DISEASES OF AQUATIC ORGANISMSDis Aquat Org

Vol. 91: 47–56, 2010doi: 10.3354/dao02245

Published July 26

INTRODUCTION

The zebrafish Danio rerio is an important researchmodel for the study of infectious disease (Dooley &Zon 2000), developmental and genetic biology(Grunwald & Eisen 2002, Ackermann & Paw 2003),cancer (Amatruda et al. 2002), immunology (Yoder etal. 2002, Trede et al. 2004), toxicology (Hill et al.2005), and drug discovery (Zon & Peterson 2005).Their small size, relative ease of husbandry, largeresearch community, and ex vivo development oftransparent embryos makes them an amenablemodel for such studies. As a result, numerous labora-tory colonies have been established containing wild-type, mutant, and transgenic strains with a wide vari-ety of genetic backgrounds.

Unfortunately, as is the case with any animal model,the zebrafish can be afflicted with a number of dis-eases, potentially confounding experimental resultsand causing growing concern among investigators. Themost common infectious diseases found in laboratoryzebrafish are mycobacteriosis (Kent et al. 2004) and mi-crosporidiosis caused by the microsporidian parasitePseudoloma neurophilia. As the name implies, the mi-crosporidium infects neural tissue in the brain andspinal cord of zebrafish (Matthews et al. 2001).

We recently detected another microsporidium inzebrafish: Pleistophora hyphessobryconis. This is thecausative agent of ‘neon tetra disease,’ which targetsthe skeletal muscle of many aquarium fishes. The pri-mary and type host is the neon tetra Paracheirodoninnesi (Characiformes: Characidae). However, this

© Inter-Research 2010 · www.int-res.com*Email: justin.sanders@oregonstate.edu

Pleistophora hyphessobryconis (Microsporidia)infecting zebrafish Danio rerio in research facilities

Justin L. Sanders1,*, Christian Lawrence2, Donald K. Nichols3, 4, Jeffrey F. Brubaker4, Tracy S. Peterson1, Katrina N. Murray5, Michael L. Kent1

1Department of Microbiology, Oregon State University, Corvallis, Oregon 97331, USA2Children’s Hospital Boston, Aquatic Resources Program, Boston, Massachusetts 02115, USA

3U.S. Army Center for Environmental Health Research and 4U.S. Army Medical Research Institute of Infectious Diseases,Fort Detrick, Maryland 21702, USA

5Zebrafish International Resource Center, University of Oregon, Eugene, Oregon 97403, USA

ABSTRACT: Zebrafish Danio rerio are important models for biomedical research, and thus, there isan increased concern about diseases afflicting them. Here we describe infections by Pleistophorahyphessobryconis (Microsporidia) in zebrafish from 3 laboratories. As reported in other aquariumfishes, affected zebrafish exhibited massive infections in the skeletal muscle, with no involvement ofsmooth or cardiac muscle. In addition, numerous spores within macrophages were observed in thevisceral organs, including the ovaries. Transmission studies and ribosomal RNA (rRNA) genesequence comparisons confirmed that the parasite from zebrafish was P. hyphessobryconis asdescribed from neon tetra Paracheirodon innesi. Ten 15 d old zebrafish were exposed to P. hyphesso-bryconis collected from 1 infected neon tetra, and 7 of 10 fish became infected. Comparison of P.hyphessobryconis small subunit rRNA gene sequence from neon tetra with that obtained fromzebrafish was nearly identical, with <1% difference. Given the severity of infections, P. hyphesso-bryconis should be added to the list of pathogens that should be avoided in zebrafish research facili-ties, and it would be prudent to avoid mixing zebrafish used in research with other aquarium fishes.

KEY WORDS: Zebrafish · Microsporidia · Pleistophora hyphessobryconis

Resale or republication not permitted without written consent of the publisher

Dis Aquat Org 91: 47–56, 2010

microsporidium shows broad host specificity, and hasbeen reported from many species of aquarium fishes inseveral families (Characidae, Cyprinidae, Cyprinodon-tidae, Poecilidae, Cichlidae), including Danio rerio andD. nigrofasciatus (Steffens 1962). Some host rangereports of this parasite were derived from cross trans-mission studies (Canning et al. 1986), but most reportsregarding the host range were from observations ofnaturally infected fishes. Therefore, it is conceivablethat some of these infections may have been caused byother related, undescribed species that are morpholog-ically indistinguishable.

We detected severe muscle infections of Pleistophorahyphessobryconis (Lom & Dyková 1992, Shaw & Kent1999) in zebrafish from 3 separate research facilities.We report here on the case histories, including macro-scopic and histological changes associated with theinfection in laboratory zebrafish. We demonstrate thatthe parasite recently found in zebrafish from researchfacilities was P. hyphessobryconis by conducting crosstransmission experiments and rRNA gene sequencecomparisons with P. hyphessobryconis obtained fromthe type host, neon tetra.

MATERIALS AND METHODS

Case histories. Zebrafish from 3 research facilitieswere examined by histology either as part of routinehealth screening or because fish exhibited clinical dis-ease. The index case (at Lab 1) was evaluated by 2 ofthe authors (D.N. and J.B.), and the specimens from theother 2 facilities were submitted to the Zebrafish Inter-national Resource Center (ZIRC) diagnostic service(http://zebrafish.org/zirc/health/index.php). Informa-tion regarding fish husbandry and quarantine proce-dures was provided by the submitting client.

