• View in gallery
    Figure 1—

    Photograph of a selection of contemporary radio transmitters (A), microacoustic transmitters (B), and PIT tags (C). The scale is in centimeters.

  • View in gallery
    Figure 2—

    Site of loss (expulsion) of a transmitter (circle) that was (A) and was not (B) associated with the incision used for transmitter insertion.

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  • 123. Close DA, Fitzpatrick MS & Lorion CM, et al. Effects of intraperitoneally implanted radio transmitters on the swimming performance and physiology of Pacific lamprey. North Am J Manag 2003; 23:11841192.

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  • 128. Connors KB, Scruton D & Brown JA, et al. The effects of surgically-implanted dummy radio transmitters on the behaviour of wild Atlantic salmon smolts. Hydrobiologia 2002; 483:231237.

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Surgical insertion of transmitters and telemetry methods in fisheries research

A. Michelle Wargo RubFish Ecology Division, Northwest Fisheries Science Center, National Oceanic and Atmospheric Administration, 2725 Montlake Blvd East, Seattle, WA 98112.

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Niels JepsenSection for Freshwater Fisheries Ecology, Institute of Aquatic Resources, Danish Technical University, 8600 Silkeborg, Denmark.

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Theresa L. LiedtkeColumbia River Research Laboratory, Western Fisheries Research Center, US Geological Survey, 5501A Cook-Underwood Rd, Cook, WA 98605.

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Mary L. MoserFish Ecology Division, Northwest Fisheries Science Center, National Oceanic and Atmospheric Administration, 2725 Montlake Blvd East, Seattle, WA 98112.

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E. P. Scott Weber IIIDepartment of Medicine and Epidemiology, School of Veterinary Medicine, University of California-Davis, Davis, CA 95616.

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Use of electronic transmitter and monitoring systems to track movements of aquatic animals has increased continuously since the inception of these systems in the mid-1950s. The purpose of the present report is to provide information about veterinary principles and their incorporation into surgical implantation procedures for fish. We also intend to provide insight into the unique challenges of field-based aquatic surgical studies. Within this context, 4 aspects of the process for surgical implantation of transmitters in fish (ie, handling, aseptic technique, anesthesia, and implantation) will be described. Effects of surgical insertion of transmitters (ie, tagging) and aspects of the surgical implantation process where collaboration and professional exchanges among nonveterinarian researchers and veterinarians may be most fruitful will be discussed. Although this report focuses on surgical implantation, the principles and protocols described here (other than incision and suture placement) are also applicable to studies that involve injection of transmitters into fish.

A Brief History of Fish Telemetry

The earliest use of telemetry in fish dates to 1956, when cumbersome electronic backpacks were externally attached to adult coho salmon (Oncorhynchus kisutch).1 Soon thereafter, the transmitter package was made smaller and the capabilities of the transmitter were expanded. Within 2 decades, telemetry techniques were firmly established and in common use throughout the world. By 1975, researchers were tagging hundreds of individuals from 40 fish species.2 Since then, telemetry has been used to investigate the movements of tens of thousands of fish as well as the movements of aquatic mammals, reptiles, amphibians, and invertebrates.2

Both sonic (also known as acoustic) and radio technologies were used in the earliest telemetry investigations. Sonic transmitters were used in both freshwater and marine environments. However, early sonic models were not extremely effective in turbulent environments because air bubbles attenuated the acoustic signal. Radio signals were effective in freshwater, even in a highly turbulent, air-enriched environment; however, radio signals were rapidly attenuated in saltwater. Both radio and sonic transmitters consisted of electronic components and a battery potted in epoxy or encapsulated within a watertight casing. Most radio transmitters also possessed a trailing antenna to increase transmission range.

A common attachment method for both types of transmitters involved threading pins, wires, or sutures attached to a transmitter through the dorsal musculature of a fish. Alternatively, a transmitter was inserted into the stomach of a fish (gastric implant). Gastric insertion of a radio transmitter with an antenna required that the antenna be positioned such that the free end of the antenna exited the fish's mouth. As telemetry was applied to more species, new external attachment methods were developed to accommodate a variety of body forms and habitats.3

With the advent of long-lived, miniaturized acoustic transmitters in the 1970s, fish researchers used surgical implantation as a means of transmitter attachment. Implantation of transmitters directly into the body cavity had several advantages, including eliminating motion hindrance for fish with externally mounted equipment and reducing transmitter loss associated with external attachment or gastric implantation (through regurgitation).4 The first attempts at surgical implantation of transmitters into fish were reported in 1975.5 Acoustic transmitters were used, and various suture materials and suturing techniques were evaluated. Results for flathead catfish (Pylodictis olivaris), large-mouth bass (Micropterus salmoides), and striped bass (Morone saxatilis) in that study5 formed the basis for most surgical methods currently in use.

Soon thereafter, new techniques were attempted for surgical implantation of radio transmitters with a trailing antenna. Initial attempts involved coiling the antenna and implanting it within the peritoneum.6 However, researchers modified this technique so that the antenna exited through the body wall and trailed alongside the fish to improve signal propagation and thus detection range. A shielded-needle technique was devised to reduce the risk of damaging internal organs during this step and to improve postoperative recovery and healing.7 At that time, recommendations were formulated for maximum transmitter weight relative to fish body weight,6 which enhanced rapid expansion of fish research that relied on telemetry.

The underlying assumption that fish with transmitters behave in a manner representative of untagged conspecifics is a fundamental principle for telemetry studies. Early research efforts focused primarily on methods to reduce fish fatalities after tagging and to increase the amount of time that fish were capable of apparently normal movement.6 However, as the variety of transmitters increased and the scope of tagging studies expanded, there was an increase in the need for attachment techniques that minimize sublethal tagging effects.8 Fish researchers conducted studies9–11 to determine the effects of transmitter attachment on a variety of fish functions, including feeding activities, swimming capacity, buoyancy control, spawning behavior, and susceptibility to predation. Results of those studies have provided fish researchers with new insights into the capabilities and limitations of telemetry techniques in fish.

Contemporary Telemetry Studies

Currently, fisheries researchers can choose from a wide variety of electronic transmitters, including radio-frequency identification transmitters (eg, PIT tags) and active radio or acoustic transmitters (Figure 1). Passive devices require an external energy source for transmission of signals, whereas active devices require a power source (eg, a battery), which enables them to transmit signals at preprogrammed intervals. A variety of sensors for factors such as pressure, temperature, and electrical activity can also be incorporated into or attached to telemetry transmitters. Information from sensors can be actively transmitted along with the transmitter's unique identification code or stored within the tag for subsequent retrieval. However, to retrieve archived data, those tags must be physically recovered from the fish.

Figure 1—
Figure 1—

Photograph of a selection of contemporary radio transmitters (A), microacoustic transmitters (B), and PIT tags (C). The scale is in centimeters.

Citation: American Journal of Veterinary Research 75, 4; 10.2460/ajvr.75.4.402

The range of technological options has evolved to enable researchers to address diverse study objectives, such as providing information on fish physiology, behavior, movement, and habitat utilization. These objectives can be achieved exclusively through telemetry devices or by combining telemetry with other methods, such as physiologic and genetic sampling. Contemporary studies range from basic monitoring of a few (10 to 30) fish over relatively short distances to more complicated monitoring that may involve tens of thousands of fish. These larger studies may involve evaluating the performance of fish over hundreds of kilometers or at multiple points along a migration corridor, such as fish passage structures, spillways, or turbines at dams.

