“Ultimate responsibility for the ethical and scientific validity of an investigation, and the methods employed therein, must rest with the investigator.” (American Society of Ichthyologists and Herpetologists 2004:3)

The popularity of implanting electronic devices such as transmitters and data loggers into captive and free-ranging animals has increased greatly in the past two decades. The devices have become smaller, more reliable, and more capable (Printz 2004; Wilson and Gifford 2005; Metcalfe et al. 2012). Compared with externally mounted devices, implanted devices are largely invisible to external viewers such as tourists and predators; exist in a physically protected, thermally stable environment in mammals and birds; and greatly reduce drag and risk of entanglement. An implanted animal does not outgrow its device or attachment method as can happen with collars and harnesses, which allows young animals to be more safely equipped. However, compared with mounting external devices, implantation requires greater technical ability to perform the necessary anesthesia, analgesia, and surgery.

More than 83% of publications in the 1990s that used radiotelemetry on animals assumed that there were no adverse effects on the animal (Godfrey and Bryant 2003). It is likely that some studies using implanted electronic devices have not been published due to a high level of unexpected mortality or to aberrant behavior or disappearance of the implanted animals, a phenomenon known as the “file drawer” problem (Rosenthal 1979; Scargle 2000). The near absence of such studies from the published record may be providing a false sense of security that procedures being used are more innocuous than they actually are. Similarly, authors sometimes state that it was unlikely that device implantation was problematic because study animals appeared to behave normally, or authors state that previous investigators used the same technique and saw no problems. Such statements are suppositions if no supporting data are provided or if the animals were equipped because there was no other way to follow their activity. Moreover, such suppositions ignore other adverse effects that affect behavior indirectly, and animals often mask the signs of infection to avoid attracting predators (Wobeser 2006).

Guidance specific to sterilization of electronic devices for implantation is limited in the wildlife record (Burger et al. 1994; Mulcahy 2003). Few biologists have been formally trained in aseptic technique, but most biologists know that electronic devices should be treated in some way to reduce the chance for infection of the host animal by bacteria, viruses, parasites, and fungi. Most biologists (73%) who implant devices into fishes believe aseptic techniques are important (Wagner and Cooke 2005). However, I maintain that many biologists find it difficult to place the concept of asepsis into practice in their work because of confusion about what constitutes aseptic technique, a lack of surgical knowledge and training, the perception of increased costs, or the belief that aseptic surgeries are impractical or unnecessary for their application. Some have even argued that, while compromising surgical techniques in the field might result in complications or mortalities, the money saved would allow for a compensatory increase in sample size (Anderson and Talcott 2006).

In this paper I define aseptic surgical techniques, document the legal and professional guidance for performing aseptic surgeries on wild animals, and present options for sterilizing electronic devices and surgical instruments for field use.

Electronic devices are frequently described in the telemetry literature as being sterilized or disinfected without any distinction between these terms (e.g., Chittenden et al. 2008; Hilmer et al. 2010; Streicher et al. 2011). Sterilization and disinfection differ in the degree that life forms are eliminated (McDonnell 2007; Rutala 2007). Sterilization occurs when all forms of life are destroyed, including spores—the microbial form most resistant to destruction. Disinfection is the reduction or elimination of microbial life forms except for spores. Some disinfectants may eventually kill spores if concentrations are high enough and contact times are extended over many hours. Strictly defined, “asepsis” is the state of being free of microorganisms that might be pathogenic, whereas “sterility” is the complete absence of all forms of life, pathogenic or not. Biologists do not know what microorganisms persist on the disinfected instruments and devices they use; therefore, asepsis is rarely achieved because of improper selection and inadequate concentrations of disinfectant solutions, insufficient contact times, and recontamination of instruments by placing used instruments back into the same disinfectant bath. Details of the modes of action and the uses of specific disinfectants and sterilants, and techniques for disinfection and sterilization are available in many reference books (e.g., Block 2001; McDonnell 2007; Rutala 2007).

