Victims or vectors: a survey of marine vertebrate zoonoses from coastal waters of the Northwest Atlantic

Abstract

Surveillance of zoonotic pathogens in marine birds and mammals in the Northwest Atlantic revealed a diversity of zoonotic agents. We found amplicons to sequences from Brucella spp., Leptospira spp., Giardia spp. and Cryptosporidium spp. in both marine mammals and birds. Avian influenza was detected in a harp seal and a herring gull. Routine aerobic and anaerobic culture showed a broad range of bacteria resistant to multiple antibiotics. Of 1460 isolates, 797 were tested for resistance, and 468 were resistant to one or more anti-microbials. 73% (341/468) were resistant to 1–4 drugs and 27% (128/468) resistant to 5–13 drugs. The high prevalence of resistance suggests that many of these isolates could have been acquired from medical and agricultural sources and inter-microbial gene transfer. Combining birds and mammals, 45% (63/141) of stranded and 8% (2/26) of by-caught animals in this study exhibited histopathological and/or gross pathological findings associated with the presence of these pathogens. Our findings indicate that marine mammals and birds in the Northwest Atlantic are reservoirs for potentially zoonotic pathogens, which they may transmit to beach goers, fishermen and wildlife health personnel. Conversely, zoonotic pathogens found in marine vertebrates may have been acquired via contamination of coastal waters by sewage, run-off and agricultural and medical waste. In either case these animals are not limited by political boundaries and are therefore important indicators of regional and global ocean health.

Introduction

As human populations in coastal areas continue to increase, coastal ecosystems may become increasingly important as reservoirs or sentinels of infectious organisms from agricultural, animal and human waste. Resultant human and wildlife disease outbreaks and mortality events that occur in the marine environment can increase awareness of the connection between diverse taxa, terrestrial ecosystems, ocean and human health and the risk of infection with zoonotic diseases. Better understanding of the ecology of infectious diseases of multiple taxa of marine animals, which share marine resources and pathogens, will allow for better prediction of the risks to human health. Influences driving the risk of zoonotic infection may include (1) changes in human activity, such as agriculture, (2) increased population density in coastal communities, (3) waste management, (4) consumption of wildlife and (5) changes in medical technology (Hauschild & Gauvreau 1985, Myers et al. 1993, Graczyk et al. 1997, Woolhouse & Gowtage-Sequeria 2005). Thus human activity is resulting in a marine environment in which pathogens, including protozoa, can thrive (Johnson et al. 1997, 1998, Fayer 2004, Boinapally & Jiang 2007).

Infectious diseases can substantially alter long-term trends in populations, including wildlife, or result in short term reductions in local abundance (Heide-Jørgensen et al. 1992). Pathogens, in combination with a weakened population, habitat loss, increased predation, climate change and anthropogenic pollution can also result in severe disease outbreaks which can ultimately lead to extinction of species (Warner 1968, Pounds et al. 2006). Throughout history, there have been numerous opportunities for the introduction of a pathogen to new hosts and the spread to new host populations (Morse 1993, Dobson & Carper 1996, Daszak et al. 2001, Wolfe et al. 2007). Some of the most novel human viruses are zoonotic: the source of these pathogens include the marine environment (Jones et al. 2008), which lacks the barriers inherent in terrestrial dispersal (Morse 1993, McCallum et al. 2003). In the case of West Nile Virus, the spread of the virus occurred along the Atlantic seaboard, a common migration route for many bird species in the northeast US (Rappole et al. 2000). Similarly, the spread of influenza A from aquatic birds is believed to be the most probable source of all influenza A virus strains in other species as was the case in the 1982 mortality event affecting harbor seals in the northeast US (Hinshaw et al. 1984, Callan et al. 1995, Webster 1998, Horimoto & Kawaoka 2001).

Marine vertebrates are no exception to the role of host, and host population, in an environment where for some, diseases are increasing (Harvell et al. 1999, Daszak et al. 2001, Lafferty et al. 2004). The Northeast United States has experienced several epizootic events resulting in mass mortalities of marine mammals and seabirds caused by a variety of viral, bacterial, parasitic, and toxic agents. These events include the 1979–1980 influenza A mortality event and the 1991–1992 phocine distemper morbillivirus (PDV) event in harbor seals Phoca vitulina of New England (Geraci et al. 1982, Duignan et al. 1995). Mass die-offs of birds have been attributed to the introduction and spread of West Nile Virus between 1999 and 2000 in New York State (Bernard et al. 2001). The largest recorded common tern Sterna hirundo mortality event in the National Wildlife Health Center epizootic database was attributed to Salmonella typhimurium at the Monomoy National Wildlife Refuge, Massachusetts (Sohn et al. 2004). Acanthocephalan enteritis has been described in common eiders Somateria mollissima in Massachusetts (Clark et al. 1958), with a recent undiagnosed mortality in Wellfleet, Massachusetts involving over 2400 birds (Jankowsky et al. 2007). A mass mortality of humpback whales in 1987 was attributed to saxitoxin ingestion (Geraci et al. 1989).

Mass beach mortalities of charismatic marine macrofauna garner the attention of many; however, it is the underlying potential for the presence and spread of disease that motivated our regional monitoring of zoonotic pathogens. Increases in human population in coastal communities, human-wildlife interactions, and recognition of the economic as well as social importance of the marine environment of the Northeast US region, contributed to our interest in assessing the prevalence of disease causing microbes. In this study we surveyed a broad cross-section of available hosts within subsets of populations that included live, stranded and fishery by-caught marine vertebrates. These animals were surveyed for bacterial, protozoan and viral pathogens. The specific pathogens targeted were those known to be prevalent in one or more of the vertebrates studied locally or elsewhere in the world. Resource limitation precluded a fully comprehensive survey of all potential zoonotic agents.

Materials and Methods

Sample Collection

Stranded and by-caught mortality samples

Stranded and by-caught birds were collected by the staff at the Seabird Ecological Assessment Network (SEANET, www.tufts.edu/vet/seanet/), Massachusetts Audubon Society, National Oceanographic Atmospheric Administration (NOAA) Northeast Fisheries Science Center (NEFSC) Observer Program and the authors. Marine mammals were collected with the assistance of the New England Aquarium, University of New England Marine Animal Rehabilitation Center, the NOAA NEFSC Observer Program and the authors. Large whale cases were necropsied at the site of stranding (usually beach), and a subset of birds were frozen and then thawed before sampling. Other animals were necropsied in a laboratory between 4 and 48 h post mortem (stored at 4°C overnight). Full necropsies of marine mammals were conducted under protocols described by Pugliares et al. (2007). Necropsies of marine birds were conducted using protocols as described by SEANET. Tissue samples and data are archived at WHOI and Tufts University. Tissue samples were collected using equipment sterilized by rinsing with 95% ethanol followed by flaming with a butane torch.

Live animal samples

Fecal samples were collected from live-caught gulls at Kent Island, Canada, Appledore Island, Maine, and Monomoy National Wildlife Refuge, Massachusetts. Adult great black-backed gulls Larus marinus, herring gulls L. argentatus, and laughing gulls L. atricilla were captured during egg incubation using chicken wire walk-in nest or drop-down traps. Each bird was banded, measured, and pharyngeal and cloacal swabs were collected to obtain samples of bacteria. A fresh sample of feces was also collected from each bird by placing it into a plastic box for <1 min just prior to releasing it; most birds responded to box placement by voiding their cloacas almost immediately. Fecal samples were transferred to sterile cryovials using plastic sterile Pasteur pipettes or syringes. The liner at the bottom of the box was replaced between each bird, so as to avoid contamination. Fecal samples were used for analyses of parasites and bacteria. Pharyngeal swabs were used for analyses of bacteria and influenza. Avian pharyngeal rather than the commonly used fecal swabs were used to allow direct comparability with mammalian nasal swabs.

Fecal samples from seals and birds were collected from beaches in the USA at the Isles of Shoals, New Hampshire/Maine; Great Island in Wellfleet, Massachusetts; Muskeget Island, Nantucket Sound, Massachusetts; Monomoy National Widlife Refuge; and Chatham Harbor, Chatham, Massachusetts. Visual identifications and photographs of the species present at each beach were made before approaching the animals and collecting feces. Animals were identified as harbor seal Phoca vitulina, grey seal Halichoerus gryphus, double-crested cormorant Phalacrocorax auritus, and herring and great black-backed gulls. If a seal haul out site was not >90% of one species, samples were recognized as a mix of the species present (i.e. grey/harbor seal). Bacterial swabs of feces were taken on site. Samples of 1 to 10g were placed on ice in sterile cryovials for molecular analysis and frozen at −70°C on return to the laboratory. Samples for aerobic and anaerobic bacteria were collected using Fisherfinest™ Amies clear gel transport swabs (Fisher Scientific) and submitted within 24 h to IDEXX Laboratories, Grafton, Massachusetts.

Pathogen determination

DNA isolation

Nucleic acids were extracted from tissue samples using the QIAGEN Tissue Kit and from fecal samples using the Mo Bio Soil Kit (Mo Bio Laboratories) following the kit instructions. Urine samples were extracted using the Mo Bio Soil Kit, but with the urine as a volume with weight equal to 250 μg. Samples tested include liver, lung, tracheo-bronchial lymph, spleen, kidney, testes, ovary, uterus, urine, bursa, gut content, feces and brain.

PCR detection

Samples collected from the environment often contain agents that inhibit amplification, so each sample was tested to ensure that it was competent for PCR amplification by using primers flanking a highly conserved fragment of the 18S rRNA gene. All samples that generated a product of the correct size were then tested for human pathogen DNA. In some samples which exhibited amplification inhibition, a 1:10 dilution of the sample eliminated the inhibition and resulted in a product. In these cases, the 1:10 dilution was used for further analysis. All PCR experiments had positive controls for corresponding parasite/pathogen DNA (10 ng per 50 μl reaction) and negative controls for contamination without added template DNA. All PCR reactions were run on agarose gels for detection of products, using a 2% gel for the Giardia products, but a 1% gel for all the others.