Many telemetry studies are conducted on threatened or endangered species in an effort to provide resource managers with sufficient information to design and implement appropriate management actions. Telemetry allows researchers to collect data that would be virtually impossible to obtain through other approaches, and telemetry has the promise of providing precise measures of performance. As such, use of this technology is becoming more commonplace. Because of this widespread use, researchers are under increasing pressure to confirm that transmitters and tagging protocols do not substantially affect the results of their studies. In particular, protocols for transmitter attachment have come under increased scrutiny by government agencies, funding organizations (eg, National Science Foundation), and even professional journals, which stipulate that tagging methods must be demonstrably humane as well as scientifically sound.

Because of their small size and cylindrical shape, most PIT tags can be injected into fish through a 12-gauge needle attached to a custom injection device.12,13 Depending on the species and size of the fish being tagged, transmitters have been injected into the peritoneal cavity, dorsal sinus, pelvic girdle, and various muscles. Several attachment methods are available for active transmitters,9,11 but contemporary researchers are primarily opting for surgical implantation. However, although this is an area of interest, detailed procedures for surgical implantation of transmitters currently are not common in the literature. Rather, most of the published surgical techniques or protocols have been developed by isolated research groups or agencies for use with a given species and within a specific context. As such, there is a lack in consistency, and they are not consistently applied and do not consistently conform with sound veterinary principles. Furthermore, although training in surgical techniques for fish is available to veterinarians through coursework affiliated with academic institutions and professional outreach programs, similar opportunities are virtually nonexistent for nonveterinary researchers.

A 2004 survey14 of 177 researchers who were actively using surgery for insertion of telemetry devices in fish indicated that they had learned transmitter implantation techniques primarily from a combination of observation of others performing the surgeries, evaluation of the literature, and instruction by mentors. Less than 10% of respondents had received professional instruction provided by an educator or veterinarian through an academic or professional development course.14 Furthermore, although most respondents reported that surgical experience was important for ensuring a successful outcome, approximately two-thirds reported that they had little experience (performed surgery on only 1 to 10 fish) before participating in their first telemetry study, and 13% indicated that they had performed implantation surgeries during their first study without having prior surgical experience.

Analysis of the responses for this survey14 indicate substantial room for improvement in the teaching and development of surgical technique for implantation of telemetry transmitters, and authors of the survey concluded that nonveterinarian fish surgeons could benefit from interaction and consultation with veterinarians. Unfortunately, comments of the survey participants also indicated that some researchers were hesitant to approach veterinarians for advice. This reluctance was attributable to a perception that veterinarians lacked specific instruction or training in fish medicine or did not have sufficient experience working in field conditions to offer practical assistance. Authors of a 2011 report15 have attempted to change this perception and to foster collaboration between veterinarians and fisheries researchers by outlining the role of veterinarians in surgical implantation of electronic transmitters in fish.

Fish Handling in Field Studies

Fish implantation surgeries are most often performed on the deck of a boat, at a makeshift tagging station on a dock, or near the water's edge. Surgery locations are often remote and without amenities such as power and fresh water, thus posing logistical challenges to researchers. As such, the most important considerations regarding design and conduct of studies are often the most simple and are related to fish handling rather than to surgical technique.

The overall goal should be to minimize handling of fish, and the importance of basic planning to enable researchers to achieve this goal cannot be overstated.8,16 Correct handling of fish can mean the difference between a successful tagging study and one that fails, regardless of the surgical procedure used for insertion of the transmitter. Netting fish and removing them from the water can elevate concentrations of analytes (eg, cortisol and glucose) thought to be related to stress.17–20 Additional actions such as pursuing, netting, and crowding fish can exacerbate these responses.21–25

In many field studies, fish are tagged at the sampling site and are placed in holding tanks only long enough to allow recovery from anesthesia prior to release. In these situations, researchers should always ensure that there is adequate tank space for holding fish prior to and after surgery as well as containers to facilitate the transfer of fish through each stage of the tagging procedure. Some studies will require additional confinement (eg, for extended postsurgical monitoring or suture removal) or transport to another location for tagging or release. These studies will likely require additional accommodations for holding or transport of fish. Arrangements for these additional accommodations should be made prior to the day that fish are scheduled to be tagged. Fish containers should provide a dark environment and have lids or covers to minimize disturbance and prevent escape.25 These simple measures will help ensure that stress-inducing situations are minimized.

Obtaining adequate amounts of freshwater or saltwater from a fish's environment is also essential because this water may be needed to address deteriorating water quality or to speed recovery from anesthesia. Intuitively, water quality in containers should be maintained at the highest possible standards. Oxygen concentrations should be monitored and carefully maintained near saturation, but supersaturation with oxygen should be avoided. Compressed air or oxygen can be pumped through diffuser stones into the containers. Critical thresholds for dissolved oxygen will differ with exposure time and water temperature as well as with fish species and size.26–29

The pH of water in tanks, containers, and anesthetic baths should be consistent with that of the original source water. The pH can be altered by fish respiration and the addition of anesthetics. Test kits or strips can be used to monitor pH, and sodium bicarbonate can be added as needed for buffering. To prevent changes in pH caused by the by-products of respiration, a neutral environment should be maintained by ensuring that fish are not overcrowded or overstimulated and by refreshing tank water frequently.

Frequent water changes are also a way to ensure a consistent water temperature. A rule of thumb is that water temperature in holding containers should be maintained within 2°C of source water temperature.16 Tagging should be scheduled and performed only when local water temperatures are within published thresholds for the specific fish species. There are thermal guidelines for use of some anesthetics (eg, tricaine methanesulfonate [MS-222]); thus, water temperature may limit or restrict their use. Performing surgery in conditions near or outside thermal tolerance limits can exacerbate handling stress, increase the risk of secondary infection at the incision site, or directly result in fish fatalities.30–34

Transfer of fish between holding tanks or containers should be kept to a minimum, and sanctuary nets should be used if possible. A sanctuary net contains a reservoir in the bottom that retains water to ensure that netted fish remain submerged. The reservoir also protects the skin and eyes of fish from rubbing or bumping against the netting. Sanctuary nets (sometimes referred to as solid-bottom nets or sock nets) can be purchased from aquaculture equipment suppliers.a Nets of this nature can be custom made by lining the bottom third of a standard dip net with a soft, nontoxic, impermeable material such as vinyl. Another effective means of benign transfer is to gently cup fish in the palm of one's hands. Researchers should be aware of possible scale and mucus loss. Fish should be handled gently when placing them on measuring boards, weighing balances, or surgery platforms. During all procedures, the gills and skin should be kept moist. This may require that shades be erected or that fish be draped with plastic or bubble wrap to prevent desiccation. Although careful attention to handling will not guarantee a successful outcome, inappropriate handling can result in serious sublethal effects or even fatalities.

