This study was conducted at two locations in California and Alaska, between May 2004 and June 2008. California sea lions (Zalophus californianus) were implanted at The Marine Mammal Center (TMMC), a stranded-animal rehabilitation center in Sausalito, California. The sea lions were released in the Marin Headlands near TMMC and near the Moss Landing Marine Laboratory South of Santa Cruz. Juvenile Steller sea lions (Eumetopias jubatus) were captured near Glacier Island in Prince William Sound, Alaska, and transported to the Alaska Sea Life Center (ASLC) at Seward, located at the Northern end of Resurrection Bay, Alaska. The animals were subsequently released into Resurrection Bay near Seward.

The life history transmitters 8 are 122 mm long, positively buoyant cylinders with an outer diameter of 42 mm, hemispherical ends, and a mass of 115 g. The tags are coated in Epo-Tek^® 302-3M medical grade epoxy (Epoxy Technology, Billerica, MA) certified to USP Class VI standards for biocompatibility with implantation in non-cured and polymerized states. This material prevents adhesion to connective tissue or omentum and, therefore, decreases the likelihood of transmitter ingestion by predators of sea lions. Tags were sterilized in ethylene oxide gas (Anprolene^®, Andersen Products Inc., Haw River, NC). The LHX transmitters were programmed to transmit abdominal temperatures for postoperative monitoring, at hourly intervals and for periods of up to two weeks following implantation, to a nearby handheld UHF receiver, on all animals except CSL6053, CSL6160, TJ22, TJ23 (Additional file 1).

Four stranded California sea lions (CSL) were selected after completion of rehabilitation procedures at TMMC (Additional file 1). One animal (CSL6053) had been treated for a gastrointestinal foreign body (fish hook, see 9), and three had been treated for domoic acid toxicity (see 10). Fifteen wild juvenile Steller sea lions (SSL) were captured for research purposes in Prince William Sound, Alaska (Additional file 1) and transported to a quarantined research facility at the ASLC 11. The first two animals of each species received single transmitter implants; all remaining animals received dual implants. This study was conducted in compliance with all applicable research ethics and animal welfare regulations. The research protocol was approved by the Institutional Animal Care and Use Committees of the ASLC (AUPs #02-015 & #03-007) and TMMC (AUP #02/04) and by the Office of Protected Resources (U. S. National Marine Fisheries Service) under research permits #1034-1685 and #881-1668.

CSL surgeries were performed at TMMC in a hospital building that included a sterile surgery room designed for pinnipeds. All SSL surgeries were performed in a specially designed portable surgical unit (Reiff Manufacturing, Walla Walla, WA) at the ASLC (Figure 1). The containerized surgical unit measures 2.8 × 6.0 m and includes power, heating, running water, ventilation, and surgical equipment. The unit can be transported on a flat bed truck and can be lifted onto the working deck of research vessels. All sea lions were fasted for approximately 12 hours prior to surgery.

California sea lion anesthesia was induced with a mixture of medetomidine (Domitor^®, Pfizer Animal Health, Exton, PA, 70 μg/kg) and zolazepam-tiletamine (Telazol^®, Fort Dodge Animal Health, Fort Dodge, IA, 1 mg/kg) by intramuscular (IM) injection delivered by hand or via dart 12. Steller sea lions were masked with 5% isoflurane (AErrane^®, Fort Dodge Animal Health) in 100% oxygen after they voluntarily entered squeeze cages 13 for induction of anesthesia 14. All animals were intubated using appropriately sized, cuffed endotracheal tubes (10 – 16 mm diameter) and anesthesia maintained with 1% to 3% isoflurane in 100% oxygen delivered via a semi-closed, partially rebreathing circuit. Depth of anesthesia was assessed based on respiratory rate and tidal volume, response to stimuli, palpebral reflex, capillary refill time, and jaw and muscle tone. Respiratory and heart rate, oxygen saturation of hemoglobin (SpO₂), end-tidal carbon dioxide (EtCO₂) and deep rectal or esophageal body temperature were monitored during anesthetic procedures. Intermittent positive pressure ventilation was provided when indicated (i.e. decreased respiratory rate, decreased SpO₂ or increased EtCO₂). Animals that showed continuous irregularities in respiratory parameters were provided with mechanically assisted ventilation through a volume-regulated ventilator (Model 2000, Hallowell EMC, Pittsfield, MA) at a rate of six to ten breaths per minute and a volume of approximately 15 – 20 ml/kg/breath. Hypothermia was prevented with recirculating water or electric heating pads, thermal insulating pads covering the surgical table, warm water bottles placed under the flippers, and/or a Bair Hugger^® Model 505 forced warm air blanket (Arizant, Eden Prairie, MN). At the end of surgical procedures, radiographs were taken for the first three CSL prior to recovery from gas anesthesia to determine exact location of implants inside the peritoneal cavity.

