Nets should be set perpendicular to shore; o Smallest mesh size should be closest to shore; and o Although net–nights will be the unit of interest, record set time and pick up time. • Fyke Nets Fig – 11.3 o Fyke nets should be set perpendicular to shore; o Nets should be set in the evening/late afternoon before electrofishing starts and retrieved the next morning; o Record set time and pick up time; and o Try to set the net so the top of the first hoop is no more than about 1 foot under the water‘s surface. • Electrofishing Docsity.com Fig – 11.4 o Electrofishing should be conducted with pulsed DC, high range 100-1000 volts, 120 cycles per second; o Standardize power output of the electrofishing unit based on the conductivity of each lake. o Electrofish starting at each randomly chosen sampling point for 600 seconds as measured by the timer on the electrofishing unit. Always record on data sheets the actual number of seconds shocked (e.g., 578 sec, 600 sec, 605 sec, etc.); o Electrofish in the same direction from the sampling point for all samples; o Electrofish petal operations (continuous or intermittent) are at the discretion of the operator, and should be designed to capture the highest number of fish. Use intermittent shocking when approaching structure such as beaver lodges, downed trees, docks and weed patches. o Stay off the pedal until close to structure, then hit the pedal; o A minimum of two dippers and one driver should be in each electrofishing boat. o Catch rates go down if you electrofish the same section over again. o Never cover the same section that you have electrofished over again. o Make sure that when fish are worked up, they are released back at the start of the section, and not near the end where they can stray into the next section to be electrofished again; and o Electrofish at night to have the highest catch rates. Processing the Catch: Docsity.com Fork length is measured from the most anterior part of the head to the median caudal fin rays (fork of tail, Figure 2). This method is commonly used in Canada (Anderson and Gutreuter, 1983) and is only appropriate for fork tailed fish such as salmon, trout, and char. Another measurement sometimes used for salmon that have undergone morphological changes associated with breeding, is the post-orbital hypural length (MacLellan, 1987). Post-orbital hypural length is the distance from the posterior margin of the eye orbit to the posterior end of the hypural bone (last vertebrate). Table 11.2: Species measured for fork length or total length. Total length is the distance from the most anterior part of the head to the tip of the longest caudal fin ray when the fin lobe of the tail are pressed together. In BC, total length is the measurement used on fish without forked tails such as burbot and sculpins (Figure 2). Docsity.com Fig – 11.7. Diagram illustrating common length measurements for BC freshwater fishes. Standard length is the distance from the most anterior part of the upper jaw to the posterior end of the hypural bone. In applying this measurement, some other external landmark is often used instead of the hypural bone. This is normally the end of the caudal peduncle or the last scale of the lateral line (Anderson and Gutreuter, 1983). As well, measurements will often be made from the most anterior tip of the head as opposed to the upper jaw. Due to the variety of ways different observers define this measurement, standard length can often be confusing and inconvenient to use. Table 2 lists species for which fork length or total length is measured. All length measurements are recorded in millimeters (mm). Many rulers, tapes, calipers, or boards are available for measuring fish. The device used should be water-proof, light-weight, durable, easy to use, and offer adequate accuracy and precision. The units and type of length measurement should be clearly recorded on the data sheets and should be consistent for all the data collected during a sampling program. This will help to avoid confusion when the data is analyzed. Weight Weight measurements are normally made in grams (g) on whole fish that have been recently captured (whole wet weight). Excess water is drained or blotted from the animal with paper towel before measurement. As with the length measurements it is important to remain consistent in the techniques used and to carefully record units and methods in the field notes. Several different types of weigh scales can be used to make weight measurements in the field. Commonly used scales include toploading electronic balances, beam balances, and spring scales. One should attempt to match the accuracy of the scale to the size of the fish. For example, fry should not be weighed on a spring scale designed for adult fish. Toploading electronic balances are available in a variety of sizes and capacities suitable for field applications. Single scale models generally measure up to approximately 700 g, with accuracy greater than 0.01 g. Multiple scale balances can measure up to many kilograms, with accuracy of 0.1 g. Although flexible in their capacities, electronic balances require a power source and may not be useful for long, remote field trips. Also, their components may be sensitive to rough handling and inclement weather, conditions often experienced in the field. Beam balances can measure up to approximately 3000 g and are normally accurate to at least 0.1 g. Although they do not require electricity, they can be bulky, sensitive to rough treatment, and very difficult to use in bad weather. Pull type spring scales can measure weights up to 2000 g although the level of accuracy is not as high as beam or electronic balances. There are spring balances available that have a range of 0 - 10 g with an accuracy of ±1 g. While they may not be as accurate as electronic or balance beam balances, spring scales are small, durable, and do not require electricity. These attributes make them good for field work and as a backup to