PATHOGEN EXPOSURE AND BLOOD CHEMISTRY IN THE WASHINGTON, USA POPULATION OF NORTHERN SEA OTTERS (ENHYDRA LUTRIS KENYONI)

Sea otters (Enhydra lutris) were extirpated from Washington State, United States, in the 18th and 19th centuries primarily due to intensive pelt harvesting (Jameson et al., 1982). Successive translocation of 59 northern sea otters (Enhydra lutris kenyoni) from Amchitka Island, Alaska in 1969 and 1970 successfully reestablished a population off the west coast of the Olympic Peninsula (Jameson et al., 1982). Washington sea otters are protected under the Marine Mammal Protection Act, and the population's small size, geographic isolation, and vulnerability to oil spills have resulted in State Endangered status (Washington Administrative Code 232-12-014; Lance et al., 2004).

To determine overall health of the Washington sea otter population, blood chemistry and pathogen exposure were determined for 30 wild sea otters captured in 2011 as part of research by the US Geological Survey (Bodkin, 2010). Every effort was made to use the same laboratories and methods that were used in a similar survey in 2001–2002 by the US Fish and Wildlife Service, the Washington Department of Fish and Wildlife, and National Oceanic and Atmospheric Administration's Olympic Coast National Marine Sanctuary (Brancato et al,. 2009). We compared results in a cross-sectional study to assess changes in clinical pathology values (serum chemistries), exposure to pathogens, and overall health of the population over the last decade. Selection of pathogens to examine was based on previous documentation in this and other sea otter populations and on new and emerging diseases in other marine mammals.

Thirty free-ranging sea otters were captured in August 2011 in the Olympic Coast National Marine Sanctuary, Washington (47°58′N, 124°41′W) as part of a coastal ecosystem study (Bodkin, 2010). Comparative data were obtained from animals captured in 2001–2002 in the same location for a study of chemical contaminant and pathogen exposure (Brancato et al., 2009). Capture and sampling methods were similar in 2011 and 2001–2002 except where noted. Individual animals or mother-pup pairs were captured in hand-held Wilson traps (Brancato et al., 2009). Adult animals were immobilized with an intramuscular injection of fentanyl and midazolam (valium was used in 2001–2002) by a licensed veterinarian experienced with sea otters. Physical examinations of adults included weighing, measuring, and determination of sex, reproductive status, and physical abnormalities. Body condition was calculated as ratio of mass (kg) to body length (straight-line distance between tips of nose and tail). A premolar was extracted from adult sea otters and sent to Matson's Laboratory (Milltown, Montana, USA) for age determination by cementum annuli analysis (Bodkin et al., 1997). Each adult was marked with a unique combination of colored plastic tags placed in the interdigital webbing of the hind flippers (Ames et al., 1983). Anesthesia was reversed with an intramuscular injection of naltrexone. Each otter was fully awake when released near capture sites. Mother-pup pairs were released together (Brancato et al., 2009).

Blood was collected via jugular venipuncture and allowed to clot. Serum clot tubes were centrifuged within 6 hr of collection and aliquoted into plastic screw-top vials. In 2011 the nares of sea otters were swabbed with a sterile polyester swab placed in a 4-mL cryovial of viral transport medium (Docherty and Slota, 1988) to detect current morbilliviral infections. Serum samples for biochemical analyses were frozen at −20 C until shipment to Phoenix Central Laboratory (Everett, Washington, USA). All other serum aliquots and nasal swabs were frozen in liquid nitrogen (−150 C), shipped to the US Geological Survey's National Wildlife Health Center (NWHC; Madison, Wisconsin, USA), and stored at −80 C until testing or shipment to various laboratories.

Sea otters in 2011 were palpated and urine was collected from full bladders via cystocentesis. Leptospira culture medium (Zuerner, 2005) was added to a urine aliquot within 6 hr of collection and incubated up to 1 wk at 29 C before submission to testing laboratory. Urine samples for domoic acid (DA) and saxitoxin (STX) testing were stored at −20 C until tested.