Finally, researchers should establish minimum standards for fish health before beginning a tagging project. The maximum proportion of descaling that will be tolerated, whether fish with visible lesions will be tagged, and the acceptable parasite load for participating fish are examples of criteria necessary to optimize study outcomes. Detailed information for developing these criteria has been reported elsewhere.35

Aseptic Techniques

For homeothermic animals, aseptic technique is considered a routine component of surgical protocols. However, this is not always the case for telemetry implantation surgeries in fish. Aseptic techniques and maintaining asepsis in an aquatic (and often remote) environment can be challenging. Furthermore, because of sensitivity of the integument of fish to most disinfectants and sterilants, certain aspects of aseptic techniques (eg, applying chemical disinfectants to the incision site) can be counterproductive or even harmful. For these reasons, aseptic practices have been inconsistent and less stringent for implantation surgeries conducted on fish in field conditions than would be expected for similar procedures conducted on homeotherms.36

In particular, there is inconsistency for the practice of transmitter sterilization. Heat sterilization methods are damaging to electronics; therefore, only chemical options are available for transmitter sterilization. Unfortunately, most chemicals capable of sterilization are highly toxic to both fish and humans. Furthermore, these chemicals generally require prolonged contact times (eg, 6 to 24 hours), and there are regulations regarding their use and disposal. As such, sterilization of transmitters is often thought to be unsafe, cumbersome, and impractical for application in field conditions.

Furthermore, there is a lack of evidence in the literature that pathogen transmission has occurred as a direct result of surgical tagging. Infections have been reported in fish after tagging37; however, these were described as secondary infections rather than primary infections introduced through a breach in asepsis. Only a few surgical tagging studies38–40 have been designed specifically to evaluate the infection risk associated with surgical methods. In all these studies, the authors concluded that aseptic practices (eg, sterilizing transmitters and surgical instruments) were without merit for healthy fish tagged in normal conditions. Investigators in 1 study38 surgically implanted pike (Esox lucius) with radio transmitters in nonsterile conditions in a field setting and did not provide prophylactic or postoperative treatments. Implanted fish were released into a reservoir; 1 year later, all implanted fish were recaptured and had no observable negative effects related to the surgery or implants. In another study,39 investigators tested whether the introduction of untreated lake water into the coelom had negative effects on survival rate and healing of bluegill (Lepomis macrochirus). Their results indicated no positive effects of reducing water entry through the incision or for the use of sterile equipment. In a similar study,40 survival rate, growth, and healing were evaluated in juvenile Atlantic salmon (Salmo salar) tagged by use of nonaseptic and aseptic techniques; investigators found no benefits for the use of sterile techniques.

These results have prompted researchers to consider whether requirements for asepsis should be relaxed for fish implantation surgeries performed under certain conditions. Veterinarians and other experts who are asked to consult on telemetry implantation studies in fish need to be aware that skepticism exists toward the potential benefits of aseptic practices for surgical implantation, and consultants should be prepared to provide justification (ie, documentation) for the use of aseptic techniques (including sterilizing transmitters) as well as to provide practical means for use of these methods in field conditions.

Considering the increase in the number of telemetry studies being conducted with endangered and threatened fish populations, optimizing the use of aseptic practices may be critical for successful long-term monitoring. For example, the authors are not aware of any published evidence that pathogen transmission has occurred during transmitter implantation surgeries. However, there is evidence that coded-wire tagging procedures enhance transmission of Renibacterium salmoninarum, the causative agent of bacterial kidney disease, among juvenile Chinook salmon (Oncorhynchus tshawytscha).41 Furthermore, water-borne transmission is the primary route of infection for many pathogens of fish.42 Pathogens can be transmitted among individual fish or populations of fish during several steps of the tagging process. For example, pathogens may be transferred by vectors used for fish handling (eg, buckets, nets, and boots), surgical instruments, suture material, and transmitters or they can be directly introduced through the surgical incision. From a regulatory perspective, there are increasing requirements by institutional animal care and use committees as well as some journals that fisheries researchers comply with standards developed primarily for studies of homeotherms. These standards invariably dictate that instruments and implants be sterilized for use during surgical procedures in which the body cavity is entered and the patient is intended to survive the procedures for subsequent postsurgical monitoring.

Despite the actual and perceived challenges posed for researches conducting telemetry implantation studies, there are general aspects of tagging procedures that are similar between a veterinary hospital setting and field settings. These include disinfection of the surgical environment, surgical instruments, and transmitters. Standard practices for asepsis can be adapted to provide effective and practical methods and allow researchers to develop defensible protocols for the specific conditions of each telemetry study.

Asepsis in field settings—Equipment for fish telemetry procedures typically includes tanks, buckets, nets, air stones, tubes and hoses, measuring boards and weighing balances, containers for transfer of fish, instrument containers, and a tagging platform. Any or all of this equipment can serve as a vector for pathogen transmission. Potential pathogens must be limited to a particular water source or group of fish. Hence, hardy, nonporous, and noncorrosive materials should be used. At a minimum, this equipment should be disinfected between locations at which tagging procedures are performed, after each tagging session, and before working with a new cohort of fish.

Disinfectants should be chosen on the basis of their efficacy against local pathogens, ease of use, and ease of disposal. The efficacy of some disinfectants commonly used in aquaculture facilities has been tested against specific pathogens of fish.43–47 However, a disinfectant may be selected on the basis of its efficacy against a specific class or group of pathogens. Currently, the only disinfectant with labeled efficacy against some fish pathogens and approval by the EPA for use in aquaculture is an oxidizing agent with potassium peroxymonosulfate as the active ingredient.b This product is available as a powder that can be mixed with water and applied as a spray or via immersion. A minimum contact time of 10 minutes is typically required for disinfection. When used appropriately and depending on the location, this product may be discharged into the water, thus affording an important convenience for field settings. However, it is illegal to use this product in some states (eg, California); therefore, researchers should always communicate with state and local authorities regarding regulations for chemical use and disposal before selecting or using any products.48

Additional chemicals used for disinfection of fish handling equipment include biguanides,c–f iodophores,g–j quaternary ammonia compounds,k–m chlorine products,n and other oxidizing agents.o Of these chemicals, chlorine products are specifically not recommended for use in field settings because of their extreme toxic effects on fish and potential to remain lethal to fish even if the products volatilize.

Another goal for asepsis is to protect humans from exposure to potentially harmful zoonotic pathogens such as Mycobacterium spp,49 although the risks posed to researchers will depend on the source of water and species of fish involved. Thus, all researchers involved in handling fish should wear impermeable gloves of appropriate length to provide a barrier for broken or unprotected skin. Researchers performing implantation surgeries should wear medical-grade examination gloves. For high-volume tagging operations, it likely will not be practical or feasible to change gloves between subsequent individual fish. However, at a minimum, gloves should be changed between cohorts or batches of fish, when the integrity of the gloves is compromised, or when gloves become grossly contaminated with blood or feces.

Fish researchers should wear comfortable clothing, which should be kept clean and dry. Closed-toed footwear is recommended to prevent injuries that could result from dropped scalpels, other surgical instruments, or heavy equipment. Similar to fish handling equipment, clothing that comes in contact with fish (ie, boots, waders, reusable gloves, and neoprene sleeves) should be washed or disinfected between tagging locations and at the end of each tagging session.

Researchers in the United States should be aware of the potential for transfer or spread of aquatic organisms such as the zebra mussel (Dreissena polymorpha) or New Zealand mud snail (Potamopyrgus antipodarum) and take appropriate precautions to mitigate such transfer or spread. Specific precautions may also be required to prevent the transmission of pathogens such as viral hemorrhagic septicemia virus, infectious hematopoietic necrosis virus, infectious pancreatic necrosis virus, infectious salmon anemia virus, Myxobolus cerebralis (the causative agent of whirling disease), and Rhabdovirus carpio (the causative agent of spring viremia of carp) in North America. Production and research facilities are required to have government-mandated policies for preventing introduction, amplification, and spread of pathogens in fish. These policies dictate sanitation procedures that must be performed at any specific facility.