Brucella spp. and Leptospira spp. Screening

Only tissue samples and urine were tested routinely for Leptospira spp. and Brucella spp. Leptospira spp. were detected using the Lep1/Lep2 16S rDNA primer set (Merien et al. 1992) and the cycling protocol 94°C (3 min), followed by 40 cycles of 94°C (1 min), 60°C (1 min), 72°C (1.5 min), with a final extension of 72°C (10 min) to produce an approximately 330 bp amplicon. Brucella spp. were detected using the Bru4/Bru5 31 kDa outer membrane protein primer set described in Bailey et al. (1992) to produce an amplicon of approximately 220 bp. The cycling protocol was 93°C (5 min), followed by 40 cycles of 94°C (1 min), 62°C (1 min), 72°C (1 min), and a final extension of 72°C (10 min). On the first 30 necropsies, frozen tissue samples for Brucella spp. determination were sent for culture to the USDA at Ames, Iowa. Frozen tissues were sent to the Oklahoma State Animal Disease Diagnostic Laboratory (OADDL), Stillwater, Oklahoma for determination of Leptospira interrogans by PCR (Acevedo-Whitehouse et al. 2003). Some Leptospira and Brucella amplicons were sequenced to confirm amplification of the correct targets.

Giardia and Cryptosporidium screening

The primer set used most extensively to detect Giardia spp. (GGL639/GGR789) targets a 171 bp fragment of the giardin gene (Mahbubani et al. 1992). These primers were applied in a nested amplification protocol that used 1 μl of the first reaction as template for the second, and each reaction had a total volume of 25 μl. Amplification parameters were 94°C (2 min), followed by 94°C (30 s), 56°C (30 s), 72°C (1 min), and a hold at 4°C. The first amplification was carried out for 25 cycles, and the second amplification was 40 cycles. Samples positive by Giardia genus amplification were tested for Giardia intestinalis using the primer set MAH433F/MAH592R (Rochelle et al. 1997), with cycling parameters 94°C (4 min), followed by 94°C (1 min), 60°C (1 min), 72°C (1 min), and a hold at 4°C. Again a nested amplification strategy was applied, with the first using 25 cycles and the second using 40 cycles. Reactions were a total volume of 25 μl each. Our samples positive for the Giardia genus were genotyped by our collaborators (Lasek-Nesselquist et al. 2008).

The PCR primers used for Cryptosporidium 18S rDNA detection were the nested set WR494F/AWA1206R and CPB DiagF/PW99R (Ward et al. 2002), resulting in an amplicon of approximately 420 bp. The nested protocol used 25 μl reactions and cycling parameters of 94°C (10 min), followed by 40 cycles of 94°C (30 s), 58°C (40 s), 72°C (40 s), and a hold at 4°C. One microliter of the first reaction was used as template for the second reaction with the same cycling parameters. Samples positive using the genus primers were tested for Cryptosporidium parvum using the primer sets Cry5/Cry6 and NCryp1/NCryp2 described previously (Mayer & Palmer 1996). The first reaction was 94°C (2 min), followed by 35 cycles of 94°C (30 s), 56°C (30 s), and a final extension at 72°C (1 min). The second reaction was 94°C (2 min), followed by 40 cycles of 94°C (30 s), 60°C (30 s), 72°C (30 s), and a final extension at 72°C (3.5 min). Some Giardia and Cryptosporidium amplicons were sequenced to confirm correct target amplification.

Bacterial culture with antibiotic sensitivity

Routine sampling sites included fecal/cloaca swabs for live animals, and thorax (using an intercostal approach to the pleural space) and abdomen or coelom (using a lateral abdominal approach to the peritoneal space) for those examined by necropsy. Thorax and abdomen/ coelom sample sites were flame seared and incised with a sterile blade. Swabs from nasal/blowhole/nares were collected as appropriate and practical on live animals and if contamination of the outside surface of dead animals was minimal. Other sites were chosen for bacterial isolation if lesions or infection were suspected. Cultures for fungal agents were only submitted if suspected at gross necropsy. All samples were collected using sterile methods. Swabs were shipped overnight to IDEXX Laboratories (Grafton, Massachusetts) and plated on blood agar, and MacConkey plates for aerobic culture, and blood agar, MacConkey and anaerobic blood agar plates for anaerobic culture.

Anaerobic and anaerobic bacteria were recovered, identified and aerobic bacteria were tested for antibiotic sensitivity using the Vitek system (bioMérieux Vitek). Requests were made for culture to include Campylobacter and Salmonella for fecal swabs. All others were requested for routine cultures. No growth was assumed after 48 h of negative culture. Antibiotics tested routinely included amikacin (AMK), ampicillin (AMP), augmentin (amoxicillin + clavulanic acid, AUG), carbenicillin (CAR), ceftazidime (CAZ), ceftiofur (CEF), cephalothin (CEPH), cloramphenicol (CHL), ciprofloxacin (CIP), gentamycin (GEN), tribrissen (TRI), piperacillin (PIP), enrofloxacin (ENR), tetracycline (TET), ticarcillin (TIC), and tobramycin (TOB). Penicillin G (PENG), vancomycin (VAN), oxacillin (OX) and erythromycin (ERY) were tested only with the following bacteria: Enterococcus spp. were only tested for AMP, CHL, TET, PENG, and VANC sensitivity; and Staphylococcus spp. were only tested for AUG, AMP, CEF, ENR, GEN, TET, OX, PENG and VANC sensitivity. Multiple antibiotic resistance (MAR) indices (Kruperman 1983) were calculated for each isolate, and ranged from 0 to 1.

Influenza virus isolation

Viral swabs of lung and nasal/blow/nares were placed in viral media (Hardy Diagnostics). Viral samples for molecular and culture analysis were frozen at −70°C and sent to the United States Geological Survey (USGS) National Wildlife Health Laboratory (NWHL), Madison, Wisconsin, for influenza A and B type analysis. Each sample was tested by RNA extraction and by the Matrix RT-PCR test for avian influenza. Influenza B was tested using the BD Directigen Flu A/B test (BD, Franklin Lakes, New Jersey). Additional diagnostic tests on a random selection of 25 oral/nasal/blow samples were tested using the Remel XPECT™ FLU A&B Test Kit (Remel) for influenza A and B identification.

Histopathology

Histopathology samples were fixed in 10% neutral buffered formalin, dehydrated and embedded in paraffin and sectioned for examination of 5 μm hematoxylin-and-eosin stained slides by Northwest ZooPath (Monroe, Washington) and the University of Tennessee, College of Veterinary Medicine (Knoxville, Tennessee).

Gross and histopathology analysis

Using histopathology, gross pathology and molecular results, each case was categorized in one of 14 categories of significant findings: (1) bycatch with no significant findings (NSF), (2) bycatch with gas emboli/bubbles, (3) bycatch with disease process, (4) bycatch other, (5) could not be determined (CBD), (6) emaciation, (7) infectious disease (bacterial/viral/fungal), (8) human interaction, (9) mass stranding NSF, (10) neurological, (11) other, (12) parasitism, (13) predation, and (14) tournament caught. We did not analyze by cause of death given the presence of significant pathology in some by-caught animals. Where an animal could fall into 2 categories, the category that most specifically described the gross and histopathological findings was chosen.

Results

Between December 2005 and August 2007 a total of 370 live, stranded, fishery by-caught and tournament-caught marine vertebrates were sampled: 165 individuals of 15 species of marine mammals, 192 individuals of 15 species of seabird, and 13 individuals of 3 species of shark (Table 1). Geographic ranges of the animals collected extended north to Kent Island, Canada (44.58°N, 66.75°W) and south to Virginia, United States (38.82°N, 75.95°W) (Fig. 1). Of the total stranded and by-caught animals, 167 cases were examined by gross necropsy including 96 stranded birds, 45 stranded marine mammals, 22 by-caught marine mammals and 4 by-caught birds. Individual species sample sizes are mostly too small for statistical tests, but the findings are relevant for the identification of host species and situations requiring further study.

Table 1. Individual marine vertebrate species surveyed for zoonoses. Values are numbers of individuals.

Species	Common name	By-caught	Live	Stranded	Total
Marine mammals					165
Balaenoptera acutorostrata	Minke whale			2
Cystophora cristata	Hooded seal			1
Delphinus delphis	Short-beaked common dolphin	1		16
Globicephala melas	Long-finned pilot whale			2
Grampus griseus	Risso's dolphin			2
Halichoerus grypus	Grey seal	5	58	2
Kogia breviceps	Pygmy sperm whale			2
Lagenorhynchus acutus	Atlantic white-sided dolphin	1		4
Megaptera novaeangliae	Humpback whale			1
Phoca groenlandica	Harp seal	5		7
Phoca vitulina	Harbor seal	3	12	2
Phocoena phocoena	Harbor porpoise	7		1
Stenella coeruleoalba	Striped dolphin			1
Mixed haul outa	Grey or harbor seal		28
Unidentified delphinid	Unidentified delphinid			1
Ziphius cavirostris	Cuvier's beaked whale			1
Seabirds					192
Charadrius melodus	Piping plover			2
Somateria mollissima	Common eider			47
Gavia immer	Common loon			9
Sterna hirundo	Common tern			1
Larus marinus	Great black-backed gull		17	9
Phalacrocorax carbo	Great cormorant			3
Puffinus gravis	Greater shearwater	3		1
Larus argentatus	Herring gull		64	12
Larus atricilla	Laughing gull		4
Sternula antillarum	Least tern			2
Moras Bassanus	Northern gannet			6
Phalacrocorax auritus	Double crested cormorant	1	6	1
Gavia stellata	Red throated loon	1		1
Melanitta perspicillata	Surf scoter			1
Melanitta deglandi	White winged scoter			1
Sharks					13
Aliopias vulpinus	Thresher shark	9
Isurus paucus	Mako shark	3
Prionace glauca	Blue shark	1
Total		40	189	141	370

^aFecal samples were collected from a haul out of a mixture of Halichoerus grypus and Phoca vitulina

Fig. 1.

Locations and types of bird and mammal samples examined for molecular analysis, microbiology and pathology.