Laboratories and methods used to determine serum chemistry values and pathogen exposure were similar between 2011 and 2001–2002 samples except where noted. Biochemistry parameters in serum were analyzed by standard methods (Table 1). Pathogens and laboratories used to determine pathogen exposure are listed in Table 2. Values for positive results were provided by the testing laboratory.

Several serum chemistry parameters were not normally distributed; therefore, analysis of variance on ranks was used to explore differences in serum chemistry parameter between years (2001–2002 vs. 2011), sexes, and body condition. In addition to exploring changes in pathogen exposure in the Washington population between 2001–2002 and 2011, we examined the effect of age, sex, and body condition on the probability of disease occurrence with multiple logistic regression. All analyses were conducted in R (R Development Core Team, 2011).

Thirty free-ranging sea otters were captured off the coast of Washington state in each study (2011 and 2001–2002). All captured animals were clinically healthy at capture. Age was similar (Table 3) and body condition was not statistically different between study years (F_1,56 = 1.58, P = 0.21). Blood chemistry values and pathogen exposure of the two pregnant sea otters captured in 2011 were within the range for other otters captured that year. Low sample size prevented statistical analysis of the effect of pregnancy on response variables.

Sea otters sampled in 2011 had higher chloride (F_1,56 = 11.8, P = 0.001), total carbon dioxide (F_1,56 = 60.1, P<0.001) and sodium:potassium ratios (F_1,56 = 4.1, P = 0.05), and lower calcium (F_1,56 = 6.4, P<0.001), cholesterol (F_1,56 = 24.3, P<0.001), creatinine (F_1,56 = 58.3, P<0.001), potassium (F_1,5  = 14.6, P<0.001), and bilirubin (F_1,56 = 5.4, P<0.001) compared to animals sampled in 2001–2002 (Table 1). Female sea otters had higher cholesterol (F_1,56 = 17.2, P<0.001) and lower creatinine (F_1,56 = 16.3, P<0.001) compared to male sea otters (Table 1). There was also significant interaction between sex and year for calcium (F_1,5  = 4.5, P = 0.04) and globulins (F_1,56 = 4.9, P = 0.03).

Results of pathogen screening are summarized in Table 2. Exposure to Toxoplasma gondii, Brucella spp., influenza A virus, and Morbillivirus was detected in sea otter serum in 2011. There were significant differences in pathogen exposure between study years for T. gondii. Antibody prevalence to T. gondii was 97% (29/30) in 2011 compared to 60% (18/30) in 2001–2002. Study year alone was the best predictor of T. gondii status in Washington sea otters (next best model >3 ΔAIC units). Sea otters in 2011 were 19 times (95% CI = 3.4–367) more likely to be positive for T. gondii exposure than otters in 2001–2002.

Morbillivirus was not detected via PCR performed on nasal swabs. There were significant differences in Morbillivirus exposure between study years. Antibodies to Morbillivirus were detected in 10% (3/30) of otters sampled in 2011 while 80% (24/30) of animals sampled in 2001–2002 were positive. All animals sampled in 2001–2002 with positive antibody titers (n = 24) had higher titers, in most animals by two or more dilutions, against canine distemper virus (CDV) and phocine distemper virus (PDV) than against cetacean morbilliviruses (dolphin morbillivirus and porpoise morbillivirus). In 75% of positive animals (18/24) from 2001–2002, titers were higher to CDV than to PDV, but typically only by one dilution. Animals sampled in 2011 were tested only for exposure to CDV and PDV, and two of the three otters that were antibody positive had 2-fold higher titers to CDV than to PDV. The model that included all predictors (year+age+body condition+sex) was the best at predicting Morbillivirus exposure in sea otters (next best model >10 ΔAIC units; misclassification rate 10%); however, year and age were the only statistically significant predictors within the model. Odds of Morbillivirus exposure were 500 times higher (95% CI = 43.7–26,410) for otters sampled in 2001–2002 compared to 2011. Likelihood of exposure increased with age, 1.5 times (95% CI = 1.1–2.4), for every increased year.