Anesthesia was partially reversed in animals induced with medetomidine-zolazepam-tiletamine using atipamezole (Antisedan^®, Pfizer Animal Health, Exton, PA, 200 μg/kg) IM at the end of the procedure 12. After surgery, isoflurane and oxygen were discontinued, and animals were maintained on room air via endotracheal tube during recovery. Animals were extubated once jaw tone and the swallowing reflex had returned and then were allowed to recover in dry anesthetic pens (CSL) or transport cages (SSL) for one to five hours before being given unrestricted access to seawater in pens with pools.

All sea lions were given the non-steroidal anti-inflammatory agent flunixin meglumine (Banamine^®, Schering-Plough Animal Health Corporation, Summit, New Jersey) at 1 mg/kg IM for postoperative analgesia.

Following implant procedures, all animals were closely monitored at each facility for periods ranging from 41 to 69 days (CSL) and 27 to 55 days (SSL). For a separate study, fecal samples were collected from animals prior to and after procedures to determine corticosteroid levels 15. SSL received periodic health screenings (including the collection of venous blood samples for hematology and clinical chemistry) at scheduled intervals prior to release for other research purposes 11. Radiographs were taken in one California sea lion (CSL6039) prior to release on day 43 post-surgery and in two Steller sea lions (TJ22, TJ23) on days 28 and 14 post-surgery, respectively, to confirm mobility of the implants.

Prior to release, all animals were marked with plastic flipper tags or by using hot iron branding (Additional file 1), and they received external, satellite-linked telemetry transmitters to monitor post-release behavior. These devices (SDR-T16 or SPLASH tags, Wildlife Computers, Inc., Redmond, WA; SRDL tags, SMRU Ltd., St. Andrews, UK) were glued to the dorsal pelage using two-component epoxy 16.

The animals were positioned securely on a stainless steel surgical table in dorsal recumbence. An area approximately 14 cm long × 8 cm wide was clipped on the ventral abdominal midline caudal to the umbilicus and cranial to the pubic symphysis and preputial opening. The surgical site was cleaned and disinfected using routine surgical preparation with povidone iodine scrub, 70% isopropyl alcohol, and povidone iodine solution. A nonporous, sterile, fenestrated drape was placed over the surgical site and secured with towel clamps.

A longitudinal skin incision of approximately 8.5 to 12 cm in length was made along the ventral midline. The linea alba was exposed using sharp-blunt dissection through blubber and subcutaneous fat layers. The linea alba was lifted with forceps to permit penetration of the abdominal wall with a single stab incision using a scalpel blade. The linea alba was then sharp dissected with scissors, avoiding the peritoneum and viscera, to a length sufficient to pass the transmitter body (approximately 8–12 cm). The peritoneum was visualized, incised with scissors, and the opening expanded manually. The transmitters were removed from the sterilization peel pouches and rinsed in sterile saline solution to facilitate insertion. The abdominal wall was grasped on either side of the incision with Allis forceps or hand held retractors and lifted up and laterally while inserting the transmitters through the incision into the ventrocaudal abdominal cavity. Bleeding was controlled when necessary with hemostatic forceps and ligatures of 2-0 PDS II absorbable monofilament suture (Ethicon^®, Inc., Somerville, NJ). The surgical incision was closed in multiple layers using size 1 or 0 PDS II sutures. If possible, the peritoneum was closed in a simple continuous pattern to contain intra-abdominal fat and viscera and ease closure of the body wall. Internal abdominal oblique muscle, linea alba, and external abdominal oblique muscle were incorporated in a simple interrupted or interrupted cruciate pattern to close the body wall. Subcutaneous fat was closed in a simple continuous pattern. Intradermal fat (blubber) was closed in a continuous mattress pattern to appose thick blubber layers. A continuous subcuticular pattern was used to appose skin layers without exposing a knot. Finally, an interrupted cruciate pattern was used on the skin to provide additional support to the skin closure. On the first two procedures in CSL, surgical stainless steel staples were used to close the skin layer. Staples were removed prior to release of the animals. Surgical skin glue was used to secure the skin incision in two SSL.