Molecular screening results

A total of 635 samples were analyzed from 236 animals (Table 2). Amplification of Brucella spp. was conducted in 109 animals. Positive tissues were found in 38 animals within 16 species of stranded and by-caught birds, dolphins, seals, and whales. Species included hooded seal Cystophora cristata, harp seal Phoca groenlandica, grey seal Halichoerus grypus, an unidentified delphinid species, common dolphin Delphinus delphis, long-finned pilot whale Globicephala melas, Risso's dolphin Grampus Griseus, Atlantic white-sided dolphin Lagenorhynchus acutus, humpback whale Megaptera novaeangliae, common eider Somateria mollissima, common loon Gavia immer, great black-backed gull Larus marinus, great cormorant Phalacrocorax carbo, greater shearwater Puffinus gravis, herring gull Larus argentatus and northern gannet Moras bassanus. The highest prevalence was in stranded seals (58%), with the highest number of positive results in stranded birds (14). Brucella amplification products were detected in tissues including brain, liver, kidney, spleen, lung, testes, and tracheo-bronchial lymph node. Brucella spp. appeared to occur more commonly in males (Table 3) and more prevalent in samples collected from winter months (Table 4). Amplicon sequencing confirmed that the correct target was detected, and speciation of Brucella types is ongoing and will be reported elsewhere.

Table 2.

Prevalence of target pathogens by PCR testing. N = number of individuals sampled. For Brucella, Leptospira and Cryptosporidium, parenthetical values are no. of individuals positive (all individuals tested); for Giardia, fraction indicates no. positive out of the no. of individuals in the tested subset

Type	Animal	N	% Brucella spp.	% Leptospira spp.	% Giardia spp.	% Cryptosporidium spp.
Live	Bird	84			16 (5/31)	2 (2)
	Seal	95			10 (4/40)	23 (22)
Stranded	Bird	34	41 (14)	18 (6)	12 (3/25)	0
	Seal	12	58 (7)	0	16 (1/6)	25 (3)
	Dolphin	19	42 (8)	11 (2)	9 (1/11)	0
	Porpoise	1	0	0	100 (1/1)	0
	Whale	7	29 (2)	28 (2)	20 (1/5)	0
By-caught	Bird	4	25 (1)	25 (1)	0	0
	Fish	10	0	0	100 (1/1)	0
	Seal	13	38 (5)	8 (1)	25 (1/4)	15 (2)
	Dolphin	2	50 (1)	0	100 (2/2)	0
	Porpoise	7	0	14 (1)	40 (2/5)	14 (1)

Table 3.

Seasonal distribution of pathogen detection (percentage positive) by PCR testing. Fall: Oct–Dec; Winter: Jan–Mar; Spring: Apr–Jun; Summer: Jul–Sep. Parenthetical values are no. of positive results out of no. of samples tested

Pathogen	Fall		Winter		Spring		Summer
Leptospira spp.	13.33	(2/15)	18.18	(4/22)	8.33	(2/24)	10.00	(3/30)
Brucella spp.	37.50	(6/16)	60.87	(14/23)	27.27	(6/22)	38.71	(12/31)
Giardia spp.	30.77	(4/13)	16.95	(10/59)	18.18	(6/33)	4.76	(1/21)
Cryptosporidium spp.	17.78	(8/45)	10.20	(5/49)	6.15	(4/65)	8.33	(4/48)

Table 4.

Prevalence of pathogen detection according to sex, based on the number of individuals of each sex tested (ND = sex could not be determined). Parenthetical values indicate no. of individuals positive

Pathogen	Female	Male	ND
Brucella spp.	24 (9)	74 (28)	3 (1)
Leptospira spp.	18 (2)	73 (8)	9 (1)
Giardia spp.	60 (6)	22 (6)	8.5 (8)
Cryptosporidium spp.	9 (2)	18 (4)	72 (16)
Influenza A & B	0 (0)	3 (1)	3 (1)

Brucella amplicons at times corresponded to pathologies that could be associated with infection. In a mass stranding of 9 common dolphins, 3 out of 5 dolphins sampled tested positive for Brucella spp. in brain and uterus. Two dolphins were diagnosed with cervicovaginolithiasis, one associated with chronic endometritis with ecchymotic hemorrhaging in the right and left uterine horns. The third dolphin exhibited mild meningio-encephalitis with evidence of renal dysfunction. Additionally, a young stranded harp seal in which multiple tissues resulted in positive Brucella amplification, exhibited a vaginal myxoid leomyoma. Infections due to Brucella spp. in birds were not identified. Brucella cultures from samples sent to USDA were all negative.

Tissues from a total of 109 animals were analyzed for Leptospira spp. Positive amplification resulted in 11 animals from 9 species including stranded common eiders, common dolphin, unidentified dolphin species, humpback whale, harp seal, herring gull, northern gannet, one by-caught greater shearwater and one by-caught harp seal. Tissues that yielded amplicons included brain, kidney, liver, spleen, testes, tracheobronchial lymph, urine, feces and gut content. Leptospira PCR was negative for all samples analyzed by OADDL, and our sequencing of amplicons indicated that the correct target was not being recovered using the Lep1/Lep2 primer set, despite the correct size of the products.

A total of 236 animals were sampled for Cryptosporidium spp.: the parasite was detected in 30 animals including live seals, stranded seals, by-caught seals, live herring gulls and by-caught porpoise (Table 2). Samples obtained in fall months resulted in more positive results (Table 4). The highest prevalence was found in live seals (23%) and stranded seals (25%), specifically stranded harp seals Phoca groenlandica, live grey seals Halichoerus grypus, and samples collected from mixed haul-out sites of grey and harbor seals. Live herring gulls Larus argentatus (n = 2) were the only bird species to test positive. Amplicon sequencing confirmed the correct target detection.

Giardia spp. amplifications are reported for only a portion of the sample set: 131 animals were tested, with a total of 22 positive for the parasite. Positives were found in animals of all species, with roughly equal prevalence for groups with sample numbers greater than 5 (Table 2). Animals included caught thresher shark Aliopias vulpinus, by-caught harbor seal Phoca vitulina, stranded and by-caught common dolphin Delphinus delphis, harbor porpoise Phocoena phocoena and Atlantic white-sided dolphin Lagenorhynchus acutus, stranded long-finned pilot whale Globicephala melas, Risso's dolphin Grampus griseus, harp seal Phoca groenlandica, and common eider Somateria mollissima. Live animals testing positive included herring gulls Larus argentatus, harbor seals P. vitulina and grey seals Halichoerus grypus. Again, a higher number of positive samples overall were recovered in fall and winter months (Table 3). Of 22 positives, 20 yielded amplification products and sequences for G. intestinalis speciation (Lasek-Nesselquist et al. 2008), but 2 yielded giardin products that shared sequence similarity with Assemblage F. Both of these samples were from live mixed grey/harbor seal populations.

Results have only been reported for samples for which the presence of Giardia has been confirmed by sequence analysis of the giardin product or via speciation of G. intestinalis (Lasek-Nesselquist et al. 2008). Amplification of fecal or gut sample extracts using the GGL/GGR primer set yielded several incorrect amplicons. One was a distinctly smaller band, which yielded a non-giardin sequence. The other was a band that appeared to be the correct size, but also yielded a non-giardin sequence.

Bacterial culture and antibiotic resistance

A total of 95 bacterial and 1 fungal species were identified to genus level at a minimum. Fecal and cloacal swabs, specifically those from live birds, had the greatest diversity of microbes cultured from routine and non-routine sites (Appendices 1 & 2). Non routine sites include those related to pathology and infection sites including lesions, abscesses, urine, organs, and abdominal or thoracic fluid. Oral swabs were not taken in all animals and are considered non pathology related for this survey. Aerobic Gram-negative bacilli comprise 76% of the isolates and 8.5% represented anaerobic organisms. Escherichia coli was most commonly isolated overall (152 isolates), especially in live and stranded birds and marine mammals. Pseudomonas spp., Clostridium perfringens, Enterobacter cloacae, Enterobacter spp., and Shewanella spp. were the next most commonly isolated. A total of 10 bacteria were only associated with non-routine culture sites. Of these, 9 were aerobic Gram-negative bacilli: Chromobacterium violaceum (kidney), Empedobacter brevis (kidney), Enterobacter sakazakii (uterus), Kluyvera spp (genital), Providencia stuartii (omentum), Pseudomonas oryzihabitans (urine, spleen), Salmonella spp. (spleen), Sphingomonas spp. (spleen) and Vibrio fluvialis (genital). Brevibacterium spp. (mandible, periaortic) was the only anaerobic Gram-positive bacillus represented in non-routine culture swabs.

Appendix 3 lists bacteria isolated in this study that are recognized as human pathogens by the American Biological Safety Association (ABSA: www.absa.org/XriskgroupsX/index.html), or other publications, along with references to published human infections where applicable. Sixty-eight of the bacterial isolates were recognized as human pathogens by ABSA (71.6%), and a greater portion were identified by searching the medical literature for cases of human infection (up to 80%). Many isolates appeared to be species-specific. Pasteurella multocida and Shewanella algae were only recovered from common eiders stranded in Wellfleet, Massachusetts. Enterococcus faecalis was only recovered from stranded birds. Isolates of Ewingella americana and Peptostreptococcus spp. were only recovered from cetaceans. Chryseobacterium indologenes was only recovered from by-caught seals, while Clostridium spp. were most common in samples from live seals.

Antibiotic resistance for each isolate ranged from 0 to 13 antibiotics. Sixty-one percent of bacterial isolates were resistant to at least one antibiotic, while 58.8% were resistant to more than one. Isolates with a MAR value of 1 (i.e. resistant to all antibiotics tested) were a Serratia marcesens and a Shewanella spp. from a stranded hooded seal, but these isolates were tested with a limited number of antibiotics (6 and 3 respectively), so it is unknown whether this value would have remained high if the others were tested. 38.7% of our isolates had a 0 MAR value, 30.9% had a MAR value >0 but <0.2, while 30.0% had a MAR value >0.2. The bacterial isolate that was resistant to the greatest number of antibiotics was a Chryseobacterium indologenes from a by-caught harp-seal that showed resistance to 13 out of 16 antibiotics. The animal yielding the greatest percentage of isolates with multiple resistances was a stranded meningio-encepahlitic Cuvier's beaked whale, where 7 of the 8 isolates tested for ABR were resistant to >4 antibiotics. Antibiotics to which isolates showed the least resistance were ciprofloxacin (2%), enrofloxacin (2%), gentamicin (4%), oxacillin, vancomycin, and erythromycin. The antibiotics with the highest number of resistant isolates included cephalothin (39%), ampicillin (34%), augmentin (26%) and carbenicillin (26%).