Histology. Fish were preserved in either 10%buffered formalin (Lab 1) or Dietrich’s fixative (Labs 2and 3) and processed for routine histology. Transversesections were made of the fish from Lab 1; sagittal sec-tions were prepared of the fish from Labs 2 and 3. Allsections were stained with hematoxylin and eosin.Additional tissue sections of the one infected fish fromLab 1 were stained with Lillie-Twort Gram stain(Culling 1974), and sections from several infected fishfrom Lab 3 were also stained with Accustain™,(Sigma-Aldrich), a commercial Brown-Hopps stain.

rRNA gene sequencing. After diagnosis of the infec-tion by histology in several fish from Lab 3, additionallive fish were euthanized and skeletal muscle wasexamined by wet mount. Infected muscle tissues from3 fish were processed for sequencing. Using the QIA-gen Blood and Tissue kit, ~25 mg of muscle tissue wasextracted following the manufacturer’s protocol for

extraction from tissue with an initial proteinase Kdigestion at 56°C for 3 h. Approximately 50 juvenileneon tetras with a suspected history of the infectionwere obtained from a private retail fish store in theCorvallis, Oregon area, USA. Muscle tissue from 1infected fish was frozen and processed for sequencingas above.

PCR was performed using the general microsporid-ian small subunit rRNA gene primers V1F (5’-CACCAG GTT GAT TCT GCC TGA C-3’) and 1492R (5’-GTT ACC TTG TTA CGA CTT-3’). Amplificationswere performed on a Peltier 200 thermocycler (MJResearch) with an initial denaturation at 94°C for2 min, 35 cycles at 94°C for 1 min, 55°C for 1 min, and68°C for 1 min with a final extension at 68°C for 7 min.PCR products were cloned into TOPO TA Cloning vec-tors (Invitrogen) and sequenced in both directionsusing primers flanking the inserted sequence. Toexclude concomitant Pseudoloma neurophilia infec-tion, a reverse primer was designed using the Primer-BLAST program available online from the NationalCenter for Biotechnology Information (NCBI; http://www3.ncbi.nlm.nih.gov/). The novel primer, PleistR(5’-TCT CGC TTG TTC GCG CCT GA-3’), was usedwith the forward primer, V1F, to perform the PCR onsamples from Lab 3 using the same thermocycling con-ditions as described above. PCR products from thesesamples were sequenced directly. All DNA analyzed inthe study was sequenced on an ABI Prism®3730Genetic Analyzer with the BigDye® Terminator v. 3.1Cycle Sequencing kit (Applied Biosystems).

Phylogenetic analysis. The 16S rRNA gene se-quence of Pleistophora hyphessobryconis was alignedwith several Pleistophora sequences and other closelyrelated genera returned by BLAST query of the Gen-Bank database (Altschul et al. 1990) using theClustalW2 software (Larkin et al. 2007). Pseudolomaneurophilia and Glugea anomala (Microsporidia) wereselected as outgroup species. The jModelTest software(Posada & Buckley 2004) was used to determine themost likely model of sequence evolution for the data-set, and phylogenetic analyses were performed usingBayesian inference as implemented in MrBayes3.1.2(Huelsenbeck et al. 2001) and maximum likelihoodalgorithms (Guindon et al. 2009) as implemented in thePHYML webserver (www.atgc-montpellier.fr/phyml/).MrBayes was run using the General Time Reversible(GTR) model of nucleotide substitution with δ-distrib-uted rate variation across sites and a proportion ofinvariable sites (GTR I + G) for 1 000 000 generations.PHYML was run using the GTR model of nucleotidesubstitution, and bootstrap support was based on 100replicates.

The following sequences were obtained from Gen-Bank and used for alignment and subsequent phylo-

48

Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities

genetic analysis: Pleistophora ovariae (AJ252955.1),P. mirandellae (AJ252954.1), Ovipleistophora miran-dellae (AF356223.1), Heterosporis anguillarum(AF387331.1), P. mulleri (AJ438985.1), P. hippoglos-soideos (AJ252953.1), Pleistophora sp. (AJ252957.1), P.typicalis (AJ252956.1), P. anguillarum (U47052.1),Pleistophora sp. 2 (AF044389.1), Trachipleistophora(AJ002605.1), Vavraia culicis (AJ252961.1), Pleisto-phora sp. 3 (AF044390.1), G. anomala (AF056016.1), G.stephani (AF056015.1), G. atherinae (U15987.1), Lomaacerinae (AJ252951.1), Pleistophora sp. 1 (AF044394.1),and Pseudoloma neurophilia (AF322654.1).

Transmission studies. All fish were maintained andtreated humanely, and the study protocol was con-ducted with approval from the Oregon State Univer-sity Institutional Animal Care and Use Committee(ACUP# 3652).

Two groups of 15 d old AB zebrafish (10 fish group–1)were obtained from the Sinnhuber Aquatic ResearchLaboratory, Oregon State University, and each groupwas placed into a 1.5 l tank. Spore counts were ob-tained by a hemocytometer. Homogenized muscle tis-sue containing 50 000 spores from a heavily infectedneon tetra was added to the tank of 1 group. The othergroup was held as an unexposed control. Tanks re-mained static for 12 h postexposure after which waterflow was slowly applied. Fish were held in a flow-through system supplied by ~100 ml h–1 of dechlori-nated tap water heated to 28°C and fed Zeigler® LarvalDiet twice daily. After 30 d, all fish were euthanized byan overdose of tricaine methanesulfate (MS-222) andprocessed for histology. In addition, 2 moribund fishfrom the parasite exposed group were collected at 20 dpostexposure and examined by wet mount.

