Vol. 91: 47–56, 2010 DISEASES OF AQUATIC ORGANISMS Published July 26 doi: 10.3354/dao02245 Dis Aquat Org

Pleistophora hyphessobryconis () 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, USA 2Children’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 is an increased concern about diseases afflicting them. Here we describe infections by Pleistophora hyphessobryconis (Microsporidia) in zebrafish from 3 laboratories. As reported in other fishes, affected zebrafish exhibited massive infections in the skeletal muscle, with no involvement of smooth or cardiac muscle. In addition, numerous spores within macrophages were observed in the visceral organs, including the ovaries. Transmission studies and ribosomal RNA (rRNA) gene sequence comparisons confirmed that the parasite from zebrafish was P. hyphessobryconis as described from neon innesi. Ten 15 d old zebrafish were exposed to P. hyphesso- bryconis collected from 1 infected , and 7 of 10 fish became infected. Comparison of P. hyphessobryconis small subunit rRNA gene sequence from neon tetra with that obtained from zebrafish 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

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INTRODUCTION Unfortunately, as is the case with any model, the zebrafish can be afflicted with a number of dis- The zebrafish Danio rerio is an important research eases, potentially confounding experimental results model for the study of infectious disease (Dooley & and causing growing concern among investigators. The Zon 2000), developmental and genetic biology most common infectious diseases found in laboratory (Grunwald & Eisen 2002, Ackermann & Paw 2003), zebrafish are mycobacteriosis (Kent et al. 2004) and mi- cancer (Amatruda et al. 2002), immunology (Yoder et crosporidiosis caused by the microsporidian parasite al. 2002, Trede et al. 2004), toxicology (Hill et al. Pseudoloma neurophilia. As the name implies, the mi- 2005), and drug discovery (Zon & Peterson 2005). crosporidium infects neural tissue in the brain and Their small size, relative ease of husbandry, large spinal cord of zebrafish (Matthews et al. 2001). research community, and ex vivo development of We recently detected another microsporidium in transparent embryos makes them an amenable zebrafish: Pleistophora hyphessobryconis. This is the model for such studies. As a result, numerous labora- causative agent of ‘neon tetra disease,’ which targets tory colonies have been established containing wild- the skeletal muscle of many aquarium fishes. The pri- type, mutant, and transgenic strains with a wide vari- mary and type host is the neon tetra Paracheirodon ety of genetic backgrounds. innesi (: ). However, this

*Email: [email protected] © Inter-Research 2010 · www.int-res.com 48 Dis Aquat Org 91: 47–56, 2010

microsporidium shows broad host specificity, and has extraction from tissue with an initial proteinase K been reported from many species of aquarium fishes in digestion at 56°C for 3 h. Approximately 50 juvenile several families (Characidae, Cyprinidae, Cyprinodon- neon tetras with a suspected history of the infection tidae, Poecilidae, Cichlidae), including Danio rerio and were obtained from a private retail fish store in the D. nigrofasciatus (Steffens 1962). Some host range Corvallis, Oregon area, USA. Muscle tissue from 1 reports of this parasite were derived from cross trans- infected fish was frozen and processed for sequencing mission studies (Canning et al. 1986), but most reports as above. regarding the host range were from observations of PCR was performed using the general microsporid- naturally infected fishes. Therefore, it is conceivable ian small subunit rRNA gene primers V1F (5’-CAC that some of these infections may have been caused by CAG GTT GAT TCT GCC TGA C-3’) and 1492R (5’- other related, undescribed species that are morpholog- GTT ACC TTG TTA CGA CTT-3’). Amplifications ically indistinguishable. were performed on a Peltier 200 thermocycler (MJ We detected severe muscle infections of Pleistophora Research) with an initial denaturation at 94°C for hyphessobryconis (Lom & Dyková 1992, Shaw & Kent 2 min, 35 cycles at 94°C for 1 min, 55°C for 1 min, and 1999) in zebrafish from 3 separate research facilities. 68°C for 1 min with a final extension at 68°C for 7 min. We report here on the case histories, including macro- PCR products were cloned into TOPO TA Cloning vec- scopic and histological changes associated with the tors (Invitrogen) and sequenced in both directions infection in laboratory zebrafish. We demonstrate that using primers flanking the inserted sequence. To the parasite recently found in zebrafish from research exclude concomitant Pseudoloma neurophilia infec- facilities was P. hyphessobryconis by conducting cross tion, a reverse primer was designed using the Primer- transmission experiments and rRNA gene sequence BLAST program available online from the National comparisons with P. hyphessobryconis obtained from Center for Biotechnology Information (NCBI; http:// the type host, neon tetra. www3.ncbi.nlm.nih.gov/). The novel primer, PleistR (5’-TCT CGC TTG TTC GCG CCT GA-3’), was used with the forward primer, V1F, to perform the PCR on MATERIALS AND METHODS samples from Lab 3 using the same thermocycling con- ditions as described above. PCR products from these Case histories. Zebrafish from 3 research facilities samples were sequenced directly. All DNA analyzed in were examined by histology either as part of routine the study was sequenced on an ABI Prism®3730 health screening or because fish exhibited clinical dis- Genetic Analyzer with the BigDye® Terminator v. 3.1 ease. The index case (at Lab 1) was evaluated by 2 of Cycle Sequencing kit (Applied Biosystems). the authors (D.N. and J.B.), and the specimens from the Phylogenetic analysis. The 16S rRNA gene se- other 2 facilities were submitted to the Zebrafish Inter- quence of Pleistophora hyphessobryconis was aligned national Resource Center (ZIRC) diagnostic service with several Pleistophora sequences and other closely (http://zebrafish.org/zirc/health/index.php). Informa- related genera returned by BLAST query of the Gen- tion regarding fish husbandry and quarantine proce- Bank database (Altschul et al. 1990) using the dures was provided by the submitting client. ClustalW2 software (Larkin et al. 2007). Pseudoloma Histology. Fish were preserved in either 10% neurophilia and Glugea anomala (Microsporidia) were buffered formalin (Lab 1) or Dietrich’s fixative (Labs 2 selected as outgroup species. The jModelTest software and 3) and processed for routine histology. Transverse (Posada & Buckley 2004) was used to determine the sections were made of the fish from Lab 1; sagittal sec- most likely model of sequence evolution for the data- tions were prepared of the fish from Labs 2 and 3. All set, and phylogenetic analyses were performed using sections were stained with hematoxylin and eosin. Bayesian inference as implemented in MrBayes3.1.2 Additional tissue sections of the one infected fish from (Huelsenbeck et al. 2001) and maximum likelihood Lab 1 were stained with Lillie-Twort Gram stain algorithms (Guindon et al. 2009) as implemented in the (Culling 1974), and sections from several infected fish PHYML webserver (www.atgc-montpellier.fr/phyml/). from Lab 3 were also stained with Accustain™, MrBayes was run using the General Time Reversible (Sigma-Aldrich), a commercial Brown-Hopps stain. (GTR) model of nucleotide substitution with δ-distrib- rRNA gene sequencing. After diagnosis of the infec- uted rate variation across sites and a proportion of tion by histology in several fish from Lab 3, additional invariable sites (GTR I + G) for 1 000 000 generations. live fish were euthanized and skeletal muscle was PHYML was run using the GTR model of nucleotide examined by wet mount. Infected muscle tissues from substitution, and bootstrap support was based on 100 3 fish were processed for sequencing. Using the QIA- replicates. gen Blood and Tissue kit, ~25 mg of muscle tissue was The following sequences were obtained from Gen- extracted following the manufacturer’s protocol for Bank and used for alignment and subsequent phylo- Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities 49