Influenza

Influenza A and B were tested in 34 samples. There were 2 positive samples for influenza A but none for influenza B. Influenza A, avian influenza H3N8 virus, was detected in one by-caught harp seal. Influenza A, negative for H5 or H7, was detected in one live herring gull from Kent Island, Canada. The avian influenza type isolated was confirmed not to be of agricultural interest but the actual type has yet to be confirmed.

Significant findings in mortality cases

A diagnosis based on significant findings or ultimate cause of death based on history, gross and histopathology results was assigned to 115/121 (96%) of animals (Table 5). Six cases could not be diagnosed based on available data. The most common significant findings were related to infectious disease (31%) followed by the category of other (12%) which includes trauma involving wing fracture, con-specific aggression, a wound of unknown origin or moving vehicle, obvious gross changes such as peritonitis with no apparent cause, gastrointestinal obstruction, congenital defect, and dependent pup or calf not able to forage independently. Parasites as the primary cause of stranding and mortality were highest in common eiders (10% of the cases). Of animals that were fishery by-caught, the majority had no significant findings (7%) other than pathology associated with drowning and/or were found by gross and histopathology to exhibit gas emboli (7%) in lymph nodes, brain, myocardium, adrenal glands, spleen, skeletal muscle, and kidney.

Table 5. Significant findings based on necropsy and histopathology findings.

Most significant finding	% (Total)
Infectious disease	26 (37)
Mass stranding—no significant findings	19 (28)
Other	10 (15)
Tournament/hunt	9 (13)
Parasites	8 (12)
Bycatch—no significant findings	6 (8)
Bycatch—bubbles	6 (9)
Could not be determined	5 (7)
Bycatch—disease	4 (6)
Human interaction	3 (5)
Emaciation	1 (2)
Bycatch—other	21 (2)
Predation	11 (1)
Total	145

In general, the pathologies were variable, but some were observed more often in particular circumstances. For example, the pathologies most often seen in stranded animals included peritonitis, septicemia, hepatitis, aspergillosis, enterotoxemia, reduced nutritional state, bacterial and verminous enteritis, verminous gastritis, and interstitial and bronchopneumonia. A summary of pathologies noted in relation to bacteria isolated can be found in Appendix 3.

Discussion

Marine mammals, sea birds and sharks of the NW Atlantic harbor zoonotic bacteria including Brucella spp. and Leptospira spp., protozoan pathogens Cryptosporidium spp. and Giardia spp., and multiple strains of zoonotic bacteria that are resistant to multiple antibiotics used in both human and animal treatment. One marine mammal and one sea bird also tested positive for avian influenza, specifically H3N8 in a by-caught harp seal and unspecified non H5 type in a herring gull.

Brucella spp. was the most commonly detected target zoonosis found in both stranded marine mammals and sea birds. Isolation and detection of Brucella spp. has been documented in harbor and harp seals along the coast of southern New England (Connecticut and Rhode Island) with no gross or histological changes associated with infection (Maratea et al. 2003). Six species of Brucella are currently recognized and ongoing research suggests 3 additional specific to marine mammals: B. pinnipedialis and B. ceti (Foster et al. 2002, 2007) or B. phocae and B. phocoenoae and B. delphini (Groussaud et al. 2007). Marine mammal Brucella strains have been isolated in association with pathology and infection in humans, although these cases did not involve direct contact with infected marine mammals: one was a laboratory technician who was infected in a laboratory (Brew et al. 1999), and the others were 2 individuals from Peru who had no contact with marine mammals (Sohn et al. 2003). In marine mammals, infection is characterized by chronic infection which can lead to weight loss, inflammation, abortion and infertility (Koneman et al. 1988, Miller et al. 1999), menigioencephalitis (Gonzalez et al. 2002) and bone disease (Dagleish et al. 2007).

With regards to the cervicovaginolithiasis described in common dolphins, over the course of 8 yr of marine mammal stranding reports by the Cape Cod Stranding Network, this was the first recorded instance of vaginal stones in cetaceans. Presence of vaginal calculi in stranded dolphins has been hypothesized to be composed of calcium phosphate and the result of ossification of a developing or aborted fetus (Sawyer & Walker 1977, Benirschke et al. 1984, Woodhouse & Rennie 1991). Immunohistochemistry and sequencing of these amplicons are underway and will be reported elsewhere.

In terms of Brucella amplicons in birds, previously published studies note positive antibody response to Brucella abortus and Brucella melitensis in domestic fowl without isolation and identification of the organism (Abdu et al. 1984, Kumar et al. 1984, Kudi et al. 1997, Junaidu et al. 2006). The presence of the bacterial amplicon in birds, and at the high frequency seen in this study, suggests wild birds could be a source of infection for other species.

Leptospirosis is considered the most widespread zoonosis in the world (Levett 2001). While the west coast of the US has experienced severe epizootics of Leptospira interrogans in pinnipeds, populations on the east coast appeared to remain naïve (Gulland et al. 1996, Stamper et al. 1998, Colegrove et al. 2005), although leptospirosis is known to be enzootic in western and central Massachusetts (Andrew & Marrocco 1977). Samples in our study that yielded amplification products included a harp seal, a humpback whale, unidentified species of dolphin, common dolphin, common eider, great cormorant, greater shearwater, herring gull and northern gannet. As pathologies corresponding to leptospirosis were not noted in these animals, the diversity of hosts could represent non-pathogenic species from the marine environment. Sequencing of amplicons from the humpback gut content sample and an eider bursa sample indicated that the approximately 300 bp products were similar to Atopobium spp. (87% and 95% respectively), members of the Coriobacteriacea (Actinobacteria). These results indicate that the primer set was not amplifying the correct product in our samples, and that the positive results we obtained are not indicative of Leptospira. We have chosen to report these results in order to document the problem, and we intend to re-analyze the samples using a different set of Leptospira primers (Cameron et al. 2008).

Giarda and Cryptospordium are intestinal protozoan parasites (Fayer 2004, Ford 2005) that infect a wide range of animals, including humans. The presence of Giardia spp. and Cryptosporidium spp. in marine mammals indicates that these animals can serve as vectors of these primarily fresh-water parasites, and could be acquiring them from anthropogenic sources. There is also the possibility that novel marine strains of these parasites exist, and this is supported by the discovery of novel seal genotypes of Cryptosporidium (Santin et al. 2005). Giardia from samples from marine mammals, sea birds and a shark from this study have been confirmed as G. intestinalis of human assemblagesA and B (Lasek-Nesselquist et al. 2008) and members of the assemblage F were also present in seals. To our knowledge, our study is the first to report on the prevalence of Giardia in wild dolphins and porpoises. Other studies have reported on the prevalence of Giardia in seals and whales, although genotyping has not routinely been accomplished (Olson et al. 1997, 2004, Measures & Olson 1999, Hughes-Hanks et al. 2005). Ringed, grey, harp and harbor seals, as well as right whales and bowhead whales have all been found to harbor Giardia spp. with a general prevalence between 20 and 30%, with one exception (Hughes-Hanks et al. 2005) of much higher prevalences for ringed seals (64.5%) and right whales (71.4%). This study showed similar, and in some cases lower, prevalence values (Table 2). There was also some variability in prevalence, and this may be due to the small sample sizes and the general health of the animal (live, stranded or by-caught). When the prevalence of Giardia was calculated without considering animal status, the values became 12% for seals, 13% for birds, 23% for dolphins, 20% for whales and 50% for porpoises.

Our results for Cryptosporidium are in distinct contrast to our findings for Giardia. Cryptosporidium was found only in seals and porpoises, and in a very small number of our birds. The prevalence values for seals and porpoises are close to those observed in other marine mammals studies (18 to 24%) (Hill et al. 1997, Deng et al. 2000, Hughes-Hanks et al. 2005, Santin et al. 2005). Sequence analysis of our amplification fragments from seals indicated that they were not harboring C. parvum, but appeared to carry species related to C. muris and a Type 2 novel seal isolate (Santin et al. 2005).

Tables 3 & 4 illustrate some of the general trends we have observed in our data. In Table 3, we examined the potential for season to influence the detection of the pathogens. The results indicate that fall and winter have higher prevalences for Giardia, Cryptosporidium and Brucella, and this raises questions about how temperature and other seasonal parameters (freshwater input, migration, mating, food resources) impact pathogen prevalence. In Table 4 we examined the detection of pathogens by sex. Of particular interest is the preponderance of Brucella in males (Table 4), which raises the question of whether the association of Brucella and abortion has lead to a misplaced focus on females. The increased prevalence of Giardia in females is also interesting, and suggests there may be behavioral factors involved in the presence of this particular pathogen.

The majority of bacteria isolated in our study were recognized as human pathogens or potential human pathogens. All pathogens found in common between marine mammals, sea birds and sharks are recognized by the ABSA as human pathogens: Acinetobacter calcoaceticus-baumannii, Citrobacter braaki, C. freundii, Enterobacter cloacae, Leclercia adecarboxylata, Morganella morganii, Pseudomonas aeruginosa, Pseudomonas spp., Shewanella spp. and Stenotrophomonas maltophilia. Other isolates recovered that are known to cause infection in humans from handling fish include Aeromonas hydrophila, Edwardsiella tarda, Vibrio cholera, and V. parahaemolyticus (Harper 2002). The routine microbiological survey did report non-pathogenic organisms when present, but did not assess antibiotic resistance.