other weighing systems. Docsity.com Determination of Fish Age: The effective management of fish populations requires knowledge of the growth rate of the fish. This requires determination of the age of fish to develop a relationship between the size and age of fish. For an inventory, this information provides insights to evaluate the potential effect of harvesting on the population and to monitor the health of a population that may be affected by developments that affect fish habitat. Age can be determined directly or indirectly from the population of interest. Age and associated length or weight can be measured empirically from individuals reared in captivity or from fish specially marked at a known age and size, and recaptured at some later date. However, the cost and space required to rear fish often precludes the use of this as a practical method. As well, it can be argued that captive or marked individuals do not demonstrate growth typical of unmolested animals in the natural environment. For many species a direct measure of age can be made by analyzing hard body structures collected in the field. As fish grow, they deposit minerals in their skeletal tissues, producing characteristic growth patterns. In bones, these patterns are called annuli and in scales they are called circuli. Different periods of growth can be determined by counting the light and dark bands typical of annuli or by observing the differences in spacing of the circuli. By assessing these patterns the age of the fish can be determined. Otoliths, scales, fin rays, cleithrum, or the operculum are some of the typical structures collected for age information. The structure that is collected should depend on the needs of the inventory and the characteristics of the species collected. For example, scales cannot be used to age fish that are very long lived, such as lake trout, because the circuli near the center of the scale become very compressed and difficult to read accurately. Also, an ageing method that requires sacrificing the animal may not be desirable when studying sensitive populations. Regardless of the structures used, individuals experienced at ageing fish should be used as ageing is as much an art as it is a science. More detailed discussions on studies of fish growth can be found in Nielsen and Johnson (1983) and Bagenal (1978). Detailed descriptions of taking, preserving, and reading samples are provided in Mackay et al. (1990). Scales: Docsity.com Otolith: The sagittal otolith bones (sagittae) from the head of the fish are another structure used for ageing fish. ‗Otolith‘ is a generic term used for small calcareous particles that are present in fluid filled sacs in the fish‘s middle ear. The paired middle ears are located latero-posteriorly (behind and to the sides) to the brain. Otoliths possess a white centre surrounded by alternating concentric opaque and clear (hyaline deposits) rings. These structures assist in giving the animal its position with respect to gravity and allowing it to balance. Collecting otoliths require killing the fish and, hence, should only be performed when other, non-lethal methods cannot be employed. The removal procedure is easily learned, relatively quick, and requires only basic dissecting tools. Otolith bones collection is the preferred method for determining age in species that do not produce reliable scale readings, or grow to great ages. The otoliths should be extracted unbroken and as clean as possible, using small forceps. Both bones should be removed (Mackay et al., 1990). Once removed from fish, any residual tissue, gelatinous membrane or blood should be rinsed from the otolith with fresh water. Other methods that can be used to remove otoliths and the procedures can be found in Jearld (1983) and MacLellan (1987). Generally otoliths can be preserved by air drying or by freezing. If stored dry, otoliths may become brittle and easily damaged by rough handling. A solution of glycerin, glycerin/water, or glycerin/alcohol can be used to preserve otoliths and prevent them from becoming dry and fragile. Glycerin has also been known to have a mild clearing effect on the otoliths making them easier to read. Formalin should not be used to preserve otoliths or fish from which otoliths may later be taken. The formalin tends to de-calcify bone resulting in chalky otoliths where the annuli are obscured. Otoliths are generally stored in small, labeled envelopes. If preserved with a liquid, the bones should be kept in a small, sealed container with a label placed inside the container with the bones. Cleithrum and Operculum: Cleithra or opercular bones are structures that can be useful for ageing pike, walleye, or perch. However, this procedure requires killing the fish and thus should be justifiable. The operculum is easily removed with dissecting scissors and cutting along the anterior boarder of the gill cover. The cleithrum is a bony structure that supports the posterior border of the gill cavity and is usually covered by the posterior portion of the gill cover. Freshly removed structures can be frozen for up to 2 months before they are analyzed. Prior to analysis they should be cleaned, soaked in hot water to remove excess tissue and oils and allowed to air dry for several days. After drying the cleithra should be read within 2 weeks. If left too long, cleithra turn opaque making it difficult to discern the annuli markings. Docsity.com Sexing and Maturity: • Determination of Sex The most accurate way to determine the sex of most species of fish is through an examination of internal sex organs. In adults, eggs are usually obvious in the ovaries and in males the testes are typically smooth, whitish organs along the dorsal surface of the body cavity. The sex organs of immature fish can be hard to find but generally they will appear as long, thin organs along the dorsal surface of the body cavity; females will be a pinkish colour while males will be translucent to whitish. Some