Aseptic surgical technique is a combination of procedures that minimize the introduction of potentially infectious microorganisms into the surgical site (contamination) that may grow and invade tissue (infection). Information on aseptic techniques for animal surgery can be found in veterinary surgery textbooks (e.g., Slatter 2003; Fossum et al. 2007; Mann et al. 2011). Aseptic techniques include the use of sterilized surgical instruments; the use of sterile surgical drapes to isolate the surgical field; application of an antiseptic to the surgical site; and the wearing of sterile surgical gloves, gown, cap and mask by the surgeon. All devices to be implanted must be sterile. Aseptic surgical procedures are appropriate for most surgeries on free-living animals, except that sterile surgical gowns are rarely worn, because they are readily contaminated in the field. Any compromises in aseptic technique made during surgical implantation of devices must be approved by an Institutional Animal Care and Use Committee or its equivalent.

Aseptic surgical technique is an important part of implanting telemetry devices, and it begins with preparing sterilized surgical instruments and devices to be implanted (Bonnet et al. 2000; Mulcahy 2003). Even with aseptic surgical techniques, the introduction of microbes into the surgical wound cannot be eliminated (Alban et al. 1999; Mellish et al. 2010). There is a threshold number of microorganisms necessary to produce infection, however, and aseptic procedures are employed to reduce the number of introduced pathogens below that threshold. The presence of foreign matter, such as an implanted electronic device, can lower the threshold number of microorganisms required for infection to occur.

Aseptic techniques must be used for surgeries from which the animals are intended to survive (Table 1). Federal regulations (9 C.F.R. 1A § 2.31(d)ix) promulgated to administer the Animal Welfare Act (7 U.S.C. §§ 2131–2159) state that, “All survival surgery will be performed using aseptic procedures, including surgical gloves, masks, sterilized instruments, and aseptic techniques” and that “Operative procedures conducted at field sites need not be performed in dedicated facilities, but must be performed using aseptic procedures.” For federal agencies, coverage of the Animal Welfare Act is extended to all vertebrates by the Interagency Research Animal Committee Principles (Office of Science and Technology Policy, Interagency Research Animal Committee 1985). The Guide for the Care and Use of Laboratory Animals (National Research Council 2011:118) states, “General principles of aseptic surgery should be followed for all survival surgical procedures.” The Guide to the Care and Use of Experimental Animals (Canadian Council on Animal Care 1993:IX. Standards for Experimental Animal Surgery. D) dictates that, “All species undergoing surgery should receive a similar level of care and attention. Recovery surgeries in all species of animals should be performed using aseptic technique. Instruments should be sterile.” and “Surgery in field conditions should be performed in as clean an environment as possible, with sterile instruments, sterile surgical gloves and aseptic technique.”

Table 1.

Key statements about aseptic techniques in major guidelines produced by professional societies and government agencies in North America. Citations are given when authors other than societies or governmental entities are named and page numbers are listed when the guidelines are paginated; otherwise section numbers are listed.

Guidelines addressing the use of specific taxonomic groups of animals in research have been issued by several professional societies (Table 1). Most acknowledge that aseptic techniques or good-quality surgical procedures, presumably meaning aseptic techniques, should be used. However, some of these documents are unclear regarding the definition of aseptic technique. For example, the guidelines from the American Society of Ichthyologists and Herpetologists (2004:22) state that “aseptic but not necessarily sterile procedures should be employed,” without defining the difference. Most perplexing is the declaration in the guidelines issued jointly by the American Fisheries Society and other organizations (American Fisheries Society, American Institute of Fishery Research Biologists, and American Society of Ichthyologists and Herpetologists 2004: § VII. Laboratory Activities. Surgical Procedures), that “Given the aquatic environment in which fish live, it is impossible to conduct surgical procedures under sterile conditions, even within laboratory settings.” The statement is false because, with very few exceptions, surgeries on fish are conducted out of the water, allowing aseptic techniques to be routinely used (Stoskopf 1993; Wildgoose 2000; Mulcahy 2003; Harms 2005), and it ignores the ongoing use of aseptic surgical techniques to implant electronic devices into marine mammals and birds that share the aquatic environment with fishes.