RESULTS

Case history: Lab 1

Zebrafish at this facility had been maintained as aclosed colony for several years. In an effort to addgenetic diversity to this colony, a group of ~400 adultzebrafish was purchased in June 2006 from a commer-cial wholesale vendor that primarily supplies pet storeswith various species of tropical fish. The new fish wereheld in quarantine for 90 d, and no disease problemswere noted during this time. While in quarantine, thenew fish were segregated from the other fish in a cir-cular 321.8 l tank with a continuous flow-through offresh, non-recycled water. The water source was thesame as that used for the established research colony:a mixture of reverse-osmosis filtered municipal waterand water from a deep on-site well. The fish were pro-vided with the same food as that given to the adult fish

in the established colony, consisting of dry flake food(TetraMin tropical fish flake food) twice daily and livebrine shrimp once daily. At the end of the quarantineperiod, a female fish from the recently purchasedgroup was mated with a male fish from the long-estab-lished colony. The eggs/developing embryos werebleach disinfected by immersion in a water bath con-taining 30 ppm sodium hypochlorite for 2 min afterwhich the embryos were transferred to 10 gal (~38 l)aquariums where they developed into fry. Theseaquariums all shared the same recirculating water sys-tem. The food provided to the fry consisted of liveTetrahymena on Days 1 to 4 post hatching, livemicroworms on Days 3 to 21, and live brine shrimp onDays 6 to 21. After Day 21, the fish were fed flake foodand brine shrimp as described above for adultzebrafish. All live invertebrates fed to the fish werefrom cultures maintained at Lab 1.

At ~1 mo of age, the young fish were transferredfrom the 10 gal (~38 l) aquariums to a single 85 gal(~322 l) circular tank with a continuous flow-through offresh, non-recycled water. In October 2007 (i.e. at13 mo of age), 8 apparently healthy fish from thisgroup were selected at random for routine healthassessment and 1 of these fish was found through his-tologic examination to be infected with microsporidianparasites morphologically consistent with Pleistophorahyphessobryconis. Over several wk after the detectionof the infected fish, the remaining 135 fish of this sib-ling co-hort group were euthanized and examinedgrossly (98 fish) and/or histologically (37 fish). No otherinfected fish were detected in this group and therehave not been any subsequent cases detected in Lab 1.

Case history: Lab 2

A microsporidian infection consistent with Pleisto-phora hyphessobryconis was detected in 1 of 30 mori-bund fish submitted to the diagnostic laboratory ofZIRC. The fish was a wild-type strain purchased froma commercial tropical fish vendor and was being heldin quarantine at the time it became ill. The fish werehatched September 2008 and submitted to the diag-nostic laboratory March 2009.

Case history: Lab 3

The infection was initially detected in 2 of 13 mori-bund fish submitted to the ZIRC diagnostic service inJune 2009. The infected animals were 18 mo old fishfrom the CG1 strain: a clonal, homozygous strain of fishused exclusively for tissue transplantation studies anddeveloped using AB fish obtained from a laboratory

49

Dis Aquat Org 91: 47–56, 2010

colony and a brass mutant line obtained from a petstore (Mizgireuv & Revskoy 2006).

The ‘founding’ CG1 fish in Lab 3 (i.e. grandparentsof the infected fish) were originally imported fromanother institution into Lab 3’s quarantine facility assurface-disinfected eggs. This quarantine facility isphysically separated from the main fish holding room,with isolated recirculating water systems, ultraviolettreatment of effluent post-filtration, restricted access,and dedicated equipment. Fish are moved from quar-antine into the main facility as eggs after being disin-fected in 30 ppm sodium hypochlorite for 2 min andtransferred to sterile water.

After this discovery, more fish in this facility werescreened for the parasite. First, 10 live fish from the samestock in which the initial infection was detected wereshipped live to Oregon State University for histology. Allof these animals, many of which presented obvious clin-ical signs of infection, tested positive for the parasite byhistology. An additional 13 ind. from the subsequent (F3)generation of CG1 in this facility were examined andconfirmed as positive for the infection by histology. Fi-nally, infections were detected in 3 more fish in August2009, 2 more from the F3 generation of the CG1 strainand another fish from a separate population of WIKstrain zebrafish. All 3 of these animals had been exposedto sublethal doses of gamma radiation to suppress theimmune system before tissue transplantation (Traver etal. 2004). The WIK fish in this facility were imported asdescribed for the CGI line and had been maintained in-ternally for many generations, with no known history ofexposure to fish outside of the colony.

In October 2009, 46 AB strain fish from Lab 3 weresubmitted to ZIRC for histological analysis as part ofthe parent institution’s health sentinel monitoring pro-gram. These fish had been directly imported into thefacility from ZIRC as surface disinfected eggs andreared in small groups on each of the facility’s recircu-lating systems. At 6 mo, all of these fish were eutha-nized and examined by histology. A single fish fromthis group was positive for Pleistophora hyphesso-bryconis, and it had been housed in the same rack asthe infected CG1 fish.

Macroscopic changes and clinical signs

The 1 infected fish from Lab 1 showed no obviousclinical or macroscopic changes. The 1 infected fishfrom Lab 2 was sluggish, appeared bloated, and exhib-ited a white area in the integument below the dorsalfin that was obvious in the swimming fish when exam-ined from above.