Mean surgical time was 90 minutes (SD = 8) and mean anesthetic time (from induction to recovery) was 125 minutes (SD = 13) for the 19 sea lions. Figure 2A, B demonstrates the relative size of the LHX implant in the smallest of the four CSL, a female (CSL6018). The single transmitter corresponds to approximately three vertebrae in length. With a mass of only 66 kg, this animal was about half the size of the juvenile Steller sea lions for which the LHX tags were primarily developed. Figure 2C demonstrates the relative size of the implants in a much larger animal, CSL6053 (195 kg), comparable in size to juveniles of the target species (SSL). In this animal, the tags correspond to approximately two vertebrae in length.

During the immediate post-procedure recovery period (1–5 hrs), the animals were mildly to moderately lethargic and rested in sternal recumbence. Once given access to pools, the animals went into the water immediately. During the first 18 hours after surgery, signs of some abdominal discomfort associated with the incision site were noted in all animals. This was characterized by the animal choosing to lie almost exclusively in lateral recumbence or by keeping the abdominal midline elevated by folding the posterior flippers under the abdomen if lying in a sternal position. However, the animals were not observed to lick or bite at the incision site or to rub it with nose or mouth. The animals spent a considerable amount of time in the water but also repeatedly entered and exited the water with both patterns reflecting normal, pre-surgery behavior. After 18 hours, signs of discomfort (abdominal midline elevation) gradually lessened, and none were noticed after 24 – 48 hours. Appetite and food intake appeared normal, and defecation and fecal consistency was normal. Immediate postoperative abdominal temperatures were slightly elevated in some animals (38 to 39.2°C) but otherwise were within normal ranges for otariids (36.0 to 37.9°C) with the exception of three instances observed in two CSL. In these two animals temperatures briefly rose above 39°C as a result of further manipulations related to other treatment exams (two events, CSL6018) or antagonistic interactions with other sea lions (one event, CSL6039). All SSL maintained body mass and exhibited no change in body condition as estimated via deuterium oxide dilution as reported elsewhere 17 through the experimental period.

Figure 3 shows the single LHX tag in CSL6039 immediately after surgery (A), and 43 days after the procedure (B), illustrating the mobility of the transmitters. In TJ22, the single LHX tag had also moved cranial in the animal 28 days after surgery (C). Wound healing progressed very well in all sea lions. Figure 4 shows incisions sites in one SSL after 26 days (A) and one CSL (B) prior to release. All animals showed minimal swelling at the incision site after surgery. A mild clear discharge was noted in two SSL, which may have been associated with skin sutures that were placed too tightly or possibly due to the use of tissue glue preventing drainage. Though surgical steel staples may shorten procedures by up to 15 minutes, they require an additional sedative procedure for removal when compared to absorbable suture on the skin and as a result are not viable for field deployments. No indication of surgical site dehiscence or wound infection was observed as a result of the procedure. No evidence of bacteremia or septicemia was noted in any of the 19 study animals.

All four CSL were deemed releasable based on demonstration of normal behavior, absence of abnormal blood values on CBC and serum chemistry, good body condition, and healed incisions. All 15 SSL were deemed clinically healthy at the time of release based on pre-established release criteria 11. The four CSL were only monitored for relatively short periods (nine to 47 days) following their release (Additional file 1). This was because release for three animals occurred near the time of annual molt, likely leading to the rapid shedding of external transmitters. These animals were not branded, and no consistent re-sight effort exists in the region. Dive behavior of the four implanted CSL was not compared to control animals since no data on post-release behavior of non-implanted, rehabilitated CSL was available.

Post-release tracking of SSL is reported elsewhere 17, but results revealed diving and ranging behavior comparable to non-implanted control animals 16. Animals were tracked following their release for 29 to 182 days (Additional file 1). These animals were released in an area with a considerable re-sight effort through research trips (Alaska Department of Fish and Game, Alaska Sea Life Center) and remote video observation systems 18. Through external tracking devices and re-sight efforts, minimum post-implant survival was confirmed for an average of 73.7 days (SE = 9.0, n = 4 CSL) and 236.6 days (SE = 71.5, n = 15 SSL), respectively (Additional file 1).