Besides legal and professional requirements, there are at least three additional and interrelated reasons for sterilizing devices and surgical instruments and using aseptic technique during implantation surgeries. These include 1) assuring the quality and reliability of the data collected, 2) being concerned for animal welfare, and 3) preventing transmission of infectious agents between individual animals and between populations.

From the standpoint of data quality, animals that develop infections but that do not die may be of greater concern than those that quickly die. Dead animals yield no incorrect data, but sublethally affected animals may. A critical principle of telemetry studies is that the attachment of an electronic device should not alter the behavior of the animal being studied (Guthery and Lusk 2004; Wilson and McMahon 2006; McMahon et al. 2011; Vandenabeele et al. 2011a, 2011b). Data gathered concerning the activity and behavior of equipped animals, however, may not be representative if sublethally affected animals do not behave the same as healthy animals (Bradfield et al. 1992). Extrapolations made to the larger, unmarked population using such flawed data could result in serious errors in wildlife population management. A period of censoring data from implanted animals often is used to guard against short-term alterations in activity and behavior resulting from research manipulations, but reasons are rarely presented for selecting the length of the censoring period, let alone data demonstrating that censoring periods are accurate (Esler et al. 2000; Iverson et al. 2006; Dechen Quinn et al. 2012).

The privilege of using animals in research is accompanied by the obligation to minimize their pain and distress. Infection and inflammation in the implanted animal caused by inadequate surgical techniques produce unnecessary pain and distress. Several researchers argue that even fish, the vertebrate animal group whose welfare has been given the least attention until recent years, can feel pain (e.g., Sneddon 2003a, 2003b, 2006, 2009; Braithwaite and Boulcott 2007), whereas others still maintain that fish do not feel pain (Rose 2002, 2007). Until these disputes have been resolved, biologists should extend to fishes and other poikilothermic vertebrates the same considerations given to mammals and birds, including the provision of analgesia.

I found no studies addressing the following scenario: potential transmission of infectious agents from one animal to another associated with implanting electronic devices into a series of animals while reusing inadequately treated surgical instruments. However, there is evidence from other invasive procedures that such investigator-caused transmission of disease agents can occur. For example, the fish pathogen Renibacterium salmoninarum can be transmitted by the repeated use of needles on equipment that injects coded wire tags into the snouts of fishes or by the injection of transponders into fish coeloms (Elliott and Pascho 2001). A previous study on the potential transmission of R. salmoninarum among fish being tagged with coded wires did not detect transmission (Zajac et al. 1988), but these researchers relied solely on detecting visible signs of infection; whereas, Elliott and Pascho (2001) demonstrated infection by histopathology, enzyme-linked immunosorbent assay, and a fluorescent antibody test. This finding demonstrates that studies purporting to test for adverse effects of implanting electronic devices by simply observing the implanted animals or by the use of simple indices may well arrive at erroneous conclusions. Total and differential white blood cell counts are recommended for evaluating transmitter implantation surgeries in small mammals, but may not be applicable to all taxa (Reynolds 1992).

To adequately perform aseptic surgical techniques in the field, either a sufficient number of surgical tools must be purchased to allow a separate, sterilized set to be used on each animal, or the means to sterilize used instruments in the field must be available. Given the availability of inexpensive surgical instruments, it is easier to purchase sufficient surgical sets than to try to clean, package, and sterilize surgical instruments in the field unless the surgeries to be done number in the hundreds. The five instruments (Olsen–Hegar needle holder, Adson tissue forceps, Kelly forceps, operating scissors, and teat cannula) in the set that I use in my transmitter-implantation surgeries can be purchased in economy grade from a single veterinary medical discount supply company for a total of US$16.05 (4 September 2012) per set. The estimate for the direct costs (logistics, transmitters, and data collection fees) to implant a large number of satellite transmitters into spectacled eiders Somateria fischeri at remote sites in Alaska was US$8,000–10,000/bird (M.G. Sexson, U.S. Geological Survey Alaska Science Center, personal communication). The cost of the loss of a single animal to infection would be vastly more expensive than the cost to purchase sufficient surgical tool sets to allow a separate set of sterilized instruments to be used for each bird. Given that these instruments are made of metal, they can be reused for years, which would amortize their costs. Long-term reuse and the amortization of the costs of instrument sets are factors sometimes ignored by those arguing that aseptic technique is too expensive for field use (e.g., Chomyshyn et al. 2011).