All 2011 serum samples were negative for antibodies to Sarcocystis neurona by indirect fluorescent antibody test (IFAT). Antibodies to S. neurona were detected using the Sarcocystis direct agglutination test (SAT) in 2001–2002 in 29% (4/14) of sampled animals, but differences in testing methods between study periods did not allow direct comparison. Antibodies to Brucella spp. were detected only in 2011 in 10% (3/30) of sampled animals. Influenza A virus exposure was found in 23% of the animals sampled in 2011; testing had not occurred in 2001–2002 for comparison. No animals had antibodies against Leptospira, and urine culture results for Leptospira were negative for all six animals examined in 2011, while one animal had a positive Leptospira interrogans serovar gryppotyphosa titer (1∶100) in 2001–2002. No detectable antibodies to caliciviruses or Coxiella burnetii were found in any sea otters sampled.

All but one animal tested for DA in 2011 (5/6 urine samples, 1/1 fecal sample) had detectable concentrations with the enzyme-linked immunosorbent assay (ELISA). Two animals had substantial amounts of DA (17.8 ng/g and 26.3 ng/g) but not as high as those associated with severe signs and death in other marine mammals (Scholin et al., 2000). No DA was detected in urine samples collected opportunistically in 2001–2002 from three males with the receptor binding assay (RBA); however, the minimum detection limit for the RBA in sample material is much higher (>50 ng/g) than with the ELISA (>4.0 ng/g). Saxitoxin was detected (detection limit in sample material >3.0 ng/g) in four of seven samples (n = 6 urine, n = 1 fecal). Two animals had substantial STX concentrations (20.6 ng/g and 26.3 ng/g); however, lethal levels of STX for marine mammals have not been established (Fire and Van Dolah, 2012) and both animals were apparently healthy at the time of sampling.

Results of the serum chemistry evaluations did not reveal significant health issues in the Washington sea otter population. The majority of serum values were within or near published reference intervals for sea otters and similar mustelid and marine mammal species, so most differences between study years and sexes were not considered clinically significant. The only potentially clinically significant differences were in cholesterol and total carbon dioxide parameters. Cholesterol levels were significantly higher in females. Two female cholesterol values were considered potentially clinically significant due to their outlier status (235 and 303 mg/dL) but were still within the range reported for other sea otter populations (Hanni et al., 2003). Two otters sampled in 2011 had total carbon dioxide levels (an indirect estimate of bicarbonate) in the mid-30s (35 mEq/L and 34 mEq/L), which could indicate primary metabolic alkalosis (possible upper gastrointestinal disease) or compensation for a respiratory acidosis (possible respiratory disease). However, blood gas and pH levels are altered during dives (Ponganis, 2011), so high levels of carbon dioxide in these animals could have been affected by differences in diving conditions.

Infectious diseases have been recognized as an important mortality factor for California's southern sea otter population (Enhydra lutris nereis; Thomas and Cole, 1996; Kreuder et al., 2003). Although the Washington sea otter population has continued to grow at a finite rate of 7.9% since 1989 when formal censuses began (Jameson and Jeffries, 2011), recovery plans emphasize the importance of monitoring for pathogens that could potentially threaten this population (Lance et al., 2004). The pathogens evaluated in this study were selected because they were correlated with die-offs and reproductive failure in other populations of sea otters or sympatric marine mammals. Exposure to new pathogens and significant changes in pathogen exposure over the last 10 yr in this population were found.

Clinical toxoplasmosis has been documented in several marine mammal species, including sea otters (Dubey et al., 2003b). Exposure to T. gondii appears to be widespread among marine mammals being documented in sea otters, seals (Phoca erignathus), sea lions (Zalophus californianus), walruses (Odobenus rosmarus), and dolphins (Tursiops truncatus; Dubey et al., 2003b). Washington sea otters from 2011 and 2001–2002 were tested for T. gondii exposure at the same lab with the same agglutination test (Table 2) and exposure rates were high, with exposure 19 times more likely in 2011 animals than those sampled 10 yr prior. Although S. neurona is more frequently the cause of mortality in protozoal meningoencephalitis of sea otters (Thomas et al., 2007), T. gondii is considered a major contributor to the slow population recovery for the California sea otter population (Conrad et al., 2005). Aberrant behaviors resulting from T. gondii infections in southern sea otters may make them more vulnerable to other factors such as shark attacks (Kreuder et al., 2003). Therefore, the 19-fold increase (95% CI = 3.4–367) in exposure to T. gondii in the Washington sea otter population over the last 10 yr suggests that monitoring for this disease should continue.

