Little is known about microsporidiosis pathogenicity in cetaceans. Here we report seroprevalence of 76% for microsporidia in blood samples from common bottlenose dolphins (Tursiops truncatus), from animals managed under human care (n=108) or captured for health assessments (n=13) and released

Cray, Carolyn

Desoubeaux, Guillaume

Flower, Jennifer E.

Fravel, Venessa

Jensen, Eric D.

Joachim, Anja

Le-Bert, Carolina

Peschke, Roman

Soto, Jeny

Wells, Randall

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University of Nebraska - Lincoln DigitalCommons@University of Nebraska - Lincoln U.S. Navy Research U.S. Department of Defense 2018 Seroprevalence Survey for Microsporidia in Common Bottlenose Dolphin (Tursiops truncatus): Example of a Quantitative Approach Based on Immunoblotting Guillaume Desoubeaux Roman Peschke Carolina Le-Bert Venessa Fravel Jeny Soto See next page for additional authors Follow this and additional works at: https://digitalcommons.unl.edu/usnavyresearch This Article is brought to you for free and open access by the U.S. Department of Defense at DigitalCommons@University of Nebraska - Lincoln. It has been accepted for inclusion in U.S. Navy Research by an authorized administrator of DigitalCommons@University of Nebraska - Lincoln. Authors Guillaume Desoubeaux, Roman Peschke, Carolina Le-Bert, Venessa Fravel, Jeny Soto, Eric D. Jensen, Jennifer E. Flower, Randall Wells, Anja Joachim, and Carolyn Cray DOI: 10.7589/2017-11-287 Journal of Wildlife Diseases, 54(4), 2018, pp. 870–873 Wildlife Disease Association 2018Seroprevalence Survey for Microsporidia in Common BottlenoseDolphin (Tursiops truncatus): Example of a Quantitative ApproachBased on ImmunoblottingGuillaume Desoubeaux,1,2,3 Roman Peschke,4 Carolina Le-Bert,5 Vanessa Fravel,6 Jeny Soto,1 Eric D.Jensen,7 Jennifer E. Flower,8 Randall Wells,9 Anja Joachim,4 and Carolyn Cray1,10 1Division of ComparativePathology, Department of Pathology & Laboratory Medicine, Miller School of Medicine, University of Miami, 1600 NW10th Avenue #7101A, Miami, Florida 33136, USA; 2Service de Parasitologie–Mycologie–Médecine Tropicale, CHU deTours, 2 Boulevard Tonnellé, Tours 37044, France; 3CEPR–INSERM U1100/Équipe 3, Faculté de Médecine, UniversitéFrançois-Rabelais, 10 Boulevard Tonnellé, Tours 37032, France; 4Institute of Parasitology, Department of Pathobiology,University of Veterinary Medicine Vienna, Veterinärpl. 1, Vienna 1210, Austria; 5National Marine Mammal Foundation,2240 Shelter Island Drive, Suite 200, San Diego, California 92106, USA; 6Six Flags Discovery Kingdom, 1001Fairgrounds Drive, Vallejo, California 94589, USA; 7US Navy Marine Mammal Program, Space and Naval WarfareSystems Center Pacific, 53560 Hull Street, San Diego, California 92152, USA; 8Mystic Aquarium, a division of SeaResearch Foundation Inc., 55 Coogan Boulevard, Mystic, Connecticut 06355, USA; 9Chicago Zoological Society’sSarasota Dolphin Research Program, c/o Mote Marine Laboratory, 1600 Ken Thompson Parkway, Sarasota, Florida34236, USA; 10Corresponding author (email: ccray@med.miami.edu)ABSTRACT: Little is known about microsporidiosispathogenicity in cetaceans. Here we reportseroprevalence of 76% for microsporidia in bloodsamples from common bottlenose dolphins (Tur-siops truncatus), from animals managed underhuman care (n¼108) or captured for healthassessments (n¼13) and released.Microsporidiosis is an opportunistic infec-tion caused by microsporidia, which areunicellular eukaryotes related to fungi (Mathiset al. 2005; Didier and Weiss 2006). It occursin immunocompromised patients and is orallytransmitted by spores contained in excreta of awide range of host species (Wasson and Peper2000; Didier and Weiss 2006). Fourteenmicrosporidian species have been describedin humans, and some are zoonotic pathogens,such as Encephalitozoon cuniculi of rabbits,dogs, and