In Lab 3, large, depigmented regions in the centraldorsal fin area were observed in infected fish while still

swimming in tanks. Some fish also displayed spinalcurvatures. Examination of infected fish with a dissect-ing microscope revealed multifocal to coalescingwhite-gray, slightly raised regions where the skinappeared depigmented (Fig 1). Removal of the skinshowed that the underlying skeletal muscle was white,soft, and edematous (Fig 1). The 1 positive AB fish fromthe Lab 3 sentinel group appeared normal when euth-anized.

Microscopic examinations

Wet mount preparations of skeletal muscle frominfected fish revealed numerous sporophorous vesi-cles, many of which contained fully developed spores.Spores in wet mounts were 7 by 4 µm (n = 20), and con-tained a prominent posterior vacuole measuring ~3 by3 µm (Fig 2). Histopathologic changes were consistentfrom all 3 laboratories and thus are described to-gether. As previously described by other investigators(Schäperclaus 1941, Dyková & Lom 1980, Canning etal. 1986), intramuscular infection by Pleistophorahyphessobryconis microsporidian organisms causeddisruption of skeletal muscle comprising myomericunits. However, in several of the fish examined as apart of this study, the severity of infection was muchmore pronounced and was associated with severechronic inflammation. In these fish, on average, 50 to80% of affected skeletal muscle myofibers displayedextensive liquefactive necrosis and marked expansionby the intramuscular microsporidial parasite. Skeletalmuscle degeneration, denoted by loss of cross-stria-tions as well as fragmented, hypereosinophilic myo-fibers and centralized nuclei, was also observed

50

Fig. 1. Danio rerio. Zebrafish infected with Pleistophorahyphessobryconis. (A) Mottled appearance with light areas(arrows) on flanks. (B) Same fish with skin removed, exhibit-ing opaque regions in muscle (arrows) representing massive

infection. Scale bar = 500 µm

Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities

among the affected myofibers. In sections, musclemyofibers contained from 2 to 30 plus P. hyphesso-bryconis organisms at varying developmental stages,ranging from rounded-up multinucleated meronts tosporophorous vesicles containing sporoblast mothercells and vesicles with mature spores (Fig. 3). Theaffected skeletal muscle myofibers were frequentlysurrounded by large numbers of intermixed histiocytesand lymphocytes with fewer eosinophilic granularcells, which tracked along the endomysial connectivetissue and were accompanied by marked endomysialedema that widely separated myofibers. Regeneratingmyofibers characterized by enhanced sarcoplasmicbasophilia and nuclear rowing were frequently adja-cent to many of the necrotic myofibers (data notshown). Numerous liberated mature spores within theendomysial spaces were surrounded by dense aggre-gates of histiocytes and lymphocytes (Fig. 3B). Phago-cytized mature spores were occasionally seen withinactivated histiocytes.

The parasitic infection in the single fish from Lab 1was limited to the skeletal muscles; however, in thefish from the other 2 laboratories, spores within phago-cytes were also detected in various other organs,including the kidney interstitium, spleen, ovaries,intestine, and mesenteries. Massive numbers of sporeswithin phagocytes were frequently observed in theconnective tissues of the ovary, but never within ova(Fig. 3G). Aggregates of spores were observed in alllayers of the intestine, extending through the serosalsurface and into mesenteric connective tissue. Smoothand cardiac muscle were not affected in any of the fishexamined. Both Gram stain methods were effective for

distinguishing spores. With the Accustain method,mature spores stained deep blue while immaturespores appeared to take up less of the stain (Fig. 3D).With the Lillie-Twort method, fully formed sporesstained deep blue to purple (Fig 3E).

All 7 of the fish from Lab 3 examined by histologyexhibited a mixed infection with Pseudoloma neu-rophilia (Fig 3C). This microsporidium was easily dis-tinguished from Pleistophora hyphessobryconis by itslocation in the central nervous system and lack of asporophorous vesicle with a prominent wall.

Ribosomal DNA sequence

Of the small subunit rRNA gene, 1361 bp was se-quenced from Pleistophora hyphessobryconis obtainedfrom an infected neon tetra and is available in the Gen-Bank database under accession number GU126672.The maximum likelihood and Bayesian analyses pro-duced phylogenetic trees with identical topology(Fig. 4). The topology is also consistent with other phy-logenetic analyses of fish microsporidian parasites(Nilsen 2000). P. hyphessobryconis was placed in theclade containing Ovipleistophora ovariae and O. mi-randellae as well as Heterosporis anguillarum.

A total of 1224 bp (position 1 to 1224) of the smallsubunit rRNA gene was sequenced from Pleistophorahyphessobryconis obtained from 3 zebrafish submittedby Lab 3. All 3 of these sequences were identical, withno insertions/deletions nor transitions/transversions.Comparisons of the 1224 bp subset of the sequenceobtained from the neon tetra and the 3 sequencesobtained from zebrafish showed no insertions/dele-tions and 2 transitions: at site 149 (T to C) and at site180 (G to A) for an overall paired distance of 0.002.

Transmission

Both moribund zebrafish collected from the groupexposed to infected neon tetra tissue exhibited massivemuscle infections based on wet mount observations at20 d postexposure. Histological examination of theremaining fish collected at 30 d showed infections in 5of 8 fish. These fish had light infections exhibiting var-ious stages of development.