Morbilliviruses have been known to cause mortality in pinnipeds and cetaceans in the northern Atlantic Ocean and Mediterranean Sea since the 1980s (Di Guardo et al., 2005). Phocine distemper virus has been particularly well documented as the cause of repeated large pinniped mortality events in the North Atlantic (Hall et al., 2006) but was not found in the Pacific region until PDV nucleic acid was detected in nasal swabs of northern sea otters in south-central Alaska (Goldstein et al., 2009). Molecular similarities between the North Atlantic PDV and the sea otter PDV suggest that the virus may have spread across the polar ice cap during thaws from recent Arctic warming (Goldstein et al., 2009). Alaskan sea otters around the Kodiak Archipelago had a 41% prevalence of antibodies to PDV (Goldstein et al., 2011). In Washington sea otters the prevalence of morbillivirus-neutralizing antibodies was significantly higher in 2001–2002 (80%) than in 2011 (10%). Sera collected from Washington sea otters during 1992–1997 (n = 14) were negative for antibodies to CDV or PDV (Ham-Lamme et al., 1999). These data suggest an incursion of a morbillivirus in this population between 1997 and 2001, but by 2011 the virus was infrequently encountered. High morbillivirus titers may indicate animals survived a recent viral outbreak (Ohashi et al., 2001), and anti-morbillivirus titers in otters in 2001–2002 were generally much higher than those in 2011. In July–August 2000 a higher than usual number of carcasses (n = 21) were detected at the southern end of the Washington sea otter range, but most of the carcasses were decomposed and the cause of the event could not be determined (NWHC, unpubl. necropsy data).

Titer differentials indicate that Washington sea otters were exposed to CDV or PDV rather than to cetacean morbilliviruses. The source of the morbillivirus in Washington sea otters is unknown. Although there is no interchange between the Alaskan and Washington otter populations, PDV could have been transported south by other marine mammals. However, a 2-fold titer difference between CDV and PDV in some otters suggests the morbillivirus is CDV. When sea otters haul out on land they could come in contact with CDV-infected terrestrial carnivores. This transmission pathway was suggested for other marine mammals (Kennedy et al., 2000). Fatal CDV infections were documented in river otters in British Columbia (Mos et al., 2003), but CDV exposure was not detected during a serologic survey of marine-foraging river otters from Washington (Gaydos et al., 2007).

Fatal infections with S. neurona have been documented in California and Washington sea otters (Thomas et al., 2007) and Pacific harbor seals (Phoca vitulina richardsi; Miller et al., 2001). Asymptomatic infections were documented in California sea lions (Zalophus californianus; Gibson et al., 2011) but not in sea otters to date. In this study, exposure to S. neurona in Washington sea otters was 29% in 2001–2002 and no otters were positive in 2011. However, direct comparison between study years was prohibited by differences in testing methods. Although the sensitivity and specificity of the SAT (2001–2002 test) are 100% and 90% (Lindsay and Dubey, 2001), respectively, in mice, this test has not been validated in sea otters in which other Sarcocystis spp. occur (Dubey et al., 2003a). The IFAT (2011 test) titer cutoff values have been established to correlate with S. neurona infection and active disease in sea otters (Miller, pers. comm.; Table 2).