rodents (Wasson and Peper 2000),and Enterocytozoon bieneusi of cats, pigs, andcattle (Mathis et al. 2005). However, very littleis known about microsporidia infection insome other animal clades, such as cetaceans.Collection of excreta is difficult in aquaticfauna and even more so in wild fauna. Directmethods of detection do not reflect the actuallevels of exposure, because spore shedding isknown to be transient (Didier and Weiss2006). Presence of spores in excreta can alsoreflect a simple gastrointestinal passage ofundigested spores, because microsporidia arereadily carried by prey fish, such as Atlanticsalmon (Salmo salar), round goby (Neogobiusmelanostomus), turbot (Scophthalmus maxi-mus), and peacock wrasse (Symphodus tinca)for which up to 43% are positive (Fayer et al.2008; Kvach and Winkler 2011). Thus, 20.5%stools from naturally exposed dolphins arepositive by PCR at the time of sampling(Fayer et al. 2008). To indicate globalexposure, serological assays for prevalenceare considered a useful screening approachthat can partly overcome the aforementionedlimitations.We obtained blood samples from commonbottlenose dolphins (Tursiops truncatus), ei-ther managed under human care (n¼108;male¼50.5%; mean age¼2165 yr) or free-ranging (n¼13; male¼35.7%; mean age¼1263yr). Plasma specimens were tested at theUniversity of Miami for antimicrosporidialantibodies by means of an inhouse quantita-tive immunoblot (Desoubeaux et al. 2017b),using E. cuniculi extract as protein antigens(Künzel et al. 2014), and horseradish perox-idase-conjugated rabbit antibottlenose dol-phin IgGhþl (Bethyl, Montgomery, Texas,USA). A global score was calculated as thesum of all the band intensities, estimated bytwo independent optical assessments, whichranged from 0 to 4 for each of the sevenproteins located at 135, 75, 50, 40, 30, 25, and19–20 kDa on the immunoblot strip. Thus, thetotal was ranked on 28 points as a maximum,870and the positive cutoff value set at 5/28. In all,75.9% samples were positive (Fig. 1). Thescores ranged from 0 to 23/28, the median was8, and the mean value 9.163.0. The twoproteins located at 135 kDa and 30 kDa,corresponding to polar tube protein-3 (Uni-Prott accession no. Q8MTP3) and polar tubeprotein-2 (Q8SRT0), showed the highestimmunoreactivity, with 71.9% and 70.2%positive sera. Specificity of the immunereaction seemed unquestionable, becausetheoretically, E. cuniculi polar tube protein-3has peptide sequence similarity only to relatedproteins of other Encephalitozoon species,such as Encephalitozoon hellem, Encephalito-zoon intestinalis, or Encephalitozoon romaleae(identity-threshold .20%; UniProt BLAST-score .450 with 500 hits allowed; UniProtexpected value 1.031047; https://www.uniprot.org/blast/). Encephalitozoon cuniculipolar tube protein-3 does not share similari-ties to fungi, bacteria, or archaea. In addition,FIGURE 1. Microsporidial antigen immunoblotting of blood samples from common bottlenose dolphin(Tursiops truncatus). Examples of three plasma specimens showing negative () and positive (þ) patterns usingprotein extract from Encephalitozoon cuniculi CH-K 2169 strain (Desoubeaux et al. 2017b) and horseradishperoxidase-conjugated rabbit antibottlenose dolphin IgGhþl conjugate. The primary bands are highlighted by thetheoretical molecular weight of the corresponding proteins indicated in kilodaltons (kDa), on the right. In rabbitsnaturally-infected with E. cuniculi, the polar tube protein 3 (UniProt accession no. Q8MTP3) located at 135 kDawas shown to be the most sensitive, whereas polar tube protein 2 (Q8SRT0) at 30 kDa was found to bestatistically associated with the report of current clinical signs (Desoubeaux et al. 2017a). Unlike proteins locatedat higher molecular weights, immunoreactivity with proteins located at 25 and 19–20 kDa was weak (i.e., positiveherein for only 7.9% and 