DISCUSSION

Identification and taxonomy

Pleistophora hyphessobryconis infections in aquar-ium fishes have been documented for many decades

51

Fig. 2. Danio rerio. Wet mount of Pleistophora hyphesso-bryconis spores. Note prominent posterior vacuole (arrow).

Scale bar = 10 µm

Dis Aquat Org 91: 47–56, 201052

Fig. 3. Danio rerio. Histological sections of zebrafish infected with Pleistophora hyphessobryconis. (A) Numerous sporophorousvesicles with spores and developmental stages within myocytes. (B) Spores in phagocytes associated with chronic infectionsand myolysis (arrows). (C) Mixed infection with P. hyphessobryconis in muscle and xenomas of Pseudoloma neurophilia in spinalcord (arrows). (D) Meronts (m) and developing (sp) and mature spores in sporophorous vesicles. Some fully developed sporesstain deep blue (arrows). (E) Mature spores stain deep blue to purple (arrows); developmental stages are pink (arrow heads).(F) Numerous spores (arrows) throughout all layers of the intestine and mesenteries. (G) Masses of spores in phagocytes (arrows)in ovaries. Scale bars: (A,B,C,F) = 20 µm, (D,E,G) = 10 µm. Stains: (A,B) = hematoxylin and eosin, (C,D,F,G) = Accustain

Gram, (E) = Lillie-Twort Gram

Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities

since the original description in 1941 by Schäperclaus(Schäperclaus 1941). Unusually for microsporidia, theparasite shows remarkably broad host specificity,infecting some 20 species of freshwater fishes in 4orders (Steffens 1962, Schäperclaus 1991, Lom &Dyková 1992). Among those are zebrafish (Opitz 1942)and the dwarf danio Danio nigrofasciatus (Spence etal. 2008). Here we found the infection in 3 separatezebrafish research colonies with no known movementof fish between the facilities. Confirmation of hostranges of parasites with limited morphological charac-ters (such as Microsporidia) often requires cross trans-mission studies or sequence comparisons. There arereports of experimental transmission experiments of P.

hyphessobryconis amongst variousfishes (Canning et al. 1986). Here weconfirmed that the infection in zebra-fish was P. hyphessobryconis by histol-ogy, rRNA gene sequence compar-isons, and cross transmission studies.

We found <1% (0.002) difference insmall subunit rRNA gene sequence be-tween parasites from the type host (theneon tetra) and from zebrafish. This isconsistent with intraspecific variationin this region of the small subunit rRNAfor other Pleistophora spp. and relatedmicroporidia. The intraspecific pair-wise distance between other availablesequences from multiple individuals inGenBank are as follows: P. typicalis,0.013; Heterosporis (syn Pleistophora)anguillarum, 0.02; Ovipleistophora(syn, Pleistophora) mirandellae, 0.069;and P. mulleri., 0.007. Further, wefound an average pairwise interspe-cific distance of 0.198 with all membersof the genera Pleistophora, Oviplei-stophora, and Heterosporis with se-quences published in GenBank. Theminor, but consistent, differences seenbetween the 3 zebrafish sequencescompared to that of neon tetra mightsuggest different strains of the parasite.Regardless, the parasite from neontetra was easily transmitted to ze-brafish. Our findings agree with previ-ous transmission studies, confirmingthe broad host specificity of this mi-crosporidium (Canning et al. 1986).

It was noted previously that thegenus Pleistophora does not form amonophyletic clade (Nilsen et al.1998). In fact, ultrastructural charac-terization and molecular analyses of

the small subunit rRNA gene sequences have resultedin the movement of P. mirandellae (Pekkarinen et al.2002) and P. anguillarum (Lom et al. 2000) to new gen-era (Ovipleistophora and Heterosporis, respectively).Prior ultrastructural descriptions of P. hyphessobryco-nis (Lom & Corliss 1967) confirm the placement of thisspecies in the genus Pleistophora as redescribed byCanning & Nicholas (1980). However, phylogeneticanalyses of P. hyphessobryconis place it more closelyto both O. mirandellae and H. anguillarum than P. typ-icalis, the type species described for the genus (Fig. 4).Interestingly, unlike the members of Ovipleistophora,this species does not infect oocytes but rather myocytesas does the type species of this genus, P. typicalis.

53

Fig. 4. Phylogenetic tree of Pleistophora hyphessobryconis obtained from neontetra and related microsporidia. 1361 bp small subunit rRNA gene sequencesfrom 20 microsporidia infecting fish were used to reconstruct phylogeny. Maxi-mum likelihood tree is shown. Branch numbers: maximum likelihood bootstrapsupport based on 100 replicates/Bayesian posterior probabilities. Genus names:as recorded in GenBank with new genus designations in parentheses. Mi-crosporidian parasite Pseudoloma neurophilia was selected as an outgroup taxa.

Scale bar represents number of base substitutions per site

Dis Aquat Org 91: 47–56, 2010

Comprehensive analysis of the genus is beyond thescope of this study, but further investigation appears tobe needed.