Although we did not detect S. neurona exposure in the 2011 live-animal samples, the prevalence of S. neurona fatalities in Washington sea otters in 2011 was 53% (eight of 15; NHWC, unpubl. necropsy data), indicating that S. neurona continues to be a major cause of mortality in this population. Discrepancies between S. neurona serology and diagnostic findings for dead otters from Washington may be due to a short prodromal period, postulated to be around 1 mo (Thomas et al., 2007). Severe meningoencephalitis accompanying S. neurona infections suggests relatively poor adaptation between S. neurona and sea otters (Thomas et al., 2007). If most S. neurona exposures result in acute fatal infections, detection of S. neurona antibodies in apparently healthy otters would be unlikely. Anecdotally, sea otter 44659-001 sampled in August 2011 had a high antibody titer for T. gondii (≥1∶200 via modified agglutination test [MAT]) but was negative for S. neurona (via IFAT). This otter was found dead in May 2012 and diagnosed with severe S. neurona meningoencephalitis (NWHC case 23958) with no evidence of active T. gondii infection (NWHC, unpubl. data).

Brucella spp. are well-known pathogens of terrestrial mammals and humans (Thorne, 2001; Franco et al., 2007). More recently, Brucella infections have also been correlated with reproductive failure and encephalitis in marine mammals (Nymo et al., 2011). Anti-Brucella antibodies were detected previously in sea otters from Russia, Alaska, and California (Hanni et al., 2003; Goldstein et al., 2011) and were detected in three Washington sea otters in 2011. All three were positive using buffered acidified plate agglutination and fluorescent polarization assay and only one was positive with the competitive ELISA. These tests were used previously to screen marine mammal sera for exposure to Brucella spp. (Hanni et al., 2003; Nielsen et al., 2005) but were developed for detection of Brucella abortus in bovine serum. Brucella abortus serology tests likely detect the cross-reacting, newly described marine-origin Brucella spp. (Brucella ceti and Brucella pinnipedalis; Foster et al., 2007), but screening with B. pinnipedalis-specific tests (Meegan et al., 2010) could potentially provide additional information on exposure in Washington sea otters. Because isolation and pathology associated with Brucella have not been documented for sea otters, the population health significance of antibodies to any Brucella spp. is unknown.

Phylogenetic studies indicate that aquatic birds are the principal reservoirs for influenza A viruses (Horimoto and Kawaoka, 2001), which are occasionally transmitted to other species including marine mammals such as seals and whales (Cetacea) (Hinshaw et al., 1986; Callan et al., 1995). In 2011 an unusual mortality event occurred in New England harbor seals (Phoca vitulina) caused by an H3N8 influenza A virus (Anthony et al., 2012). The identified virus resembled one circulating in North American waterfowl since 2002; however, the novel virus was suggested to have mutations that made it virulent in mammals and, therefore, also a potential public health concern (Anthony et al., 2012). Antibodies to influenza A virus were detected in seven Washington sea otters sampled in 2011, representing the first report of exposure in sea otters. Although influenza-associated mortality has not been documented in sea otters, several influenza-associated mortality events have occurred in seals, and exposure in Washington sea otters suggests this pathogen should be considered in future sea otter health surveillance projects.

Leptospirosis is both endemic and capable of causing acute disease outbreaks every 4–5 yr in California sea lions (Lloyd-Smith et al., 2007). Exposure to Leptospira spp. was previously reported for a southern sea otter (Hanni et al., 2003) and northern sea otters (Goldstein et al., 2011). One Washington sea otter (adult female sampled in 2001) was found to have antibodies to Leptospira spp. (1∶100 to L. interrogans serovar gryppotyphosa; Brancato et al., 2009). Exposure to L. interrogans serovar gryppotyphosa was previously detected in marine-foraging river otters from Washington (Gaydos et al., 2007). Given the importance of this disease to other sympatric species (e.g., sea lions and harbor seals; Dierauf et al., 1985; Stamper et al., 1998) and its potential as a zoonotic disease, continued consideration of these bacteria as a potential disease agent in sea otters is warranted.