Transmitters—Best practice guidelines dictate that implants be sterile.36 Unfortunately, as mentioned previously, sterilization of transmitters is inconsistently performed prior to telemetry implantation in fish. This is primarily because of concerns that chemical sterilization is hazardous to researchers and difficult to accomplish. In light of these concerns, we believe that veterinarians can greatly improve the level of asepsis practiced during telemetry implantation studies by providing nonveterinarian researchers with instruction regarding the safe use and disposal of liquid sterilants. Ideally, transmitters should be sterilized several days before implantation procedures, with sterilization performed in the controlled conditions of a laboratory or veterinary medical facility. Sterile transmitters could then be transported to the surgery site in appropriate packaging, such as sterile sample bags,p which would eliminate the need to use chemical sterilants in field settings. Alternatively, researchers could seek out human medical or dental facilities with the capability to use ethylene oxide or hydrogen peroxide gas to sterilize heat-sensitive equipment.

Regardless of the method used for sterilization, a sterile transmitter can be provided to researchers at a field setting, but transmitters are most likely to be contaminated with pathogens in field conditions when the transmitter is handled and activated (if necessary) prior to or during implantation. Transmitters as well as surgical instruments and suture materials should be considered contaminated at the first contact with nonsterile objects or surfaces (eg, the outside of a fish, a surgery platform, or unsterile water used for anesthesia); therefore, researchers should take appropriate precautions.

Instruments—A fresh set of sterile surgical instruments should be available for each procedure. A small steam autoclave can be used for on-site sterilization of instruments, provided a power source is available. Alternatively, instruments can be sterilized in a laboratory or medical setting and transported to field settings in appropriate packaging. In lieu of heat sterilization, instruments can be sterilized with chemical-based methods.

Researchers have used a combination of sterilization and disinfection in high-volume tagging operations where hundreds of fish from the same cohort are tagged each day. If this is the selected option, researchers should, at a minimum, ensure that instruments are sterilized between cohorts or populations of fish as well as between tagging locations. Within a given cohort of fish, multiple sets of instruments can be rotated through the surgery and disinfection processes. This rotation would involve a series of instrument baths so that the flow of the procedures (anesthetizing, processing [eg, examination, determination of body weight, and obtaining measurements], and tagging) is not interrupted. Instruments should be cleaned before disinfection, and care should be taken to meet the required contact times to ensure efficacy of chemical disinfectants. Disinfected instruments must be rinsed thoroughly with sterile or deionized water or saline (0.9% NaCl) solution prior to use. Instruments that become contaminated with obviously infected tissue (eg, abscess or granuloma) should not be used until they can be sterilized. In addition, because organic material (eg, mucus and scales) may inactivate or decrease the efficacy of some disinfectants, the solutions in the instrument baths should be refreshed or replaced at prescheduled intervals. Researchers should also remain conscious of the need for adequate ventilation when using disinfectants. Broad-spectrum disinfectant solutions, including oxidizing agents, chlorhexidine compounds, iodine compounds, and ethyl alcohol, have been used to disinfect instruments in field settings.

Surgical field—Most implantation surgeries are conducted with the fish out of water and positioned in dorsal recumbency while anesthetic is applied over the gills. The area of the incision can be protected with a surgical drape or plastic wrap. Drapes are particularly useful for prolonged surgeries in which the incision is relatively large and fish remain out of water for more than a few minutes. Plastic drapes are appropriately suited for this purpose because they are waterproof and easy to use in field settings. Researchers should ensure that anesthetic-containing water is applied locally near the head and opercula and does not flow into the incision. When the anesthetic is applied in a bath, the surgical platform should be slanted to ensure the head and gills of the fish are in or near the water and the area of the incision is well above the bath mixture.

Antimicrobial prophylaxis—A variety of disinfectants, antibiotics, and other antimicrobials have been used prophylactically for fish surgery.5,31,50–53 However, none of these agents has been recommended for use in mitigating the effects of implantation procedures. Disinfectants can irritate or damage skin,5,50 and disinfectants and antimicrobials may delay healing and enable or promote growth of opportunistic pathogens such as fungi.53 More tolerable agents, such as iodine compounds, have not been proven effective for promoting healing or preventing secondary infection after surgery.8,54 Investigators in 1 study34 observed that some commonly used compounds, such as hydrogen peroxide and a commercial water conditioner,q when applied in a postoperative bath, actually adversely affected the survival rate of juvenile Chinook salmon.

Finally, there are general concerns that the use of such products may promote antimicrobial resistance. Use of approved products and withdrawal guidelines are additional issues that must be addressed when considering the use of antimicrobials and disinfectants in animals that may potentially enter the human food chain. The US FDA website has a list of drugs currently approved for use in aquaculture.55

Anesthesia

Fish generally have a strong flight response when they are netted and handled. Therefore, application of some form of sedation or general anesthesia prior to handling is considered a best practice for surgical implantation procedures of fish, particularly for small fish and fish in juvenile life stages. In addition to reducing fish movement during the surgical procedure, general anesthesia will minimize handling stress resulting from physical restraint and will increase safety for both the fish and researchers.56

Guidelines of the American Fisheries Society,57 Canadian Council on Animal Care,58 and most US university institutional animal care and use committees currently recommend that whenever possible, general anesthesia be used when fish undergo surgical implantation procedures. However, whether fish cognitively perceive painful stimuli is debatable.59,60 In some situations in which the risks involved with anesthetizing a particular fish or cohort of fish (eg, high potential for overdose or prolonged recovery) are thought to outweigh the benefits, the use of anesthesia is not recommended.56,61 If used, anesthetics should be administered to provide the lightest plane of anesthesia possible to safely conduct the specific procedure. For most procedures, this is defined as the point at which there is loss of equilibrium and failure to respond to all tactile stimuli during the surgical procedure.56

Anesthetic options for fish—Similar to an ideal anesthetic for mammalian surgeries, the ideal anesthetic for fisheries research would be inexpensive and easy to obtain and administer, have a wide margin of safety for fish and researchers, provide smooth and rapid (eg, within a few minutes) anesthetic induction and recovery, provide consistent performance for various species and life stages, perform consistently despite differences in environmental conditions (eg, water conditions), and possess a zero-withdrawal FDA approval for use in food fish. Disposal of the anesthetic should be easy and economical.

Unfortunately, an ideal anesthetic does not exist for all circumstances and fish species. Furthermore, the location (eg, field or laboratory), environmental conditions, and legal constraints under which a study is conducted, rather than what may be ideal for fish performance, often dictate the choice of anesthetic. This is particularly evident for field studies in which researchers release fish directly to a fishery, which could potentially result in subsequent consumption by humans. Currently, only 2 MS-222–based productsr,s have been approved by the US FDA for use in fish intended for human consumption. Both products require a 21-day withdrawal time and are restricted to use in a limited number of fish families (Ictaluridae, Salmonidae, Esocidae, and Percidae), in a laboratory or hatchery, and at water temperatures > 10°C. The MS-222 is available as a white crystalline powder that mixes well with water to form a stock solution. However, when mixed with water of neutral pH, the resulting solution is acidic. Therefore, depending on local water conditions (eg, pH and hardness), anesthetic baths may need to be buffered (eg, sodium bicarbonate) to avoid potential metabolic (eg, increases in BUN and ACTH production) and behavioral (eg, agitated swimming behavior) changes associated with the sulfonic acid moiety of MS-222 solutions.62 Thus, some research protocols for field settings automatically include the addition of sodium bicarbonate to MS-222 anesthetic baths.