The recovery of antibiotic resistant bacterial isolates from marine animals was not unexpected. Other studies have reported the presence of antibiotic resistant bacteria in marine animals (Johnson et al. 1998, Smith et al. 2002, Foster et al. 2004, Stoddard et al. 2005, Buck et al. 2006) and bacteria isolated from marine birds at rehabilitation facilities in California also frequently yielded Escherichia coli (Steele et al. 2005), with over 50% of isolates resistant to ampicillin. Our study is the first to examine such a wide range of organisms and sample types (tissues as well as external swabs). One of the most interesting findings in the study was the presence of multiple antibiotic resistant bacteria in a Cuvier's beaked whale, a species associated with deep water (1000 to 3700 m) and rarely seen in coastal environments (Ferguson et al. 2006). Isolates included: Photobacterium damiela resistant to augmentin, ampicillin, carbenicillin, cephalothin, and ticarcillin; several Pseudomonas spp. resistant to augmentin, ampicillin, carbenicillin, ceftazime, cephalothin, and chloramphenicol; and Acinetobacter calcoaceticus-baumannii resistant to ampicillin, ceftazime, cephalothin, and chloramphenicol. Clostridium perfringens, Candida glabbata, Pseudomonas oryzihabitans, and Staphlococcus spp. were also isolated from this animal. Where this animal acquired bacteria with levels of antibiotic resistance indicative of significant contamination is a question that remains unanswered, and merits further study. However, given the finding of significant PCB burdens in deep sea fish (Stegeman et al. 1986), it would seem that terrestrial sources of these drug resistance genes may similarly have deep water sinks. It is also interesting that high ambient pressure may in itself enhance antibiotic resistance development (Hind & Attwell 1996).

Thresher and mako sharks sampled off Martha's Vineyard, Massachusetts in this study also exhibited bacterial isolates with multiple antibiotic resistances. The isolates averaged resistance to 4 antibiotics, with a range of 0 to 8, augmentin and cephalothin resistance being the most common. Although our sample set was small, the resistance in sharks corroborates previous findings of antibiotic resistant bacteria in smooth dogfish shark Mustelus canis from the same area and a study of nurse Ginglymostoma cirratum, bull Carcharhinus leucas and spinner sharks Carcharhinus brevipinna (Blackburn 2003). Some of these sharks do forage in coastal environments, and even those that do not may encounter food that has come from the coastal environment. Our lack of knowledge regarding the natural histories of many of these animals limits our ability to identify sources of contamination.

The overall prevalence of isolates resistant to multiple antibiotics, and the number of isolates that had MAR indices >0.2 (30.9%) were surprising. MAR values greater than 0.25 are considered to represent exposure to point-source contamination (usually human fecal) (Kruperman 1983, Kaspar et al. 1990, Parveen et al. 1997). In our results 27.4% of our isolates had a MAR value greater than 0.25, suggesting that the animals in our study were potentially being exposed to significant contamination. It seems reasonable to begin considering where and/or how these animals are being exposed, not only with concern for their health, but the fact that they can serve as vectors of antibiotic resistant bacteria over ranges that can exceed 10° of latitude/longitude.

The presence of multiple antibiotic resistance in isolates that are not recognized as pathogens is also extremely important, as this indicates that commensal or environmental bacteria can serve as reservoirs for resistance genes. While it is generally agreed that the widespread use of antibiotics has resulted in significant increases in antibiotic resistance, recent work has shown that even after the removal of the selective pressure of individual or groups of antibiotics, resistance levels have been slow to decline (Heuer et al. 2002, Sørum et al. 2006). This suggests that the maintenance of resistance genes is not necessarily detrimental to cells, and that there may be other factors associated with the maintenance of these genes (e.g. heavy metals) (Sjogren & Port 1981, Baker-Austin et al. 2006). The overall concern is that commensal and environmental bacteria are not only able to acquire and maintain resistance genes, but that they are able to multiply and spread them to others, including back to pathogenic species either in the environment or in the host. Most of the marine animals sampled have extensive migratory and foraging ranges, and it is likely that they could serve as vectors in the spread of antibiotic resistance in the marine environment.

Our results indicate that marine mammals, fish and seabirds may not only suffer as victims of disease from zoonotic pathogens, but also act as vectors, moving these human bacterial and protozoal pathogens to different geographic locations in the ocean and terrestrial environments. Marine animals interact with each other as predators, scavengers and through the shared use of marine and beach environments. Documentation of seals predating on sea birds, sea birds predating and scavenging on marine mammals and sharks predating on marine mammals out of rehabilitation facilities are a few examples that support this hypothesis (Tallman & Sullivan 2004). They come into contact with humans and terrestrial animals as food resources, during stranding events and through shared use of beach environments. The prevalence of human genotypes of Giardia intestinalis in both seals and gulls that share local beach environments is intriguing, and whether this represents contamination of the marine populations from human sources remains a question. While our knowledge regarding the presence of zoonotic agents in marine animals is progressing rapidly, very little is known about the potential impacts of these agents on both marine animal health and potential risks to human health.

The long-distance migration of marine vertebrates and their specific ocean usage areas of the marine environment may allow for specific patterns in anthropogenic movement of infected host pathogen pollution (Daszak et al. 2001). In combination with global climate change, fishery decline, poor nutritional status, and overlap of new populations, pathogen exchange in these areas can occur. For instance, it is widely believed that harp seals foraged further south in 1987 when fish populations decreased in the Barents Sea, which in turn allowed harp seals carrying phocine distemper virus (PDV) to interact with naïve populations, initiating the 1988 PDV outbreak in harbor seal (Dietz et al. 1989, Heide-Jørgensen et al. 1992, Gulland & Hall 2003, Härkönen et al. 2006). Leptospira outbreaks, and the presence of Toxoplasma gondii in sea otters in California are also examples of increasing human populations, interactions with wildlife and disease transmission (Stamper et al. 1998, Miller et al. 2002). Transmission from humans to marine life is evident in the unique case of influenza B transmission to a seal (Osterhaus et al. 2000). This evidence reminds us that while wildlife may act as vectors of zoonotic disease, their role as sentinels to the abundance and distribution of human waste is one that needs more attention. It is important to establish what may be endemic to marine environments (e.g. Vibrio spp. carrying antibiotic resistance) versus what is introduced by anthropogenic activity. Ongoing studies are using microbial source tracking methods to determine whether seabirds (gulls in particular) harbor fecal pathogens derived from anthropogenic sources, and if these pathogens are transmitted via birds to other coastal animals such as seals.

The overall goal of this research was to assess prevalence of subclinical and clinical zoonoses in marine mammal and birds of the Northwest Atlantic and bring awareness of the diversity of emerging and potential pathogens. This region supports some of the largest fisheries in the world, is recognized as an important breeding and nesting region for coastal birds, supports a large diversity of marine mammals and is an area where coastal human communities are becoming increasingly overcrowded. Concern regarding wildlife and human interactions raise the need to understand what pathogens are able to infect both, and which ones may be increasing in occurrence due to anthropogenic activities. More importantly, wide distribution of information between researchers and users of the ocean environment will help determine where pathogens originate, where they might be going and how best to prevent exposure. In this regard it is significant that a recent review of emerging infectious diseases shows the northeast of the United States as having the highest relative risk for emerging infectious diseases in the USA (Jones et al. 2008).

Appendix 1.

Number of bacterial isolates cultured compared between live, stranded and by-caught marine vertebrates.

Microorganism	Marine mammal			Marine bird			Shark	Total
	Live	Stranded	By-caught	Live	Stranded	By-caught	Caught
	N = 33	N = 37	N = 16	N = 54	N = 50	N = 5	N = 8
Aerobic Gram-negative Bacilli
Achromobacter (alcaligens) xylosoxidans subsp. xylosoxidans					1			1
Acinetobacter calcoaceticus-baumannii		3	4		3		1	11
Acinetobacter spp.	3	4	4	1	4			16
Aeromonas hydrophila		1			6			7
Aeromonas spp.		3		2	1	2		8
Alcaligenes faecalis					1			1
Burkholderia cepacia	1	1		4				6
Chromobacterium spp.							1	1
Chromobacterium violaceum		2						2
Chryseobacterium indologenes			2					2
Citrobacter braaki	2		1	2			1	6
Citrobacter freundii	1	2		5	4		2	14
Citrobacter koseri					1			1
Citrobacter spp.				1				1
Edwardsiella hoshinae	1	1						2
Edwardsiella tarda	4	7		1				12
Edwardsiella spp.	1				2			3
Empedobacter brevis					1			1
Enterobacter amnigenus				3				3
Enterobacter cancerigenus					1			1
Enterobacter cloacae		2	2	10	5	1	3	23
Enterobacter intermedius					1			1
Enterobacter sakazakii			1					1
Enterobacter avium					1			1
Enterobacter spp.		5	2	12	5			24
Escherichia coli	33	15	3	51	47	3		152
Escherichia hermanii				2				2
Escherichia spp.	1			5	2			8
Ewingella americana		3						3
Flavimonas odoratum					1			1
Hafnia (Enterobacter) alvei		2		8				10
Hafnia alvei-doxy sensitive							1	1
Klebsiella oxytoca		2		4	1			7
Klebsiella pneumoniae		1		8	1			10
Klebsiella ozaenae				1				1
Kluyvera spp.		1						1
Leclercia adecarboxylata		2	1	1		2	1	7
Moellerella wisconsensis		1			10			11
Morganella morganii	2	3			2		1	8
Pantoea (Enterobacter) agglomerans		3	1	2	3	2		11
Pasteurella multocida					3			3
Pasteurella spp.					1			1
Pasteurella spp. (not P. multocida)					1			1
Photobacterium damsela	4	4						8
Plesiomonas shigelloides		1		2	1			4
Proteus mirabilis	1	3		16	5			25
Proteus penneri				1	1			2
Proteus vulgaris		1			1			2
Providencia rettgeri						2		2
Povidencia stuartii		1						1
Pseudomonas (flavimonas) oryzihabitans		1		1	1			3
Pseudomonas aeruginosa	1				1		3	5
Pseudomonas spp.	3	13	4	10	13	2	1	46
Pseudomonas stutzeri		1						1
Salmonella spp.				1				1
Serratia liquefaciens		1		11	4			16
Serratia marcesens		5		1		1		7
Serratia spp.				1				1
Shewanella algae					14			14
Shewanella spp.	1	16		1	10	2	2	32
Sphingomonas multivorium			1					1
Sphingomonas paucimobilis		2		1	1			4
Sphingomonas spp.				1				1
Stenotrophomonas maltophilia		2	1		1		1	5
Vibrio alginolyticus		2			8			10
Vibrio cholerae					1			1
Vibrio fluvialis		1						1
Vibrio parahaemolyticus		2		8				10
Vibrio spp.		3				2		5
Yersinia ruckeri				1				1
Aerobic Gram-negative Coccobacilli
Campylobacter spp.				1				1
Aerobic Gram-positive Bacilli
Bacillus spp.	1	5		2	3			11
Corynebacterium aquaticum		1						1
Corynebacterium spp.		4	1	6	3			14
Aerobic Gram-positive Cocci
Enterococcus avium				1				1
Enterococcus faecalis				3				3
Enterococcus spp.	3	9	1	15	14	1		43
Staphlococcus coagulase positive		2	2		5			9
Staphlococcus spp.		2		1				3
Staphlococcus-hemolytic	1							1
Staphlycoccus coagulase negative-Non hemolytic		5	4	4	3			16
Staphylococcus-Non-hemolytic				7				7
Streptococcus-alpha		3	2	3	9			17
Streptococcus-beta hemolytic		2	4					6
Streptococcus-gamma				10				10
Anaerobic Gram-positive Bacilli
Actinomyces spp.	1							1
Brevibacterium spp.		1					1	2
Clostridium bifermentans	1	1						2
Clostridium perfringens	8	7		4	12			31
Clostridium spp.	8	3			1			12
Propioibacterium acne		1						1
Anaerobic Gram-negative Baccili
Bacteroides spp.	2	1		1	2			6
Anaerobic Gram-positive cocci
Peptostreptococcus spp.		3						3
Total	84	173	41	237	223	20	19	797

Appendix 2.