Caliciviruses have been associated with vesicular disease and abortion in several species of pinnipeds (Smith and Boyt, 1990). Serologic surveys have shown exposure to be common in both pinnipeds and cetaceans even in the absence of outbreaks (Smith et al., 1976). We found no evidence of calicivirus exposure in Washington sea otters, which is consistent with negative findings from serologic surveys of Alaskan and southern sea otter populations (Hanni et al., 2003). Caliciviruses are ubiquitous marine pathogens (Smith et al., 1998a) with zoonotic potential, and exposure was documented in a southern sea otter in rehabilitation (Hanni et al., 2003); therefore, these viruses should continue to be regarded as a potential disease threat for the Washington sea otter population.

Coxiella burnetii, the etiologic agent of Q fever, has a wide host range including humans, wild and domestic mammals, birds, reptiles, and ticks (McQuiston and Childs, 2002). The first report of C. burnetii infection in marine mammals was from a harbor seal with placentitis from California in the late 1990s (Lapointe et al., 1999). Coxiella burnetii infections also were found in Pacific harbor seals, harbor porpoises (Phocoena phocoena), and Stellar's sea lions (Eumetopias jubatus) from the Pacific Northwest (Kersh et al., 2012). No evidence of exposure to C. burnetii was found in Washington sea otters sampled in 2011, but infections observed in sympatric species within the Washington sea otter range suggest that C. burnetii could still pose a risk for sea otters.

Algal toxins have been implicated in mortality events for marine mammals including humpback whales (Megaptera novaeangliae; Geraci et al., 1989), California sea lions (Scholin et al., 2000), and southern sea otters (Kreuder et al., 2003). The sample size available for biotoxin exposure testing in Washington sea otters was small, yet two otters had substantial concentrations of DA and two others had substantial levels of STX. Although all otters appeared healthy at capture, indicating concentrations were not high enough to result in acute signs, repeated or prolonged exposure to DA has been postulated to trigger the progression of myocarditis to dilated cardiomyopathy in southern sea otters (Kreuder et al., 2005). Repeated subacute, low-level exposure to DA can also increase sensitivity to subsequent high-level exposure to DA in zebrafish (Danio rerio; Lefebvre et al., 2012). Further work is needed to understand the cumulative effects of biotoxin exposure in sea otters given the ongoing occurrence of DA and STX in shellfish in coastal Washington (Lewitus et al., 2012).

Significant changes in general health status, as determined from serum biochemistry parameters, were not identified in Washington sea otters over the last decade. Exposure to several pathogens varied in the 10 yr since initial testing. Reintroduction of pathogens, such as CDV, may increase the vulnerability of a naïve population to an epizootic. For other pathogens, such as T. gondii, implications are unclear because prevalence has increased but corresponding mortality has not been observed in the population. Latent T. gondii infections could be reactivated by immunosuppression from concurrent infections with other pathogens or toxicants (Dubey et al., 1989). Brucella and influenza A virus exposure were observed in the population for the first time in 2011. The population significance of these pathogens and those not detected (caliciviruses, C. burnetii) or detected at low prevalence (L. interrogans) is unknown. Given their ability to cause mortality in other sympatric species, or their zoonotic potential, continued monitoring in the Washington sea otter population is advisable. Washington's northern sea otter population has not experienced recent population declines as seen in California's southern sea otter population. The comparatively pristine habitats of Washington sea otters may play an important role in the relative health of this population. As development continues in coastal Washington, however, this population may experience changes in habitat quality and pathogen transmission dynamics.

This project was supported by the US Fish and Wildlife Service, the Washington Fish and Wildlife Office, Division of Listing and Recovery, Interagency Agreement 4500029854 (13320-A-H008). Additional support was provided by US Geological Service–National Wildlife Health Center. Sample collection was done in collaboration with the US Geological Survey–Alaska Science Center and Monterey Bay Aquarium. We thank J. Bodkin, M. Murray, G. Esslinger, B. Weitzman, B. Hatfield, M. Kenner, and J. Tomeleoni for assistance with capture and sampling. We thank R. Hornsby, D. Lindsay, R. Ford, and A. Smith for assistance with testing and interpretation. We are grateful to D. Lynch for logistic support and V. Shearn-Bochsler for review of earlier drafts of the manuscript. Use of trade, product, or firm names is for descriptive purposes only and does not imply endorsement by the US Government.