Transmission

The 3 facilities had no history of sharing fish and arelocated in different areas of the United States. Thus,we presume that the infections arose in 3 independentinstances by exposure to other species of infectedaquarium fishes. Indeed, the CG1 line was actuallyderived from zebrafish purchased from a pet store(Mizgireuv & Revskoy 2006). The affected fish fromLab 2 had also been purchased from a pet store and thematernal parent of the infected fish at Lab 1 had beenpurchased from a vendor that supplied fish to thecommercial pet trade. Transmission of Pleistophorahyphessobryconis, ostensibly per os, was achieved byplacing large numbers of spores in water with fish, amethod that has been used by others (Canning et al.1986). As with other microsporidia, it is assumed thatinfection is initiated by ingestion of spores. Sporesreleased from dead fish are a likely source of infection,but could also be released from the intestines.Schäperclaus (1941) suggested that spores may bereleased from the skin or urinary tract of infected fish.

Zebrafish spawn frequently within aquaria, and tankmates quickly eat available eggs (Lawrence 2007,Spence et al. 2008). Although spores were not foundwithin eggs, the massive numbers seen within ovariesof some infected zebrafish suggest that infectiousspores could be released during spawning and wouldthus be available to fish feeding on eggs or to infect thenext generation of fish. Maternal transmission, includ-ing true vertical transmission within eggs, has beenverified or implicated for other microsporidia of fishes(Docker et al. 1997, Kent & Bishop-Stewart 2003,Phelps & Goodwin 2008). Schäperclaus (1941) sug-gested the possibility of maternal transmission as hefound infections in 8 d old neon tetras derived frominfected parents. Once established in zebrafish, it isconceivable that the infection could be maternallytransferred to the next generation. This could evenoccur with spores outside the egg as microsporidianspores are very resistant to chlorine (Ferguson et al.2007). This provides one explanation for the occur-rence of the infection in fish derived from surface-disinfected eggs. Alternatively, these fish may havebecome infected by an unrecognized breach in biose-curity protocols.

The infected fish from Lab 1 was the 13 mo old off-spring of a female fish that had been purchased froman outside vendor and brought into the laboratory. Ifthis fish was infected as an embryo, this would mean

that the fish was subclinically infected for more than1 yr. All of the fish purchased from the vendor hadbeen removed from Lab 1 more than 7 mo before theinfected fish was detected. Even if the fish had notbeen infected as an embryo but instead at a later timethrough accidental cross-contamination with aninfected fish from the group purchased from the ven-dor, it indicates that this fish was subclinically infectedfor more than 7 mo.

As with other microsporidian infections, it is likelythat immune status influences susceptibility of zebra-fish to Pleistophora hyphessobryconis. Recently, Ram-say et al. (2009) showed that stress enhances infectionsof Pseudoloma neurophilia in zebrafish. Infections byP. hyphessobryconis were widespread in only onestrain (CG1) in one of the labs with the infection sug-gesting that this line may be particularly susceptible tothe parasite. The microsporidium was also found in fish(CG1 and WIK) that were irradiated and thus wereimmune compromised. Although immune status likelyaffects the susceptibility and progression of the diseasein zebrafish, apparently immunocompetent fish canbecome infected as demonstrated by reports of P.hyphessobryconis in zebrafish from aquaria (Opitz1942, Steffens 1962), and experimental infections inpresumably healthy zebrafish reported here. The latterwere exposed at 15 d old, a time at which fish haveinnate immunity but adaptive immunity has not fullydeveloped (Lam et al. 2004, Trede et al. 2004).

Pathology

The infection in zebrafish was consistent with reportsfrom neon tetras and other species (Dyková & Lom 1980,Canning et al. 1986, Schäperclaus 1991). The mostremarkable pathological feature of Pleistophora hyphes-sobryconis is the severe intensity of infection in theskeletal muscle, with well over half of the myofibers con-taining numerous spores and developmental stages insome fish. Parasites within myocytes were not associatedwith inflammation; however, mature spores releasedfrom degenerate myofibers into interstitial spaces wereconsistently associated with inflammation, and these lib-erated spores were often engulfed by phagocytes. This issimilar to that seen with other intramuscular parasites offishes, such as Kudoa thyrsites of salmon (Moran et al.1999). Likewise, other microsporidia show a similarsequelae of infection in which spores elicit significantinflammation only after they are released from theirintracellular environment within xenomas (Dyková &Lom 1980, Kent & Speare 2005). Microsporidian sporesremain intact within phagocytes, and thus may be trans-ported to sites beyond where they originally developed(Kent et al. 1999). The occurrence of large numbers of

54

Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities

spores of P. hyphessobryconis within phagocytes inorgans other than the skeletal muscle, as seen in the pre-sent study, has been previously observed (Lom & Dyková1992). Spores of microsporidia are Gram positive in his-tological sections (Gardiner et al. 1998, Bruno et al.2006). Both Lilly-Twort and Accustain Gram stains werevery effective in demonstrating spores and particularlyuseful for visualizing individual spores within visceralorgans.

Some infected zebrafish exhibited concurrent in-fections by Pseudoloma neurophilia, a common micro-sporidian parasite of zebrafish that demonstratesmyelinotropic behavior and is directly associated withencephalomyelitis and polyneuritis involving the peri-pheral nerves and spinal nerve roots. The 2 micro-sporidian infections can be easily differentiated byhistology. Skeletal muscle is the primary site of de-velopment of Pleistophora hyphessobryconis, withprominent developmental stages and spores withinthick-walled sporophorous vesicles. In contrast, the cen-tral and peripheral nervous systems are the primary sitesof infection for P. neurophilia and finding individualspores or xenomas in extraneural tissue is uncommon. Incases of myositis attributed to P. neurophilia, few sporesare found in the muscle, but these are frequently associ-ated with severe chronic inflammation (Matthews et al.2001). In the case of coinfections, the opaque, depig-mented muscle lesions were clearly caused by P. hyphes-sobryconis. However, it is possible that P. neurophiliacontributed to other clinical changes.