Diversity and number of bacterial and fungal isolates cultured in routine sites from live, stranded and by-caught marine vertebrates. N = no. of individuals sampled at each routine swab site.

Microorganism	Fecal/ cloaca	Coelom	Abdomen	Thorax	Oral/nares/blow	Total
	N = 129	N = 35	N = 42	N = 38	N = 52	N
Aerobic Gram-negative Bacilli
Achromobacter (alcaligens) xylosoxidans subsp. xylosoxidans	0	0	0	0	1	1
Acinetobacter calcoaceticus-baumannii	0	3	1	2	2	8
Acinetobacter spp.	4	0	4	3	4	15
Aeromonas hydrophilia	2	0	0	0	4	6
Aeromonas spp.	2	2	0	0	7	11
Alcaligenes faecalis	1	0	0	0	0	1
Burkholderia cepacia	3	0	0	1	3	7
Chryseobacterium indologenes	0	0	0	2	0	2
Chromobacterium spp.	0	0	0	0	1	1
Citrobacter braakii	5	0	0	0	0	5
Citrobacter freundii	3	0	0	0	6	9
Citrobacter koseri	0	0	0	0	1	1
Citrobacter spp.	1	0	0	0	0	1
Edwardsiella hoshinae	1	0	0	1	0	2
Edwardsiella spp.	3	1	0	0	0	4
Edwardsiella tarda	5	0	3	3	1	12
Enterobacter amnigenus	3	0	0	0	0	3
Enterobacter avian	1	0	0	0	0	1
Enterobacter cancerigenus	0	0	0	0	1	1
Enterobacter cloacae	12	1	0	2	6	21
Enterobacter intermedius	1	0	0	0	0	1
Enterobacter spp.	5	2	0	2	6	15
Escherichia coli	107	4	3	8	20	142
Escherichia hermanii	2	0	0	0	0	2
Escherichia spp.	5	2	0	0	0	7
Ewingella americana	0	0	0	0	2	2
Flavimonas odoratum	0	0	0	0	1	1
Hafnia alvei	7	0	1	0	4	12
Klebsiella oxytoca	3	0	0	0	1	4
Klebsiella ozaenoe	1	0	0	0	0	1
Klebsiella pneumoniae	6	1	0	1	5	13
Leclercia adecarboxylata	1	2	0	1	0	4
Moellerella wisconsensis	2	1	0	0	5	8
Morganella morganii	0	0	0	0	1	1
Pantoea agglomerans	3	2	0	2	2	9
Pasteurella multocida	0	0	0	0	3	3
Pasteurella spp. (not multocida)	0	0	0	0	1	1
Photobacterium damselae	4	0	1	2		7
Plesiomonas shigelloides	2	0	0	0	1	3
Proteus mirabilis	6	0	2	2	8	18
Proteus penneri	0	0	0	0	2	2
Proteus vulgaris	1	0	0	0	2	3
Providencia rettgeri	2	0	0	0	0	2
Pseudomonas aeruginosa	2	0	0	0	0	2
Pseudomonas (flavimonas) oryzihabitans	0	0	0	0	1	1
Pseudomonas spp.	17	9	2	5	14	47
Pseudomonas stutzeri	0	0	0	0	1	1
Salmonella spp.	0	0	0	0	1	1
Serratia liquefaciens	3	0	0	0	9	12
Serratia marcesens	1	0	1	2	1	5
Serratia spp.	0	0	0	0	1	1
Shewanella algae	3	0	0	0	6	9
Shewanella spp.	7	4	1	3	7	22
Sphingomonas multivorium	0	0	0	1	0	1
Sphingomonas paucimobilis	1	0	1	0	2	4
Stenotrophomonas maltophilia	0	1	0	2	0	3
Vibrio alginolyticus	6	1	0	0	9	16
Vibrio cholerae	0	0	0	0	1	1
Vibrio parahaemolyticus	3	2	0	0	1	6
Vibrio spp.	0	0	2	2	1	5
Yersinia ruckeri	0	1	0	0	0	1
Aerobic Gram-negative Coccobacilli
Campylobacter spp.	2	0	0	0	0	2
Aerobic Gram-positive Bacilli
Bacillus spp.	2	0	1	0	5	8
Corynebacterium aquaticum	0	0	0	1	0	1
Corynebacterium spp.	4	0	3	0	4	11
Aerobic Gram-positive Cocci
Enterococcus avian	1	0	0	0	0	1
Enterococcus faecalis	3	0	0	0	0	3
Enterococcus spp.	29	3	4	5	12	53
Staphlococcus coagulase positive	3	1	1	1	0	6
Staphlococcus-hemolytic	1	0	0	0	0	1
Staphylococcus-Non-hemolytic	0	0	0	0	7	7
Staphlycoccus-non-hemolytic coagulase negative	1	1	5	3	1	11
Streptococcus- gamma	1	0	0	0	9	10
Streptococcus-alpha	4	0	1	1	7	13
Streptococcus-beta hemolytic	0	0	3	2	0	5
Anaerobic Gram-positive Bacilli
Actinomyces spp.	0	1	0	1	0	2
Clostridium bifermens	1	0	1	0	0	2
Clostridium perfringens	17	1	1	1	4	24
Clostridium spp.	8	0	0	2	0	10
Propioibacterium acnes	0	0	2	0	0	2
Anaerobic Gram-negative Baccili
Bacteroides spp.	3	0	0	0	2	5
Anaerobic Gram-positive Cocci
Peptostreptococcus spp.	0	0	1	0	1	2
Fungi
Aspergillus spp.	0	1	0	1	0	2
Total	327	47	45	65	208	692
Total per swab	2.5	1.3	1.1	1.7	4

Appendix 3.

Number (N) of animals (see Table 1) where microbial organisms were isolated, source of isolate, pathology findings in animals isolated and nature of pathology when isolated in human patients.