2.6% samples, respectively). WB¼western blot.LETTERS 871E. cuniculi polar tube protein-2 shares simi-larity only with Encephalitozoon spp., Vitta-forma corneae, Nosema spp., and E. bieneusi(identity-threshold .20%; UniProt BLAST-score .125 with 500 hits allowed; UniProtexpected value 8.43105). There was statis-tical impact of dolphin sex (odds ratio¼0.50,95% confidence interval¼0.25–0.99; P¼0.05),but neither of age (P¼0.91) nor geographicalorigin and lifestyle (captive or wild; P¼0.37)upon the overall seroprevalence (using ageneral linear statistical model of analysis ofcovariance for measuring association betweenvariables and the immune status (Wu 2018).Associations with the intensity of globalantibody score were not significant for allthe variables (P¼0.55; 0.87 and 0.94, respec-tively)The hypothesis of an origin of microsporid-ia contamination by marine species is unlikely:among the various microsporidial species ofthe fishes eaten by dolphins, none is describedas zoonotic. Most microsporidia belong to theclade 5 of Marinosporidia, such as Pleisto-phora, Tetramicra, Microgemma, and Kaba-tana genera (Fayer et al. 2008). Thus,environmental contamination with spores ofspecies that come from terrestrial warm-blooded animals is more likely. A similarland-to-sea transmission scheme was demon-strated for Toxoplasma gondii, another im-portant pathogenic parasite in various marinemammals (Cooper et al. 2016), where oocystsare excreted by felids, primarily domestic cats(Felis catus), and flushed into coastal waterswhere they can be ingested directly bysusceptible hosts or are concentrated intofiltering shellfish which are thereafter eaten(Długońska 2017). Because Toxoplasma sero-prevalence is high in dolphins, oocysts havebeen assumed to get massively collected in thecoastal environment through sewage andsurface water runoff. For microsporidia,75.9% seroprevalence suggested a high levelof exposure. Microsporidial spores have beendetected in estuaries that collect runoff waterscontaminated with feces from terrestrialanimals or humans, leaky septic tanks, orwastewater treatment plants, especially afterevents such as rainfall and sewage overflows.Likewise, spores have been found in recrea-tional beach waters, mostly observed duringweekends and after increased human activities(Graczyk et al. 2010). Infected humans canexcrete millions of microsporidia spores daily;a median of 5.03107 spores/mL was describedin human immunodeficiency virus-positivepatient stools (Goodgame et al. 1999), andmicrosporidia prevalence was found to be upto 60% in some of domestic terrestrialmammal species (Li et al. 2014).Overall, our serology results indicated amassive exposure to microsporidia in theexamined dolphins, but was not able toindicate if the cetaceans are dead-end hostsor if they contribute to the dissemination ofthe pathogens by excretion. If the latter istrue, one could imagine that spores can bedispersed into recreational water for swim-ming, because dolphins live in coastal waters.Spores could also be readily concentrated infiltering shellfish, such as mussels used forhuman consumption or as food for othermarine mammals (Ghozzi et al. 2017). Thepathogenicity of microsporidia in sea mam-mals is unknown. Around 45% of the dolphinswe studied had undergone routine biopsy ornecropsy (for alternative diagnostic investiga-tions in light of clinical signs involving thebowel or the lungs), but no specific histopath-ological lesions were reported. Neither spe-cific staining nor immunohistochemistry wereused to confirm the presence or absence ofspores in tissue. Additional studies are nowneeded in order to determine the in vivopathological consequences of microsporidiafor wildlife health.We conducted our study using archivedsamples collected from Six Flags Aquariumand US Navy marine mammals as part of theirroutine care. The