Products containing benzocainet and eugenolu have been granted status as INADs by the FDA CVM.63 This means that researchers can apply to use these products as sedatives or anesthetics in fish under certain conditions. Benzocaine is used as a local anesthetic for humans. Similar to MS-222, benzocaine is available to researchers as a white crystalline powder. However, benzocaine is less acidic and less water soluble than is MS-222. Thus, stock solutions of benzocaine typically are prepared with ethanol or acetone. Because it is less water soluble than MS-222, benzocaine is not as potent and therefore generally has a wider margin of safety between the effective and lethal doses. The benzocaine-based productt has the potential to be approved as a zero-withdrawal fish anesthetic. This would mean food fish could be anesthetized with this product and released into a fishery or harvested for human consumption immediately after recovery. Currently, this product can only be used in food fish (in accordance with INAD 11–740) provided there is a withdrawal time of 72 hours.

The eugenol-based productu is also being investigated as a potential zero-withdrawal fish anesthetic. On the basis of early data, FDA approval for use in food fish seems likely. The withdrawal period (in accordance with INAD 11–741) was recently reduced from 72 to 0 hours for field studies in freshwater fish. However, researchers working in hatcheries must still adhere to a withdrawal period of 72 hours.

Eugenol is also the primary ingredient in clove oil, which is commonly used as a tranquilizer or anesthetic in fish. Although clove oil was never granted official approval for use as a fish anesthetic, eugenol is listed in the US Code of Federal Regulations (21 CFR 582.60) as generally recognized as safe if used as a flavoring aid in food. For this reason, many biologists have considered clove oil as a safe product for use in fish that would potentially be consumed by humans. Unfortunately, isoeugenol, which is 1 of 3 active ingredients in clove oil, is carcinogenic in male mice, as determined by the US National Toxicology Program. This determination was made during the period when isoeugenol was being tested as the active ingredient in an early fish anesthetic.v As a result, the FDA CVM officially rescinded authorization for the use of this anesthetic (under INAD 10–541) and also has strictly prohibited the further use of clove oil in food fish.

Some researchers in the fisheries research community consider CO2 and NaHCO3 to be viable fish anesthetics.64–66 However, use of these compounds for this purpose is controversial because of the mechanism of action (ie, cerebral hypoxia). Norway has banned the use of CO2 as an anesthetic for farm-raised fish prior to slaughter. However, because it does not require a withdrawal period, some researchers in the United States have continued to use CO2, primarily because there are no legal alternatives for anesthetizing fish that may be harvested within 2 weeks after the anesthetic procedure for consumption by humans.

Anesthesia of fish with CO2 results in a state of cerebral hypoxia, alters blood gas concentrations, and can lead to metabolic acidosis. Furthermore, it has been observed that fish anesthetized with CO2 have struggled violently as they transition in and out of consciousness. Some researchers have attempted to restrain fish during these stages of excitement to prevent injury to the fish or their eggs. Another drawback of CO2 is that it is difficult to determine an exact dose. Finally, CO2 can lower water pH, depending on the alkalinity. Similar to the situation with MS-222, NaCl and NaHCO3 have been used as buffering agents in conjunction with CO2. In lieu of CO2, some researchers have used a solution of NaHCO3 to anesthetize fish. However, when NaHCO3 is used alone, it is questionable whether enough CO2 is released into solution to cause a surgical plane of anesthesia. Thus, it has been recommended66 that acetic acid be added to NaHCO3 solutions to promote additional release of CO2 and create stronger hypercapnia. However, this further complicates the use of CO2 for field settings.

Injectable anesthetics have not been widely used for telemetry implantation surgeries. No injectable anesthetics are currently approved by the FDA for use in fish. Their availability to researchers is dependent on institutional, state, and federal veterinary regulations. In addition to issues with availability, there are also several potential drawbacks with the use of injectable anesthetics. It can be difficult to achieve the appropriate depth of anesthesia. Without prior sedation, fish must be restrained so that the anesthetic can be injected. Some tranquilizers or anesthetics can result in necrotic lesions at the injection site when too large a volume is administered at a single location.67

Oral administration of anesthetics is precluded by the general practice of withholding food from fish prior to surgery. Furthermore, no orally administered anesthetics are currently approved by the FDA for use in fish.

Nonchemical methods, such as hypothermia (typically induced via immersion in an ice bath) and electroimmobilization, for anesthesia of fish have been investigated. However, nonchemical methods differ from true anesthetics in that they do not induce loss of sensation.68 Thus, use of nonchemical methods for restraining fish during surgical procedures is questionable from a humane perspective.

The main advantage of these methods is a lack of residual chemical effects. For this reason, the use of electroimmobilization has been gaining popularity within the fisheries research community. However, electroimmobilization is not safe for all fish species. Injuries that include vertebral column fractures and death have been associated with the use of this method,69–71 and there appears to be a narrow margin of safety between effective and injurious or lethal doses. Furthermore, an effective dose appears to be closely dependent on water conductivity, temperature,72 and fish size.69 Successful applications, such as electroimmobilization of adult broodstock to facilitate egg collection, have been reported.73–79 However, these studies were conducted primarily on homogeneous populations or groups of fish whereby researchers were able to use very specific settings. Currently, the Animal Welfare Division of the AVMA does not consider electroimmobilization to be an acceptable form of anesthesia or analgesia.80 Therefore, it is clear that more information is needed about this method before it can be recommended for use in transmitter implantation surgeries.

The use of analgesics for postoperative pain management has not been widely practiced by fisheries researchers. Similar to injectable anesthetics, no analgesics are currently approved by the FDA for use in fish, and their availability to researchers is dependent on institutional, state, and federal veterinary regulations. Furthermore, the responses to various analgesics can differ among fish species.81–86 A number of studies suggest that various opioids,60,81,86–89 NSAIDs,85,86,89 and local anesthetics85,89 may provide analgesia in some fish. However, it has been expressed in 1 report83 that pharmacokinetic data will need to be collected on a number of fish taxa before valid guidelines can be developed. Information on the use of analgesics, corresponding doses, and species that have been evaluated to date has been published elsewhere.90

Anesthetic administration and patient monitoring—Administration of anesthetics via bath-immersion is the most common method for inducing anesthesia in fish. Anesthetics can also be manually sprayed with a pump onto the gills to induce anesthesia. Once a surgical anesthetic plane has been achieved, fish typically are removed from the induction bath and transferred to a surgical platform. In the experience of the authors, telemetry implantation surgeries generally can be completed in < 2 minutes (not including the time needed for anesthetic induction and recovery). Surgeries often are completed without the administration of additional anesthetic, but some surgeons prefer to provide additional anesthesia throughout the duration of the procedure. In those cases, maintenance anesthetic solution is perfused over the gills; the anesthetic solution can be provided via gravity feed, with a pump, or by partial immersion of the fish in a second water bath.