species of trout/salmon develop specific secondary sex characteristics during the spawning phase and the larger the fish the more obvious the distinction. External observations of trout in spawning colour would include body shape and jaw shape. Females will tend to have a rounder girth while larger males may develop a slight hook in the jaw. However, these changes may be difficult to distinguish. For reconnaissance level inventory it is not necessary to sacrifice fish for determining its sex, unless the animal is collected as a voucher specimen. • Determination of Maturity Recording the maturity of specimens is important information as the onset of sexual Johnson (1983) present detailed classification schemes for categorizing the maturity of fish. Accurate determination of maturity is best accomplished through direct observation of the gonads. However, classification can also be done based on external observations. The following provides basic descriptions for 6 stages of sexual maturity (brackets include abbreviations for coding): 1. Immature (IM): Young individuals that have not yet reproduced; fish with underdeveloped gonads. 2. Maturing (MT): Ovaries and testes begin to fill out and take up a large part of the body cavity; eggs distinguishable to the naked eye. 3. Mature (M): Fish in full spawning colours; gonads at maximum size; body cavity feels full; especially females; roe or milt is not produced if the body cavity is lightly squeezed. 4. Spawning (SP): Fish in full spawning colours; eggs and milt are expelled when body cavity is lightly squeezed (also referred to as gravid). 5. Spent (ST): Still have spawning colours; eggs and sperm totally discharged; body cavity feels empty and genital opening is inflamed; gonads empty except for a few remaining eggs or residual sperm. 6. Resting (R): Adult sized fish; spawning colours not as apparent; gonads are very small and eggs may not be visible to the naked eye. FISH PRESERVATION TECHNIQUES Docsity.com Fig – 11.11 Careful and correct preservation procedures in both the field and laboratory are important for ensuring the quality of the collected specimens or tissues. Fixatives of the correct concentration, appropriate containers, clean and sharp dissecting tools, waterproof data form/labels, and complete observations will all affect the quality and value of the sampling. Preservation techniques vary depending on how the samples will be used. All voucher specimens must be submitted in 50% isopropyl alcohol in the prescribed jars. The following sections outline some of the most common techniques, and describes the Ministry‘s requirement regarding the submission of samples. • Voucher Specimens Voucher specimens are representative samples of species identified in the field, collected and preserved to verify the field identification. Only one representative sample of each red/blue listed species should be collected. For species that are neither rare nor endangered, two to three specimen can be collected. These specimen should represent the size variability encountered at the sampling site. Any mortalities that occur during fish capture for sampling can be submitted as voucher specimens. • Field Preservation o Anaesthetizing to kill All fish must be killed prior to fixation. This is can be achieved by leaving the fish in high doses of the anaesthetizing solution. This is an ethical treatment of a live animal and also serves a scientific purpose: anaesthetized fish relax and can be preserved in a more natural state. o Fixatives Docsity.com Fig – 11.12 Ideally, one would wish to collect the appropriate tissues without sacrificing the donor. Tissue samples required can vary from organ tissue, to fin clips, scales and epithelial tissue. Many tissue preparation techniques are available and laboratories vary in techniques. Before sampling begins, sampling protocol should be confirmed with the lab conducting the analyses to ensure that appropriate samples are taken. This section describes basic field collection and storage techniques for genetic samples. The type and amount of tissue required will depend on the analyses to be conducted. There are three basic categories: (1) Protein electrophoresis: requires organ tissue including heart, muscle, eye and kidney either fresh or fresh frozen, stringent quality requirements. (2) DNA analyses with no PCR amplification: requires relatively large amounts of tissue often fresh and in buffer solution, stringent quality requirements. (3) DNA analyses with PCR amplification: requires very little tissue (e.g. fin clip, scale) and can be preserved in ethanol or dried; fairly lax quality requirements where some tissue degradation is often not a problem. The third category is becoming increasingly popular as conservation and non-lethal sampling requirements become more of an issue. In addition, this sampling for this category is far less arduous in the field and requires very little equipment. Collection and Preservation of Parasites and Other Health Related Specimens Fish can be infected with a variety of internal and external parasites. Sampling specifically for parasites can take a great deal of time and is not within the scope of a reconnaissance inventory. Given the number and types of parasites that may infect fish, no single sampling procedure can be recommended. Much of the thorough examination for parasites requires a full necropsy and use of stereo or compound microscopes to examine the tissues. It is best that this work be done Docsity.com back at the lab or some permanent station where a parasitologist can be consulted to verify observations. Parasites infect almost any tissue group. If sampling for parasites is a priority, the entire fish should be retained and preserved for laboratory analysis. Ideally, live fish offer the best information as some parasites will leave as the fish dies. Preserving the entire fish in a fixative should only be done if the previous alternative is not possible. Docsity.com