Microorganism	N	Host(s) and notes	Human pathogen?	Associated pathology or other findings (host species) in this study	Nature of pathogen in humans
Escherichia coli	122	152 isolates, with multiple resistances in piping plover, common eider, common loon (0–5 ABR), live herring gull (0–4 ABR), great black-backed gull (0–6 ABR) and seal (0–3 ABR)	Yes (ABSA)		Shiga toxin-producing Escherichia coli (STEC): hemorrhagic colitis, intestinal disease, cramps, abdominal pain, low grade fever
Pseudomonas spp.	42	52 isolates from all taxa and species. Most multiply resistant to AUG, AMP, CAR, CEF, CEPH, TIC	Yes (ABSA)	Often a secondary infection	Associated with bacterial meningitis, abscesses, endocarditis, pneumonia
Clostridium perfringens	21	27 isolates Most common in common eiders (10), Cuvier's beaked whale, live seals (8), live gulls (2), and stranded Atlantic white-sided (1) and common dolphin (4)	Yes (ABSA)	Encephalitis (Cuvier's beaked whale) and bacterial septicemia and enteritis (eider)	Necrotic enteritis, food poisoning, soft tissue infection
Enterobacter cloacae	22	26 isolates in all taxa. AUG, CEPH and TIC common resistance in many isolates. Found in stranded minke whale (1), hooded seal (1), and stranded common eider (3) and piping plover (1), sharks (3) live herring gulls (11), by-caught harbor seal (1) and harp seal (1)	Yes (ABSA)	Mostly in cloacal samples. Acanthocephalan enteritis in many eiders	Bacteremia, endocarditis, osteomyelitis, lower respiratory tract, skin, urinary tract, and ophthalmic infections
Enterobacter spp.	20	20 isolates in live gulls, stranded eiders, 5 species of stranded cetacean, and 1 by-caught grey seal. Many resistant	Yes (ABSA)		Associated with surgical wound infections
Shewanella spp.	18	32 isolates in all taxa and many tissue sources. AUG, AMP, CAR, CEF resistance common in isolates from Atlantic white-sided dolphin (1), hooded seal (1), pilot whale (1), striped dolphin (1), unknown delphinid (1) and minke whale (2). Also in common eiders (5), thresher shark (2), by-caught shearwater (1), red breasted cormorant (1) and live grey seal and herring gull	Yes (ABSA)	Secondary bacterial infections	Cellulitis, otitis media, ocular infections, endocarditis, abcesses, osteomyelitis, peritonitis and septicemia
Proteus mirabilis	17	33 isolates in gulls (19), eiders (1), humpback (1), one common dolphin (1) and minke whale (1). Many bird isolates TET resistant. Herring gull and eider isolates had multiple resistance to AUG, AMP, CAR and CEF. Minke whale isolate also resistant to TIC	Yes (ABSA)	Lung abcess in minke whale	Urinary tract infection, infections, septicemia and pneumonia
Staphlycoccus (non-hemolytic, coagulase-negative)	18	18 isolates in 2 stranded and 3 by-caught harp seals, live herring gulls (5), by-caught porpoise (1), stranded unknown dolphin species (1), minke whale (1), Risso's dolphin (1), striped dolphin (1) and eiders (3). Eiders and striped dolphin isolates resistant to AMP and PENG
Serratia liquefaciens	16	16 isolates in 11 live herring gulls, stranded common eiders (4) and Atlantic white-sided dolphin (1)	Yes (Grohskopf et al. 2001)		Bloodstream infection, bacteremia
Streptococcus- alpha	16	16 isolates in a by-caught Atlantic white-sided dolphin and harp seal, 7 eiders from one die-off event, live herring gulls (3), live grey seal (1) and stranded common dolphin (1), harp seal (1) and harbor porpoise (1)	Yes (ABSA)	Bacterial septicemia and enteritis (eiders). Secondary infection (eiders)	Bacterial pneumonia, meningitis
Vibrio alginolyticus	15	24 isolates, most common in eider (9). Also in 1: minke whale, pygmy sperm whale, unknown delphinid, cormorant and live laughing gull and grey seal. All isolates resistant to AMP and CAR, eider isolates resistant to TIC	Yes (Schmidt et al. 1979)	Enteritis and endocarditis (pygmy sperm whale) and associated with verminous gastritis and enteritis in many eiders	Necrotizing fasciitis, gastroenteritis, septicemia
Acinetobacter spp.	14	17 isolates in 7 species of live, stranded and by-caught marine mammals, and common eider. Pygmy sperm whale isolate multiply resistant, eider isolates show no resistance	Yes (ABSA)		Lower respiratory and urinary tract infecions
Corynebacterium spp.	13	14 isolates in stranded Atlantic white- sided dolphin, Cuvier's beaked whale, harbor porpoise,humpback whale, live (2) and stranded (1) herring gull, stranded great black-backed gull (1), and common eider (1)	Yes (ABSA)	Endocardiosis and arteriosclerosis, and thoracic fluid in the thoracic cavity with associated inflammation of the prescapular and thoracic lymph nodes of a dolphin	Endocarditis
Bacillus spp.	13	In live herring gulls (2), live harbor seal (1), stranded herring gull (1), common eiders (3), 6 marine mammal species (stranded pygmy sperm whale, humpback whale, beaked whale, Risso's dolphin, and porpoise)	No (ABSA)
Clostridium spp.	12	Most common in live seals (8). Also in stranded animals: 1 common eider, 1 pilot whale, 1 unknown cetacean and 1 hooded seal	Yes (ABSA)	Associated with large amount of fluid in thoracic cavity of unidentified delphinid species	Gas gangrene of wounds, necrotizing enterocolitis in infants
Edwardsiella tarda	12	In live herring gull (1), stranded white sided dolphins (2), stranded common dolphin (1), live seal (4), pilot whale stranded (1), stranded harp seal (2) and humpback whale (1). Only 2 live grey seal isolates resistant to AMP, CARB and one to CEF	Yes (ABSA)	Various organs from a pilot whale with multisystem infection and enteritis. Also in a humpback whale with enteritis	Gastroenteritis, cellulitis, peritonitis, septicemia
Acinetobacter calcoaceticus-baumannii	12	11 isolates, all CAZ, AMP, CHLO, and CEPH-resistant, isolated from a stranded beaked whale, by-caught harbor porpoise, harbor seal and grey seal, stranded eiders (4), stranded pilot whale (1) and thresher shark (1)	Yes (ABSA)		Nosocomial: bacteremia, septicemia, pneumonia
Moellerella wisconsensis	11	11 animals and 11 isolates.AMP and CAR resistance only. Found in eiders (all part of mass die-offs) and 1 pygmy sperm whale (also resistant)	Yes (Quevedo et al. 2006)	Eiders all part of mass die-off. Histologic changes are consistent with emaciation, bacterial septicemia, bacterial enteritis, and intestinal trematodiasis. Bacteria are small bacilli	Abdominal pain, fever, cramps, and diarrhea
Pantoea agglomerans	11	Many resistant or intermediate to CEPH: stranded eider (1), stranded common dolphin (2), stranded minke whale (1) and by-caught shearwater (1), harbor porpoise (1) and cormorant (1)	Yes (Cruz et al. 2007)		Nosocomial pathogen wound and urinary tract infections, septic arthritis
Citrobacter freundii (and complex)	11	AUG resistance common. 11 animals with 2 thresher sharks, 4 herring gulls live, 2 stranded birds, 2 stranded dolphins and 1 live seal	Yes (ABSA)		Neonatal meningitis
Shewanella algae	11	Only in common eiders from Wellfleet, MA. CEPH resistance common, 2 eiders have multiple ABR (0-4).)	Yes (ABSA)	Impacted cloaca in (eider)	Bacteremia and ear infections, osteomyelitis
Klebsiella pneumoniae	10	8 live herring gulls, stranded eider and stranded common dolphin. All resistant to CAR and TIC, most to AMP	Yes (ABSA)	Collapsed lung (dolphin) chronic enteritis interstitial pneumonia (eider)	Pneumonia, septicemia, wound/burn infection, cystitis, ankylosing spondylitis
Escherichia spp.	10	7 live and 1 stranded herring gull, live seal, and stranded eider with various ABR	Yes (ABSA)		Gastroenteritis
Streptococcus- gamma	10	9 live herring gulls, 1 stranded great black-backed gull
Aeromonas hydrophila	9	1 harbor seal, 7 eiders, 1 greater black back gull with multiple resistance, primarily resistant to CEPH, AUG, AMP and CARB	Yes (ABSA)	Immuno-compromised neonate harbor seal with bacterial pneumonia	Septicemia, ocular, respiratory tract infections, pneumonia and urinary tract infections
Vibrio parahaemolyticus	9	Most common in eiders (7), one hooded seal and a striped dolphin. Resistance to AMP and CAR common, but not for 2 eiders	Yes (ABSA)	Peritonitis, septicemia, lymphocitic inflammation, enterotoxemia, splenic and hepatic necrosis in eiders. Associated with fluid and lung lesions in striped dolphin	Gastroenteritis, wound infection
Staphlococcus-coagulase positive	9	Only ABR to ENR and GEN in 1 by-caught harp seal. Present in harp seals (by-caught and stranded) (2), grey seal by-caught (1), striped dolphin (1) with mutiple resistance and stranded eiders (5)
Aeromonas spp.	8	13 isolates present in 3 stranded cetaceans (white sided dolphin, minke whale and common dolphin), by-caught shearwater and red throated loon, live herring gulls (2), and stranded eider (1). AMP, CARB, CEPH primarily resistant	Yes (ABSA)	Pancreatic fluid (minke whale) and coelom (eider). Pericardial fluid associated with Aspergillus (eider)	Gastroenteritis (variable)
Hafnia (Enterobacter) alvei	8	8 animals, all resistant to CEPH and AUG: thresher shark (1), live herring gulls (5), stranded harbor seal (1), and stranded unknown cetacean (1)	Yes (ABSA)	Abdomen of immunosuppressed neonate harbor seal	Diarrhea
Morganella morganii	8	All isolates AUG- and CEPH-resistant except one harp seal isolate. Many TET-resistant. All taxa: common eider (2), pilot whale (1), thresher shark (1), Risso's dolphin (1), harp seal (1) and live seals (2)	Yes (ABSA)	Testicular abcess in pilot whale and genital discharge and inflammmatory cells in Risso's dolphin	Nosocomial: bacteremia, urinary tract, pneumonia, wounds musculoskeletal, central nervous system, pericarditis, and spontaneous bacterial peritonitis
Photobacterium damselae	8	11 isolates: stranded minke whale (1), white sided dolphin (1), striped dolphin (1), beaked whale (1), and live seals (4). CARB, AMP, TIM resistance primarily	Yes (ABSA)	Isolated in abdominal fluid of a 24 h post mortem Atlantic white sided dolphin with marked pancreas autolysis (and no other pathology). Also in urine of Cuvier's beaked whale with cystitis	Fish pasteurellosis, necrotizing fasciitis
Streptococcus-beta hemolytic	6	Only pinnipeds represented: (2) stranded and (4) by-caught, harbor, grey and harp seals	Yes (Henkel 1970)	Isolated in multiple sites of one neonate harbor seal with suppurative bacterial bronchopneumonia and one harp seal with terminal bacterial sepsis	Bacteremia, endocarditis, cellulitis, wound infection
Burkholderia cepacia	6	Isolated in live gulls (4), live grey seal (1), stranded common dolphin (1):all with multiple resistance	Yes (ABSA)	Pulmonary edema (common dolphin)	Pneumonia