Perfusion of the gills with anesthetic solutions throughout the procedure can provide the added benefit of maintaining adequate oxygenation. A system can be modified to allow researchers to switch between water that contains anesthetic and water without anesthetic to maintain the desired plane of anesthesia.

Anesthetic baths or sprays should be prepared with the water to which study fish are acclimated. Anesthetic performance will differ with species, life stage, and fish size as well as with local water conditions. Anesthetic baths may need to be refreshed periodically because their potency may wane. Each fresh mixture should be tested initially on a small number of fish to confirm the efficacy and safety. Finally, adequate supplies of anesthetic stock solution, buffer solutions, and oxygen should be readily available to avoid disruption of the surgery flow or inadvertent stranding of a fish midway through the tagging process.

The maximum amount of time a fish should remain anesthetized will differ depending on the anesthetic, dose, and water conditions as well as the fish species, life stage, and size. However, researchers should strive to keep the overall anesthetic time to a minimum. In some cases, stress associated with induction can be reduced by introducing a sedative dose (a lower concentration of anesthetic) before fish are exposed to the full anesthetic dose.91 A well-oxygenated, dark recovery tank should be available before fish are anesthetized. Fish can be actively resuscitated in the recovery tank (eg, gently pushed forward through the water) if they lose equilibrium (ie, roll over) too quickly or fail to recover within 5 minutes after the end of the surgical procedure.67

Fish progress through several stages of anesthesia, which range from active swimming to total loss of gill movement and cardiac arrest.67,68,92 However, some stages may be transient and not readily observed. Therefore, individual fish may proceed from excitement to death with little or no warning. Visual cues that can be used by researchers to determine the stage of anesthesia include opercular movement (or lack thereof), time required for a fish to lose equilibrium (ie, roll over), and response to stimulation.65,93 Researchers conducting experiments with new anesthetics, species, or life stages will benefit from measuring and recording anesthesia induction and recovery times. Ideally, this would be performed in a laboratory setting before being applied in field settings so that logistical problems can be eliminated or mitigated.

Incision Placement and Closure

Researchers should be familiar with the appropriate surgical tools (eg, needle holders, tissue forceps, and scalpels) needed to perform telemetry implantation surgeries as well as the proper use of those tools. Furthermore, appropriately sized instruments must be chosen for each study (eg, No. 10 scalpel blades for large adult fish and microsurgical blades for small fish and juvenile fish). Obtaining good tissue and suture-handling skills will require practice and repetition.

Incision placement—Incision placement will depend on the body type and size or life stage of fish. In many fusiform fish, incisions are made on or just parallel to the ventral midline. In flatfish, a common incision site is the dorsal aspect directly over the peritoneal cavity.91 In some anguilliform fish, a paramedian incision is made in the lateral body wall ventral to and just cranial to the insertion of the dorsal fin.94,95 Of paramount importance for all fish species is the presence of a suitable cavity to accommodate the selected transmitter because the transmitter can cause organ and tissue (pressure) necrosis when adequate space is not available. Generalized peritoneal inflammation and localized inflammation at the site of the incision have also been observed in fish implanted with acoustic transmitters.30,31,33,34

In general, researchers should strive to make the shortest incision necessary to accommodate insertion of the transmitter so that the number of sutures required and the surgery time can be minimized. Scales are not routinely removed before making an incision because this can result in additional damage to the dermis. However, if required, a minimal number of scales can be removed with surgical forceps and gentle, constant retraction in a caudal direction at a 45° angle.96 Care should be taken to avoid damaging the dermal scale bed. In some species of fish, a needle can be inserted underneath the scales for implantation of a transmitter; the needle should be inserted in a manner that minimizes disruption of the scales.

Suture materials—Materials for incision closure will likely be recognized as foreign by a fish's immune system.97 As such, they may elicit an inflammatory reaction, regardless of the closure technique. Additionally, suture materials are not typically developed for use in aquatic environments or for use with poikilotherms. Therefore, they may not perform in fish as would be expected in mammals. For example, absorbable sutures have been found in fish recaptured in cool, temperate, or cold-water environments up to 6 months after tagging.98 Furthermore, retention of absorbable suture materials in fish beyond that expected in mammals has been observed and described as having negative impacts on fish.34,95,99,100 It has been recommended96 that sutures be removed 2 weeks after surgery to avoid complications associated with suture retention. However, this is not an option when tagged fish are released shortly after recovering from implantation surgery. Therefore, regardless of the classification (eg, absorbable or nonabsorbable), closure materials should be considered as permanent or at least semipermanent when used in fish.

Closure materials can elicit tissue responses similar to those observed for transmitters, including fibrous tissue encapsulation, acute or chronic inflammation, and necrosis attributable to tissue strangulation. Chronic inflammation may develop through a chemical response to the suture material or a physical response to surface roughness, sharp edges, pressure, or implant movement. Researchers should select the smallest suture material that will effectively hold tissues in apposition without tearing. The importance of minimizing the introduction of foreign materials in fish cannot be overemphasized.

No significant differences in fish performance (as measured by survival rate) have been found between incisions closed with synthetic sutures and those closed with organic materials, such as surgical catgut or silk. As would be expected, the least reactive suture materials typically are monofilament synthetics such as polyamide, polyglecaprone, and polydioxanone.99,101–103 As a rule, braided or polyfilament materials such as polyglactin 910 should not be used in aquatic environments because of the potential for wicking of water or foreign material into the muscle layer or peritoneal cavity. However, we found no evidence that this wicking phenomenon has been documented. Researchers that use braided sutures because of their ease of handling should consider changing to pseudomonofilament, coated-cable-type sutures. These sutures have the superior handling properties of a braided suture but without a capillary effect. Stainless steel sutures in the form of staples are not common, but they have been used successfully in fish in which the integument was strong enough to be held in place.104–106 Cyanoacrylates (ie, surgical glues) are not recommended for use in fish because it has been observed that incisions closed with this material have reopened before tissues had time to heal,107,108 and this reopening resulted in transmitter loss. Surgical glue acts as a physical barrier to healing, and surgical glue that is not completely shed can lead to granuloma or fistula formation.69

No single closure material is ideal for all situations. The life stage, habitat, and behavior of study fish must be considered when making the suture selection. For example, adult salmon near the end of their life cycle will likely contribute little energy toward healing; therefore, a nonabsorbable suture material may be preferable to an absorbable material to provide maximum strength over time. Juvenile fish that are actively growing will likely heal quickly; thus, a prudent choice would be the most rapidly absorbed suture material.

Incision closure—Of all the aspects of the surgical process, incision closure techniques are arguably the most studied and described in the literature. However, we believe they remain the least understood. The primary goal of incision closure after telemetry implantation should be to close the body wall in a manner that will promote the most efficient healing. This will be achieved when disruption of tissue is kept to a minimum, tissue is maximally apposed, and materials used for incision closure are benign. Additional goals unique to telemetry research are to ensure transmitter retention and minimize impacts on fish behavior.