Several methods are available for sterilizing surgical instruments and electronic devices (Table 2). All have advantages and disadvantages. Regardless, surgical instruments and electronic devices sterilized by any method do not remain sterile forever, even with barrier packaging. The expiration date of sterility varies with the type of barrier packaging used and the conditions of storage.

Table 2.

Options that can be used to sterilize surgical instruments and electronic devices for implantation into animals. Instruments must be cleaned of all organic matter prior to sterilization. Exact details regarding the use and operation of these techniques are available from the manufacturer's literature.

If a liquid is needed for on-site use, glutaraldehyde and peracetic acid are the only liquids available in commercial preparations that are readily available and capable of sterilization (Table 2). These solutions require a contact time measured in hours and are highly toxic, so they must be thoroughly rinsed from instruments and devices with sterile saline or water in an aseptic manner. This limits their use for sterilizing electronic devices to just prior to implantation surgery. The concentration of the solution and duration of exposure are critical parameters that determine effective sterilization. The solutions are only effective for a certain number of days after preparation (usually a month or less), and they represent a problem for disposal in the field. Any device or attachment that can absorb liquids, such as an antenna collar or anchoring mesh, should not be sterilized using liquids because these solutions cannot be effectively rinsed free of the toxic chemicals.

Steam autoclaving is the most commonly used technique for general sterilization because it is nontoxic, inexpensive, and readily available, although it cannot be used with electronic devices. Autoclaving is the preferred method for sterilization of steel surgical instruments because they are not damaged by heat, moisture, or pressure (Table 2). Objects to be sterilized typically are placed in barrier packaging (double-wrapped in cloth or in special paper sterilization wraps or envelopes that permit ready passage of steam) that prevents recontamination after sterilization is complete. Two layers of packaging are usually used, assuring the maintenance of sterility especially during transportation to the field. Autoclaves usually operate at temperatures of 121°C and require 15–30 min of operation after pressure in the chamber reaches 105 kPa (15 psi; Table 2). Typically, autoclaving is accomplished in a permanently installed machine because most autoclaves require electricity and a source of water. Portable autoclaves, such as those used in small medical or dental offices, can be used for small loads, and require only a source of electricity; water can be manually added to the chamber before the process begins. Field-portable autoclaves and large culinary pressure cookers have been used for onsite sterilization in the field and are marketed through large biological supply houses. Field autoclaves have a larger capacity than sterilizers designed for use in medical offices. Autoclaving is never used to sterilize electronic devices because of the potential for thermal damage to electronic components, batteries, and heat-sensitive coatings such as waxes and plastics.

Chemical gases are the best options for sterilizing delicate electronic devices intended for implantation of animals. The low-temperature ethylene oxide sterilizer is most commonly used (Table 2), and is increasingly popular (e.g., Beaumont et al. 2002; Horning et al. 2008; Green et al. 2009; Hyslop 2009; Hinton and Chamberlain 2010; Mulcahy et al. 2011). Ethylene oxide is a gas at room temperature. Because it is flammable, explosive, and a serious health hazard to humans, it requires the use of specialized but comparatively inexpensive equipment. The manufacturer's instructions for use of their system must be closely followed to assure safety and efficacy (e.g., Andersen Products 2011).