Given the potential for severe infections and long-term subclinical chronic infections, Pleistophora hy-phessobryconis should be added to the list of pathogensthat should be avoided in zebrafish research facilities.As suggested for Pseudoloma neurophilia (Kent et al.2009), the most feasible strategy would be to hold broodfish in quarantine and screen them and their progenyfor the infection using a PCR test specific to the para-site. Sentinel fish programs and sampling of moribundfish are also recommended as a means of surveillanceof colony health. Notably, 2 of these cases were fromfish that had been in contact with commercial pet storefish. It would be prudent to not mix zebrafish used in re-search with other aquarium fishes. Further studies onthe role of maternal transmission, susceptibility ofspores to disinfectants, and the role of age and fishstrain in severity of disease are all warranted.

Acknowledgements. This study was supported by grants fromthe National Institutes of Health (NIH NCRR 5R24RR017386-02 and NIH NCRR P40 RR12546-03S1). Mention of a brandname does not imply endorsement of the product by the U.S.Federal Government. Opinions, interpretations, conclusions,and recommendations stated are those of the authors andare not necessarily endorsed by the U.S. Army. We thankG. Sanders, University of Washington, for manuscript review,

M. Schuster for assistance in translation of German refer-ences, and K. Berkenkamp for histology slide preparation andtechnical support.

LITERATURE CITED

Ackermann G, Paw BH (2003) Zebrafish: a genetic model forvertebrate organogenesis and human disorders. FrontBiosci 8:d1227–d1253

Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990)Basic local alignment search tool. J Mol Biol 215:403–410

Amatruda JF, Shepard JL, Stern HM, Zon LI (2002) Zebrafishas a cancer model system. Cancer Cell 1:229–231

Bruno DW, Nowak B, Elliott DG (2006) Guide to the identifi-cation of fish protozoan and metazoan parasites in stainedtissue sections. Dis Aquat Org 70:1–36

Canning E, Nicholas JP (1980) Genus Pleistophora (PhylumMicrospora): redescription of the type species,Pleistophora typicalis Gurley, 1893 and ultrastructuralcharacterization of the genus. J Fish Dis 3:317–338

Canning EU, Lom J, Dyková I (1986) Description of speciesinfecting fish: Pleistophora hyphessobryconis Schäper-claus, 1941. In: The microsporidia of vertebrates. Acade-mic Press, London, p 101–107

Culling CFA (1974) Microorganisms. In: Handbook ofhistopathological and histochemical techniques. Butter-worths, London, p 391–403

Docker M, Devlin R, Richard J, Khattra J, Kent M (1997) Sen-sitive and specific polymerase chain reaction assay fordetection of Loma salmonae (Microsporea). Dis Aquat Org29:41–48

Dooley K, Zon LI (2000) Zebrafish: a model system for thestudy of human disease. Curr Opin Genet Dev 10:252–256

Dyková I, Lom J (1980) Tissue reactions to microsporidianinfections in fish. J Fish Dis 3:265–283

Ferguson JA, Watral V, Schwindt AR, Kent ML (2007) Sporesof two fish microsporidia (Pseudoloma neurophilia andGlugea anomola) are highly resistant to chlorine. DisAquat Org 76:205–214

Gardiner CH, Fayer R, Dubey JP (1998) Microspora. In: Anatlas of protozoan parasites in animal tissues. ArmedForces Institute of Pathology, Washington, DC, p 12–13

Grunwald DJ, Eisen JS (2002) Headwaters of the zebrafish—emergence of a new model vertebrate. Nat Rev Genet 3:717–724

Guindon S, Delsuc F, Dufayard J, Gascuel O (2009) Estimat-ing maximum likelihood phylogenies with PhyML. Meth-ods Mol Biol 537:113–137

Hill AJ, Teraoka H, Heideman W, Peterson RE (2005)Zebrafish as a model vertebrate for investigating chemicaltoxicity. Toxicol Sci 86:6–19

Huelsenbeck JP, Ronquist F, Nielsen R, Bollback JP (2001)Bayesian inference of phylogeny and its impact on evolu-tionary biology. Science 294:2310–2314

Kent ML, Bishop-Stewart JK (2003) Transmission and tissuedistribution of Pseudoloma neurophilia (Microsporidia) ofzebrafish, Danio rerio (Hamilton). J Fish Dis 26:423–426

Kent ML, Speare DJ (2005) Review of the sequential develop-ment of Loma salmonae (Microsporidia) based on experi-mental infections of rainbow trout (Oncorhynchus mykiss)and Chinook salmon (O. tshawytscha). Folia Parasitol(Prague) 52:63–68

Kent ML, Dawe SC, Speare DJ (1999) Resistance to reinfectionin chinook salmon Oncorhynchus tshawytscha to Lomasalmonae (Microsporidia). Dis Aquat Org 37:205–208

Kent ML, Whipps CM, Matthews JL, Florio D, Watral V,

55

Dis Aquat Org 91: 47–56, 2010

Bishop-Stewart JK, Poort M, Bermudez L (2004) Mycobac-teriosis in zebrafish (Danio rerio) research facilities. CompBiochem Physiol C Toxicol Pharmacol 138:383–390