In some situations, (eg, high water temperatures), no closure may be the best option. Subyearling Chinook salmon were monitored after surgical implantation with PIT tags by use of protocols (including length of incision and number of sutures) designed for implanting acoustic transmitters in this species and size of fish.34 Investigators in that study34 observed that survival to 28 days after surgery was significantly lower for fish that had retained the 2 original ligatures at 7 days after surgery than for cohorts that had lost 1 or both sutures by day 7. Many sutures were missing from the incisions on day 28, and it appeared in some fish that intact sutures had been forcibly pulled or torn from the incisions as indicated by dermal wounds perpendicular to the initial incision site. Those investigators also noted that retained sutures served as structures for accumulation of organic matter, with some fish developing secondary ulcers associated with the accumulated mass of foreign material.34

In that same study,34 suture retention appeared particularly problematic for Chinook salmon tagged and held at temperatures > 15°C. Similarly, investigators in another study109 found fewer adverse effects in perch (Perca fluvitilis) with no closure of the incision than in those with incisions closed with a single suture. Other researchers have also observed complications related to retained ligatures.30,31,37,95,110 Clearly, there is a need to balance the risk of transmitter loss with complications associated with extended suture presence.

In current fisheries studies, the choice of needle is often based on the needle in the suture package or on personal preference of the researcher, rather than on the basis of the most appropriate needle for the tissue involved. Taper-tip needles may be adequate for juveniles and small fish. However, for most adult fish, needles with cutting tips should be used to minimize tissue trauma.96

A simple interrupted suture pattern is recommended for closure of the body wall following transmitter implantation.5,96,102 The integument of fish is often tightly adhered to underlying muscles, so a single-layer closure is usually sufficient and is also the least traumatic and most easily accomplished technique.5,96 A simple interrupted pattern is uncomplicated and can be quickly inserted; this pattern offers an advantage over continuous suture patterns in that if one ligature becomes untied, the integrity of the entire incision will not be compromised.15 There is also less risk of a trail of dangling suture for the interrupted suture pattern.

A square knot is sufficient to secure each ligature. A surgeon's knot can be used if the incision is under tension or if the suture material has a low coefficient of friction.111,112 To ensure that ligatures remain in place long enough for the incision to heal, additional throws may be used in the knots. However, it is important to mention that additional throws will increase knot security but not knot strength. Bulky knots can create more resistance on the ligatures during fish movements, which can ultimately lead to tearing of tissues and secondary infection. In addition, the more material that is used to create a knot, the greater the surface area available for colonization by microbes, algae, and fungi. Therefore, complex knots with added throws are generally not recommended.

Evaluating the Effects of Handling and Tagging Procedures

Ultimately, fish used in telemetry implantation studies must have behavior that is similar to that of their unimplanted cohorts if results are to be useful. There is little value in observations about the survival, behavior, migration, or spawning activity of tagged fish that differ from those of the untagged population. Despite this obvious constraint, there is little evidence that the performance of tagged fish in field conditions is similar to that of untagged fish. It is hard to provide such evidence for aquatic animals. It is challenging to perform visual observation, and it is often impractical because of the long distances traveled by migrating fish.

There are some solid field-based evaluations of survival after tagging of lake-dwelling fish such as pike, pikeperch (Sander lucioperca), and resident trout (Salmo trutta).38,98,113,114 However, it is far more difficult to study posttagging survival of migratory fish such as salmon and sea-run trout (family Salmonidae), shad (family Clupeidae), sturgeon (family Acipenseridae), lamprey (order Petromyzontiformes), and eel (Anguilla spp). Even resident populations or particular life stages pose challenges for field-based evaluation of long-term survival and transmitter retention. For both acoustic and radio technologies, it is sometimes difficult to distinguish between fatalities, loss or expulsion of the transmitter, and transmission failure.115 These technologies can also be used to track an implanted fish after the fish has been consumed by a predator.

For these reasons, most studies91,115,116 designed to examine the effects of handling and tagging have been conducted in captive or closed settings such as laboratories, hatcheries, and ponds. We anticipate that this will continue. However, some species or life stages are more sensitive to confinement than others, and special consideration must be made in these cases. For example, anadromous fish, such as salmonids, captured during migration and confined in a laboratory or hatchery pond may become stressed. This can result in high mortality rates for both treatment and reference groups.115,116 In particular, juvenile Chinook and Atlantic salmon are known to develop frustrated smolt syndrome if their seaward migration is delayed or they are confined after having adapted physiologically to enter seawater.117

In studies33,34 conducted to examine long-term survival and retention of acoustic transmitters, investigators captured migrating Chinook salmon smolts, maintained them in river water for 2 weeks, and then transferred them to seawater. Two weeks was the time at which the smolts would have entered seawater naturally had the migration not been interrupted by inclusion of the fish in the experiments. Transitioning the fish to seawater sufficiently replicated the naturally occurring conditions, which allowed researchers to maintain the fish for 90 days33 and 120 days.34

Although a laboratory-based study may be the best (or only) option for evaluation of the effects of tagging, researchers should be aware that these types of studies can yield biased results. For example, fish may have higher rates of survival and growth because they are no longer subjected to metabolic stressors in the natural environment, such as finding food, swimming against currents, avoiding predators, and migrating long distances. Thus, laboratory conditions may underestimate the effects of tagging. In studies33,34 conducted to evaluate survival differences between juvenile salmon injected with PIT tags and those surgically implanted with both PIT tags and acoustic transmitters, investigators found that performance and survival metrics within each treatment group were consistently lower for fish released into a river immediately after tagging than for their cohorts that were transported to a laboratory after tagging.

On the other hand, captive fish in a controlled environment may be subjected to stress from overcrowding, delayed migration, or poor water quality. These stressors can lead to delayed growth, secondary infections of the incision, an increase in the risk of developing disease, or even death. Effects attributable to maintaining the fish in controlled conditions may entirely overshadow the effects attributable to tagging.91 Effects attributable to tagging procedures have been reviewed elsewhere.9–11

To avoid these biases and improve telemetry implantation methods, additional field-based evaluations of tagging effects are needed. Researchers should explore the use of nonlethal methods (eg, radiography and ultrasonography) for determining placement and retention of transmitters. In addition, more studies are needed to identify the etiology behind adverse tagging effects that have already been reported. Toward this goal, we encourage researchers to incorporate diagnostic methods, such as necropsy and histologic examination, into study designs. Thoughtfully designed and carefully executed studies will provide reliable data on tagging effects as well as opportunities for collaboration between field-based researchers and veterinary diagnosticians.

Fish survival and transmitter retention—Fish survival and transmitter retention (often evaluated together) are important to any evaluation of a study that involves tagging procedures. Holding fish for 1 or 2 days after tagging will allow evaluation of short-term rates for survival and transmitter retention. Clearly, the handling or tagging method cannot be considered appropriate if numerous tagged fish die. On the other hand, high rates for short-term survival and transmitter retention do not ensure that handling or tagging procedures did not have effects. Sublethal effects are likely, and there can be surgery-related fatalities or transmitter loss days or even weeks after fish are tagged.33,34,115

Transmitter loss can occur passively through the incision or actively through expulsion (Figure 2). Expulsion can take place at the incision site long after the original incision has healed. In some cases, up to 70% of study fish have expelled the transmitters.110 The propensity to actively expel transmitters appears to be a species-specific phenomenon; catfish in particular are known to have extremely low transmitter retention rates. However, expulsion may also be related to water temperature, physical characteristics of the transmitter (including the material used to coat the transmitter), or the ratio of the size of the transmitter to the size of the fish. Transmitter loss may not affect survival rate or health of fish. However, when the rate of transmitter loss is unknown, interpretation of data may be confounded because it may be incorrectly assumed that the fish have died.