US Navy Marine MammalProgram houses and cares for a population ofdolphins in San Diego Bay (California, USA).The Marine Mammal Program is accreditedby the Association for Assessment and Ac-creditation of Laboratory Animal Care andadheres to the national standards of the USPublic Health Service Policy on the HumaneCare and Use of Laboratory Animals and theAnimal Welfare Act. Samples obtained from872 JOURNAL OF WILDLIFE DISEASES, VOL. 54, NO. 4, OCTOBER 2018free ranging dolphins in Sarasota Bay (Flor-ida, USA) were collected under NationalMarine Fisheries Service Scientific ResearchPermit 15543 and Mote Marine LaboratoryInstitutional Animal Care and Use Commit-tees approval, with support from DolphinQuest, Inc., the Chicago Zoological Society,and the Office of Naval Research.LITERATURE CITEDCooper MK, Phalen DN, Donahoe SL, Rose K, Šlapeta J.2016. The utility of diversity profiling using Illumina18S rRNA gene amplicon deep sequencing to detectand discriminate Toxoplasma gondii among the cyst-forming coccidia. Vet Parasitol 216:38–45.Desoubeaux G, Del Carmen Piqueras M, Pantin A,Bhattacharya SK, Peschke R, Joachim A, Cray C.2017a. Application of mass spectrometry to elucidatethe pathophysiology of Encephalitozoon cuniculiinfection in rabbits. PLoS One 12:e0177961.Desoubeaux G, Pantin A, Peschke R, Joachim A, Cray C.2017b. Application of western blot analysis for thediagnosis of Encephalitozoon cuniculi infection inrabbits: Example of a quantitative approach. ParasitolRes 116:743–750.Didier ES, Weiss LM. 2006. Microsporidiosis: Currentstatus. Curr Opin Infect Dis 19:485–492.Długońska H. 2017. Are poikilothermic animals real hostsfor Toxoplasma gondii? Ann Parasitol 63:3–5.Fayer R, Fair PA, Bossart GD, Santı́n M. 2008.Examination of naturally exposed bottlenose dolphins(Tursiops truncatus) for Microsporidia, Cryptospo-ridium, and Giardia. J Parasitol 94:143–147.Ghozzi K, Marangi M, Papini R, Lahmar I, Challouf R,Houas N, Dhiab RB, Normanno G, Babba H,Giangaspero A. 2017. First report of Tunisian coastalwater contamination by protozoan parasites usingmollusk bivalves as biological indicators. Mar PollutBull 117:197–202.Goodgame R, Stager C, Marcantel B, Alcocer E, SeguraAM. 1999. Intensity of infection in AIDS-relatedintestinal microsporidiosis. J Infect Dis 180:929–932.Graczyk TK, Sunderland D, Awantang GN, Mashinski Y,Lucy FE, Graczyk Z, Chomicz L, Breysse PN. 2010.Relationships among bather density, levels of humanwaterborne pathogens, and fecal coliform counts inmarine recreational beach water. Parasitol Res 106:1103–1108.Künzel F, Peschke R, Tichy A, Joachim A. 2014.Comparison of an indirect fluorescent antibody testwith western blot for the detection of serumantibodies against Encephalitozoon cuniculi in cats.Parasitol Res 113:4457–4462.Kvach Y, Winkler HM. 2011. The colonization of theinvasive round goby Neogobius melanostomus byparasites in new localities in the southwestern BalticSea. Parasitol Res 109:769–780.Li W, Li Y, Li W, Yang J, Song M, Diao R, Jia H, Lu Y,Zheng J, Zhang X, et al. 2014. Genotypes ofEnterocytozoon bieneusi in livestock in China: Highprevalence and zoonotic potential. PLoS One 9:e97623.Mathis A, Weber R, Deplazes P. 2005. Zoonotic potentialof the microsporidia. Clin Microbiol Rev 18:423–445.Wasson K, Peper RL. 2000. Mammalian microsporidiosis.Vet Pathol 37:113–128.Wu H. 2018. Approximations to the distribution of a teststatistic in covariance structure analysis: A compre-hensive study. Br J Math Stat Psychol 71:334–362.Submitted for publication 28 November 2017.Accepted 15 February 2018.LETTERS 873

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Seroprevalence Survey for Microsporidia in Common Bottlenose Dolphin (\u3ci\u3eTursiops truncatus\u3c/i\u3e): Example of a Quantitative Approach Based on Immunoblotting

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