Small ethylene oxide sterilizers operate using the chemical supplied in liquid form in a single-use glass ampoule sealed in a gas-permeable plastic envelope. The ampoule-containing envelope is placed into a specific-purpose plastic bag with the items to be sterilized (in gas-permeable envelopes or wraps) and a source of humidity (water-soaked gauze sponges typically are used). Sterilization is dependent on the presence of an adequate level of humidity (40% is ideal). The bag is placed into the sterilization chamber and a vacuum tube is inserted into the mouth of the bag, which is then tied tightly. The ampoule is then broken, the liquid vaporizes, the door is immediately closed, and the automated processing is initiated. Following the complete cycle, the spent ampoule may be disposed of in the domestic trash.

Except for items made of glass and metal, sterilized objects absorb ethylene oxide and must be allowed to outgas for at least 24 h in a fume hood or in an open, well-ventilated area before use (Andersen Products 2011). Objects to be sterilized must be wrapped in cloth or placed into sterilizing envelopes. The envelopes are marked with spots of chemical indicators that turn color when exposed to ethylene oxide gas. Objects in the bag must be loosely packed to permit penetration by the gas. Additionally, a single-use dosimeter is placed into the bag with the objects to be sterilized.

Hydrogen peroxide gas plasma is an effective way to sterilize electronic devices (Table 2). Like ethylene oxide sterilization, low-temperature gas plasma requires a dedicated piece of equipment and proper packaging and monitoring (Sterrad Systems; Advanced Surgical Products, Irvine, CA; www.aspjj.com/us/products/sterrad-sterilization). Unlike ethylene oxide gas sterilization, gas plasma is very rapid and leaves no toxic residue, but the equipment is expensive and its availability is limited because it is a newer technology. Hydrogen peroxide gas plasma has been used to successfully sterilize several different types of transmitters used in wild animals (Wild et al. 1998; Ferrell et al. 2005; Crawshaw et al. 2007; Gaydos et al. 2011; Lentini et al. 2011).

Other processes (chlorine dioxide, peracetic acid) for chemical sterilization exist (Table 2), but have not been used for field projects on wild animals. These methods may be suitable, but should be used with caution and thoroughly tested for unexpected complications before being introduced for routine use.

The use of aseptic surgical techniques and sterilized surgical instruments and electronic devices for implantation into wild animals is mandated by laws and in guidelines from most professional societies. Nevertheless, biologists continue to use procedures and materials that do not conform to the laws and guidelines, or to modern standards of care for surgery. I believe that is because surgical procedures are typically passed down from one generation of graduate students to another without outside input. Also, most biologists assume that if their equipped animals live, then they are normal and there was no effect of instrumentation. This ignores the potential for sublethal effects, including infection and inflammation that may alter the validity of the data being collected. The first step in reducing the potential for sublethal effects should be to use the best possible surgical techniques.

In terms of practicality and availability, the best practice is to sterilize instruments by autoclaving and electronic devices by either ethylene oxide gas or gas plasma. Institutions without these capacities can obtain them on a fee-for-service basis from large veterinary hospitals, human hospitals, or universities. A separate set of sterile instruments should be used on each animal, and ideally, sufficient surgical instruments should be available to permit sterilization of instrument packs before leaving for the field. However, field-portable autoclaves can be used to sterilize used instruments after they are cleaned. Electronic devices cannot be autoclaved and should be sterilized before leaving for the field. Chemical sterilizing solutions such as glutaraldehyde can be effectively used in the field to sterilize electronic devices, but they require long contact times, devices must be thoroughly rinsed of solution before implantation, and sterilization must be done immediately before surgery.

I thank M. Murray and C. Harms for reviewing the manuscript. Marcus Peterson made extensive modifications to the manuscript. I thank all anonymous reviewers and the Subject Editor for their comments.

Any use of trade, product, or firm names is for descriptive purposes only and does not imply endorsement by the U.S. Government.

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Author notes

Mulcahy DM. 2013. Legal, ethical, and procedural bases for the use of aseptic techniques to implant electronic devices. Journal of Fish and Wildlife Management 4(1):211-219; e1944-687X. doi: 10.3996/092012-JFWM-080

The findings and conclusions in this article are those of the author(s) and do not necessarily represent the views of the U.S. Fish and Wildlife Service.