Figure 2—
Figure 2—

Site of loss (expulsion) of a transmitter (circle) that was (A) and was not (B) associated with the incision used for transmitter insertion.

Citation: American Journal of Veterinary Research 75, 4; 10.2460/ajvr.75.4.402

Rates of fish survival and transmitter loss can be compared among transmitter types or related species to assess effects of tagging.118–120 In particular, additional research is needed to identify the ideal shape, size, and coating material for transmitters. Factors such as incision location, suture pattern, and closure material need to be identified and characterized to promote maximum healing rates with minimal long-term effects (eg, suture retention or secondary infections). One specific factor is suture materials with predictable rates of decay that are suitable and approved for use in fish.

Growth and performance—Depending on the size of the transmitter relative to the size of the body cavity, a transmitter may prevent a tagged fish from obtaining adequate nourishment.98 Therefore, assuming the survival rate for tagged fish is the same as it is for untagged fish, a subsequent logical step would be to evaluate growth and swimming performance (standard measures of well-being of fish in fisheries). However, similar to studies on long-term survival, most studies on growth have been performed in laboratories. Although the results of those studies may be valid for sedentary or nonmigratory species, such studies are often plagued by the problems associated with migratory species that are held in captivity.91 Hence, the best results are from field-based experiments whereby tagged fish are released into their natural environment and recaptured after a prolonged period.38,98,121 Results from most of these studies have indicated that there is an initial negative effect of tagging on growth followed by a compensatory period or that there have been no effects attributable to tagging.

The physical presence of a transmitter has been hypothesized to affect buoyancy compensation by preventing the airbladder from fully expanding.122 The ability to compensate can directly affect swimming performance and related activities, such as avoiding predators and being involved in typical social interactions (ie, schooling, dominance, and courtship). These factors have been evaluated in only a few species of fish,120,123 and the results have not been consistent.

In some studies,117,124–126 predators consumed large numbers of tagged fish after the fish were released. However, results from studies98,117,127 designed to address predation indicate that tagged fish are not necessarily easier prey than are untagged fish. Studies128,129 focused on the effect of handling and tagging on a fish's position in the social hierarchy have also yielded inconsistent results. However, experiments of this nature are typically conducted in artificial environments, so their results may not be representative of studies conducted in field conditions.

Physiology and reproduction—Physiologic processes and reproductive performance might be compromised for reasons related to the surgical tagging process. However, physiologic and reproductive effects attributable to handling and tagging have been examined in only a few fish species. Physiologic evaluations have been limited to measuring cortisol concentrations or the stress response,123,130 basic physiologic indicators such as the Hct and leukocrit,131,132 and the ability of fish to heal.95,133

Surgical implantation of a transmitter into the coelom can influence development and release of gametes. In studies that rely on surgically implanted fish to evaluate spawning behavior, it would be particularly important to know how the implantation method may affect reproductive physiology and behavior. However, few studies have been conducted to evaluate reproductive performance as an indicator of tagging effects. In 1 study,134 investigators detected an adverse effect of tagging on egg retention in steelhead trout. However, investigators in 2 other studies observed no reproductive effects in perch135 or Pacific lamprey (Entosphenus tridentatus).123

In the future, advances in technology should provide researchers with additional options for evaluating the effects of handling and tagging. In particular, the use of 3-D real-time telemetry appears promising. In 1 lake study,136 researchers performed real-time monitoring of both predators and prey by use of a high-resolution acoustic telemetry system. This system provided data for analyses of survival rates, general behavior, and predation.136 Such technologies would allow detailed field-based tests of various tagging methods and effects of tagging in general.

Currently available technologies can be used in a more creative manner to study the effects of tagging. For example, in a hybrid field-laboratory study, researchers evaluated the survival rate of fish in a field setting and also evaluated transmitter retention and incision healing in a subsample of fish maintained in a laboratory.33,34 Those investigators also collected fish at 2 locations along their migration route and necropsied them to evaluate physiologic effects.

Conclusions and Recommendations

Millions of dollars are spent annually for telemetry technologies to enable researchers to study the performance of fish. Such expenses place researchers under increasing pressure to use justifiable and repeatable tagging methods and provide highly accurate and representative results. Most researchers recognize the value of adhering to veterinary principles and best practices when performing transmitter implantation surgeries, but they have not always received the training needed to understand and implement these practices. Veterinarians have the medical and surgical background necessary to assist with making decisions regarding various aspects of surgery (eg, anesthesia, aseptic technique, and incision closure), but they may lack the knowledge needed to make practical recommendations for improving surgical protocols for fish. We believe that improved telemetry methods and technical training will be achieved through a strong and consistent collaboration between veterinarians and nonveterinarian researchers. We hope that the information and recommendations provided here will serve to promote and enhance communication between the 2 groups.

In the future, we hope to see more opportunities for formal exchanges between nonveterinarian researchers and aquatic animal veterinarian experts through continuing education and outreach programs, such as those conducted at national and regional professional meetings. We also hope to see more research collaborations between the fisheries and veterinary communities to address issues currently plaguing some tagging studies, such as the lack of anesthetic options for use with food fish or the lack of suture materials suitable for use in poikilotherms.

ABBREVIATIONS

CVM

Center for Veterinary Medicine

INAD

Investigational New Animal Drug

PIT

Passive integrated transponder

a.

Aquatic Eco-Systems Inc, Apopka, Fla.

b.

Virkon Aquatic, Western Chemical Inc, Ferndale, Wash.

c.

Chlorhexidine, Durvet Inc, Blue Springs, Mo.

d.

Nolvasan, Fort Dodge Animal Health, New York, NY.

e.

Chlorhex, Vedco Inc, St Joseph, Mo.

f.

Virosan, Boehringer Ingelheim Inc, St Joseph, Mo.

g.

Argentine, Argent Chemical Laboratories, Redmond, Wash.

h.

Betadine, Purdue Products LP, Stamford, Conn.

i.

Povidone, Horse Health Products, Phoenix, Ariz.

j.

Wescodyne, STERIS Corp, St Louis, Mo.

k.

Simple Green, Sunshine Makers Inc, Huntington Beach, Calif.

l.

Roccal-D, Pfizer Animal Health, New York, NY.

m.

Super Germiphene, Germiphene Corp, Brantford, ON, Canada.

n.

Chlorox bleach, Clorox Co, Oakland, Calif.

o.

Oxy-Sept 333, Food and Beverage Division, Ecolab Inc, Saint Paul, Minn.

p.

Whirl-Pak bags, Nasco, Fort Atkinson, Wis.

q.

PolyAqua, Kordon LLC, Hayward, Calif.

r.

Finquel (99.5% tricaine methanesulfonate), Argent Chemical Laboratories Inc, Redmond, Wash.

s.

Tricaine-S (99% tricaine methanesulfonate), Western Chemical, Ferndale, Wash.

t.

Benzoak (20% benzocaine), Frontier Scientific Inc, Logan, Utah.

u.

Aqui-S E (10% eugenol), Aqua Tactics Fish Health/AQATAQ Vaccines, Kirkland, Wash.

v.

Aqui-S (54% isoeugenol), Aqui S New Zealand Ltd, San Luis Obispo, Calif.

References

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