Serratia marcesens	6	Mostly marine mammals: common dolphin (1), white sided dolphin (1), hooded seal (1) and harbor seal (1). Only one stranded common dolphin with no resistance. Birds: gannet (1) and cormorant (1). Resistance to CEPH common	Yes (ABSA)	Bacterial suppurative bronchopneumonia (neonate harbor seal), Chronic lymphandenopathy (common dolphin)	Opportunistic infections of the endocardium, eyes, blood, wounds, urinary and respiratory tracts
Citrobacter braakii	6	All resistant to AUG and some to CEPH with6 animals represented: thresher shark (1), herring gulls live (2), by-caught (1) and live grey seals (2).	Yes (ABSA)		Cellulitis, peritonitis
Leclercia adecarboxylata	6	6 animals total: stranded pygmy sperm whale (1), and harbor porpoise (1), by-caught shearwater (1), loon (1) and harbor seal (1) andone thresher shark (1) No resistance	Yes (ABSA)		Rare bacteremia
Bacteroides spp.	5	One stranded and live (hooded and grey) seals (2), stranded eider (1) and live herring gull (1).	Yes (ABSA)	Associated with imflammed perigastric lymph and abdominal cavity of a hooded seal	Diarrhea, bacteremia
Klebsiella oxytoca	5	Five animals, stranded and live, all resistant to CAR, all but 3 to TIC and AMP: (1) stranded hooded seal and common dolphin, (1) stranded herring gull, (2) live herring gulls	Yes (ABSA)	Marked proliferative and ulcerative dermatitis, bacterial with underlying viral condition (dolphin)	Urinary and pulmonary infections; wound infections; secondary infection in lungs of patients with chronic pulmonary disease; enteric pathogenicity (enterotoxin); ozena (atrophy of nasal mucosa) and rhinoscleroma
Vibrio spp.	5	Found in 4 stranded marine mammals and one by-caught cormorant. Includes a pygmy sperm whale, a hooded seal, 2 harp seals. AMP resistance common, with the pygmy sperm whale with highest number of antibiotic resistance (4)	Yes (ABSA)	Enteritis (harp seal); verminous gastritis (harp seal)	Gastroenteritis
Stenotrophomonas maltophilia	5	One by-caught harbor porpoise, stranded pygmy sperm whale, risso's dolphin, stranded plover and a thresher shark	Yes (ABSA)		Nosocomial, very low risk, aquatic envrioment, urinary tract infections
Aspergillus spp.	4	Three stranded eiders, one by-caught Atlantic white sided dolphin	Yes (ABSA)	Fungal granulomae and necrosis on serosa of coelom, viscera, and air sacs (eiders)	Pulmonary; invasive aspergillosis
Pasteurella multocida	4	Four common eiders	Yes (ABSA)	Eiders from one large die-off event	Enteritis, bacteremia, septicemia, respiratory pathogen, endocarditis, osteomyelitis, meningitis and brain abcess
Plesiomonas shigelloides	4	One stranded eider, 2 live herring gulls, 1 stranded striped dolphin, all with 3+ ABR	Yes (ABSA)	Thoracic swab of striped dolphin with pneumonia	Gastroenteritis, occurs mainly in tropical and subtropical areas
Sphingomonas paucimobilis	4	One stranded common loon, pygmy sperm whale and harbor porpoise and live herring gull. Pygmy sperm whale AMK and GEN resistant	Yes (ABSA)	Abdomen of pygmy sperm whale with enteritis	Bacteremia
Pseudomonas aeruginosa	4	One thresher, (2) mako shark, (1) stranded common eider and (1) live grey seal. Multiple resistance, all resistant to AUG, AMP, CEF, CEPH, CHL, and TET. Mako 10 ABR including GEN	Yes (ABSA)		Nosocomial: urinary tract infections, respiratory system infections, dermatitis, soft tissue infections, bacteremia, bone and joint infections, gastrointestinal infections
Ewingella americana	3	CEPH resistant, only found in dolphins: 2 common dolphin and one unknown species	Yes (ABSA)	Ascending infection from the urinary bladder (dolphin); interstitial pneumonia and congestion (dolphin)	Nosocomial: septicemia, peritonitis, pneumonia, bacteremia
Peptostrepto-coccus spp.	3	Two stranded common dolphins and (1) minke whale	Yes (ABSA)	Suppurative peritonitis, dermal abcess,perithoracic duct granulomas, peritonitis, hepatitic necrosis, and chronic dermal abscessation (common dolphin). Peritonitis, pulmonary edema,interstitial pneumoniaand plasmacytic enteritis in a common dolphin. Lung abscess in chronically entangled minke whale	Brain abcess, skin infection, upper respiratory and dental infections, peritonitis, liver and spleen abcess, osteomyeltis, arthritis, fatal endocarditis
Pseudomonas (flavimonas) oryzihabitans	3	One stranded beaked whale,eider, and live herring gull: no resistance	Yes (ABSA)	Isolated in urine of beaked whale	Nosocomial: peritonitis
Enterococcus faecalis	3	Birds only: no resistance in one stranded common loon, common eider and northern gannet	Yes (ABSA)		Endocarditis, as well as bladder, prostate, and epididymal infections, endocarditis
Proteus vulgaris	3	Minke whale, gull (8 ABR) and eider (3 ABR)	Yes (ABSA)		Urinary tract infection, urinary calculi
Providencia rettgeri	3	One stranded pygmy sperm whale and 2 by-caught shearwaters. All TET, CHL resisant with multiple resistance, 7 ABR in pygmy sperm whale isolate	Yes (ABSA)		Invasive diarrhea, bacteremia, ocular infection
Brevibacterium spp.	2	One pygmy sperm whale and one thresher shark	Yes (ABSA)	White plaque build up in sinuses and mandible of pygmy sperm whale	Malodor, otitis, endocarditis
Chryseobacterium indologenes	2	Two by-caught seals; harbor and harp with resistance to 11 ABR	Yes (ABSA)	Acute pulmonary congestion in 2 seals	Nosocomial bacteremia
Clostridium bifermentans	2	One minke whale and one scat from a live seal	Yes (ABSA)		Food poisoning, bacteremia, metastatic osteomyelitis
Enteroccocus spp.	2	Two live herring gulls	Yes (ABSA)		Surgical wound infections, bacteremia, subacute endocarditis
Escherichia hermannii	2	One black-backed and herring gull (AUG, AMP, CEPH resistant), live	Yes (ABSA)
Proteus penneri	2	One live herring gull (6 ABR) and stranded eider (3 ABR)	Yes (ABSA)		Chronic urinary tract infections, bacteremia, pneumonia, neonatal meningoencephalitis, empyema, osteomyelitis, cystitis, pyelonephritis, prostatitis
Achromobacter (alcaligenes) xylosoxidans ss xylosoxidans	1	One eider sampled in October 2006 die-off event	Yes (ABSA)		Bacteremia, meningitis, pneumonia, endocardidtis
Actinomyces spp.	1	One live seal	Yes (ABSA)		Actinomycosis- oral and cervicofacial swelling with suppuration, abscess formation, tissue fibrosis, and draining sinuses
Alcaligenes faecalis (Bordatella avium)	1	One eider AMP and CHL resistant	Yes (Bizet & Bizet 1997)		Endocarditis, meningitis, otitis, hepatitis and diarrhea
Actinomyces spp.	1	One live seal	Yes (ABSA)		Actinomycosis- oral and cervicofacial swelling with suppuration, abscess formation, tissue fibrosis, and draining sinuses
Candida glabrata	1	One stranded Cuvier's beaked whale	Yes (ABSA)	Green fluid in respiratory system	Candidiasis, mucosal/ oral/ esophageal
Chromobacterium violaceum	1	One stranded white sided dolphin	Yes (ABSA)		Cutaneous inflammation, sepsis, diarrhea, liver abscesses and ocular infections
Corynebacterium aquaticum	1	One common dolphin	Yes (Moore and Norton 1995)	Associated to thorax of dolphin with multisystem pathologies proliferative dermatitis, sepsis, atherosclerosis, and degenerative cardiac disease	Septicemia, urinary tract infection, meningitis
Enterococcus avium	1	One stranded loon, AMP, VANC resistant	Yes (ABSA)		Meningioencephalitis, brain abcess, bacteremia, endocardititis, and osteomyelitis
Empedobacter brevis	1	One eider, no resistance	Yes (ABSA)		Nosocomial: endophthalmitis
Enterobacter cancerogenus	1	One eider stranded. Oct 2006 die-off. AUG, AMP, CEPH resistant	Yes (ABSA)		Nosocomial: bacteremia, wound infections, osteomyelitis
Enterobacter sakazakii	1	One by-caught harbor porpoise uterus	Yes (ABSA)	Pregnant near full term fetus (porpoise)	Neonatal sepsis, meningitis, or necrotizing enterocolitis
E. coli sorbitol negative	1	One stranded common eider	Yes (ABSA)		diarrhea, abdominal cramps
Klebsiella ozaenae	1	One live herring gull, no resistance	Yes (ABSA)		Atrophic rhinitis,nasal congestion, headache, sinusitis
Kluyvera spp.	1	One stranded white sided dolphin with AMP, CARB, CEPH, TIC resistant	Yes (ABSA)	Genital region with renal dysfunction and peritoneal serositis (dolphin)	Pyelonephritis, bacteremia, acute appendicitis
Salmonella spp.	1	One live herring gull	Yes (ABSA)		Nausea, vomiting, abdominal cramps, diarrhea, fever, and headache
Serratia spp.	1	One live herring gull, AUG and CEPH resistant	Yes (ABSA)		Opportunistic infections of the endocardium, eyes, blood, wounds, urinary and respiratory tracts
Sphingobacterium multivorum	1	One by-caught harbor porpoise only. 8 ABR (AMK, AMP, CAR, CEF, CHL, GEN, TIC, and TOB)	Yes (ABSA)	Isolated in thorax of porpoise with verminous bronchopneumonia	Bacteremia
Streptococcus uberis (viridans strep)	1	One stranded common eider	Yes (ABSA)		Mastitis (in cattle)
Propionibacterium acnes	1	One stranded common dolphin	Yes (ABSA)	Chronic-active peritonitis (dolphin)	Endocardidtis, brain abscess, subdural empyema, dental infections, endocarditis, peritonitis
Pseudomonas stutzeri	1	One stranded harbor porpoise only	Yes (Goetz et al. 1983)	Blow/respiratory system of severely emaciated porpoise with moderate interstitial pneumonia and hemmorhage sepsis	Fever, shaking chills, nausea, and vomiting
Providencia stuartii	1	One stranded hooded seal and with multiple resistance	Yes (ABSA)		Bacteremia, urinary origin, endocarditis
Vibrio cholerae	1	One stranded herring gull, AMP resistance only	Yes (ABSA)		Cholera, gastroenteritis
Vibrio fluvialis	1	One pygmy sperm whale, 5 ABR (AUG,AMP, CAR, CEPH, TIC)	Yes (ABSA)	Genital slit of pregnant pygmy sperm whale with cervicitis and uterine edema but no inflammation	Gastroenteritis, diarrhea
Chromobacterium spp.	1	One thresher shark, multiple ABR	No (ABSA)
Edwardsiella hoshinae	2	No resistance present: (1) stranded harbor seal and (1) live seal	No (ABSA)
Yersinia ruckeri	1	One stranded eider, no resistance	No (ABSA)	Acanthocephalan enteritis (eider)
Flavimonas odoratum	1	One common eider (AB resistance to AMK, CAZ, CEF, GEN, TOB)	No (ABSA)

ABR: antibiotic resistance(s); AMP: ampicillin; AUG: augmentin; CAR: carbenicillin; CAZ: ceftazidime; CEF: ceftiofur; CEPH: cephalothin; CHL: cloramphenicol; CIP: ciprofloxacin; ERY: erythromycin; ENR: enrofloxacin; GEN: gentamycin; OX: oxacillin; PENG: penicillin G TET: tetracycline; TIC: ticarcillin; TOB: tobramycin; VAN: vancomycin. ABSA: American Biological Safety Association (www.absa.org/XriskgroupsX/index.html)

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