Kent ML, Feist SW, Harper C, Hoogstraten-Miller S andothers (2009) Recommendations for control of pathogensand infectious diseases in fish research facilities. CompBiochem Physiol C Toxicol Pharmacol 149:240–248

Lam SH, Chua HL, Gong Z, Lam TJ, Sin YM (2004) Develop-ment and maturation of the immune system in zebrafish,Danio rerio: a gene expression profiling, in situ hybridizationand immunological study. Dev Comp Immunol 28:9–28

Larkin MA, Blackshields G, Brown NP, Chenna R and 9 oth-ers (2007) ClustalW and Clustal X version 2.0. Bioinfor-matics 23:2947–2948

Lawrence C (2007) The husbandry of zebrafish (Danio rerio):a review. Aquaculture 269:1–20

Lom J, Corliss JO (1967) Ultrastructural observations on thedevelopment of the microsporidian protozoon Plistophorahyphessobryconis Schäperclaus*. J Eukaryot Microbiol14:141–152

Lom J, Dyková I (1992) Microsporidia (Phylum MicrosporaSprague, 1977). In: Protozoan parasites of fishes. Elsevier,Amsterdam, p 125–157

Lom J, Dyková I, Wang CH, Lo CF, Kou GH (2000) Ultrastruc-tural justification for the transfer of Pleistophora anguil-larum Hoshina, 1959 to the genus Heterosporis Schubert,1969. Dis Aquat Org 43:225–231

Matthews J, Brown A, Larison K, Bishop-Stewart J, Rogers P,Kent M (2001) Pseudoloma neurophilia ng, n. sp., a newmicrosporidium from the central nervous system of thezebrafish (Danio rerio). J Eukaryot Microbiol 48:227–233

Mizgireuv IV, Revskoy SY (2006) Transplantable tumor linesgenerated in clonal zebrafish. Cancer Res 66:3120–3125

Moran JDW, Margolis L, Webster JM, Kent ML (1999) Devel-opment of Kudoa thyrsites (Myxozoa: Myxosporea) in net-pen-reared Atlantic salmon determined by light micro-scopy and a polymerase chain reaction test. Dis Aquat Org37:185–193

Nilsen F (2000) Small subunit ribosomal DNA phylogeny ofmicrosporidia with particular reference to genera thatinfect fish. J Parasitol 86:128–133

Nilsen F, Endresen C, Hordvik I (1998) Molecular phylogenyof microsporidians with particular reference to speciesthat infect the muscles of fish. J Eukaryot Microbiol 45:535–543

Opitz H (1942) Mikrosporidienkrankheit Plistophora auchbeim Hemigrammus ocellifer und Brachydanio rerio.Wochenschrift für Aquarien- und Terrarienkunde 39:75–77

Pekkarinen M, Lom J, Nilsen F (2002) Ovipleistophora gen.n., a new genus for Pleistophora mirandellae-like micro-sporidia. Dis Aquat Org 48:133–142

Phelps NBD, Goodwin AE (2008) Vertical transmission ofOvipleistophora ovariae (microspora) within the eggs ofthe golden shiner. J Aquat Anim Health 20:45–53

Posada D, Buckley TR (2004) Model selection and modelaveraging in phylogenetics: advantages of Akaike infor-mation criterion and Bayesian approaches over likelihoodratio tests. Syst Biol 53:793–808

Ramsay JM, Watral V, Schreck CB, Kent ML (2009) Pseudo-loma neurophilia (Microsporidia) infections in zebrafish(Danio rerio): effects of stress on survival, growth andreproduction. Dis Aquat Org 88:69–84

Schäperclaus W (1941) Eine neue Mikrosporidien-krankheitbeim Neonfisch und seinen Verwandten. Wochenschriftfür Aquarien- und Terrarienkunde 39/40:381–384

Schäperclaus W (1991) Plistophora disease of neon and otheraquarium fish (Pleistophorosis). In: Schäperclaus W,Kulow H, Schreckenbach K (eds). Fish diseases, Vol 2, 5thedn. Amerind Publishing, New Dehli

Shaw R, Kent M (1999) Fish microsporidia. In: Wittner M,Weiss L (eds) The microsporidia and microsporidiosis.ASM Press, Washington, DC, p 418–444

Spence R, Gerlach G, Lawrence C, Smith C (2008) The behav-iour and ecology of the zebrafish, Danio rerio. Biol RevCamb Philos Soc 83:13–34

Steffens W (1962) Der heutige Stand der Verbreitung vonPlistophora hyphessobryconis Schäperclaus 1941 (Sporo-zoa, Microsporidia). Parasitol Res 21:535–541

Traver D, Winzeler A, Stern HM, Mayhall EA and others (2004)Effects of lethal irradiation in zebrafish and rescue byhematopoietic cell transplantation. Blood 104:1298–1305

Trede NS, Langenau DM, Traver D, Look A, Zon LI (2004)The use of zebrafish to understand immunity. Immunity20:367–379

Yoder JA, Nielsen ME, Amemiya CT, Litman GW (2002)Zebrafish as an immunological model system. MicrobesInfect 4:1469–1478

Zon LI, Peterson RT (2005) In vivo drug discovery in thezebrafish. Nat Rev Drug Discov 4:35–44

56

Editorial responsibility: Dieter Steinhagen,Hannover, Germany

Submitted: March 8, 2010; Accepted: May 19, 2010Proofs received from author(s): July 3, 2010