genetic analysis: Pleistophora ovariae (AJ252955.1), in the established colony, consisting of dry flake food P. mirandellae (AJ252954.1), Ovipleistophora miran- (TetraMin tropical fish flake food) twice daily and live dellae (AF356223.1), Heterosporis anguillarum once daily. At the end of the quarantine (AF387331.1), P. mulleri (AJ438985.1), P. hippoglos- period, a female fish from the recently purchased soideos (AJ252953.1), Pleistophora sp. (AJ252957.1), P. group was mated with a male fish from the long-estab- typicalis (AJ252956.1), P. anguillarum (U47052.1), lished colony. The eggs/developing embryos were Pleistophora sp. 2 (AF044389.1), Trachipleistophora bleach disinfected by immersion in a water bath con- (AJ002605.1), Vavraia culicis (AJ252961.1), Pleisto- taining 30 ppm sodium hypochlorite for 2 min after phora sp. 3 (AF044390.1), G. anomala (AF056016.1), G. which the embryos were transferred to 10 gal (~38 l) stephani (AF056015.1), G. atherinae (U15987.1), Loma where they developed into fry. These acerinae (AJ252951.1), Pleistophora sp. 1 (AF044394.1), aquariums all shared the same recirculating water sys- and Pseudoloma neurophilia (AF322654.1). tem. The food provided to the fry consisted of live Transmission studies. All fish were maintained and Tetrahymena on Days 1 to 4 post hatching, live treated humanely, and the study protocol was con- microworms on Days 3 to 21, and live brine shrimp on ducted with approval from the Oregon State Univer- Days 6 to 21. After Day 21, the fish were fed flake food sity Institutional Animal Care and Use Committee and brine shrimp as described above for adult (ACUP# 3652). zebrafish. All live invertebrates fed to the fish were Two groups of 15 d old AB zebrafish (10 fish group–1) from cultures maintained at Lab 1. were obtained from the Sinnhuber Aquatic Research At ~1 mo of age, the young fish were transferred Laboratory, Oregon State University, and each group from the 10 gal (~38 l) aquariums to a single 85 gal was placed into a 1.5 l tank. Spore counts were ob- (~322 l) circular tank with a continuous flow-through of tained by a hemocytometer. Homogenized muscle tis- fresh, non-recycled water. In October 2007 (i.e. at sue containing 50 000 spores from a heavily infected 13 mo of age), 8 apparently healthy fish from this neon tetra was added to the tank of 1 group. The other group were selected at random for routine health group was held as an unexposed control. Tanks re- assessment and 1 of these fish was found through his- mained static for 12 h postexposure after which water tologic examination to be infected with microsporidian flow was slowly applied. Fish were held in a flow- parasites morphologically consistent with Pleistophora through system supplied by ~100 ml h–1 of dechlori- hyphessobryconis. Over several wk after the detection nated tap water heated to 28°C and fed Zeigler® Larval of the infected fish, the remaining 135 fish of this sib- Diet twice daily. After 30 d, all fish were euthanized by ling co-hort group were euthanized and examined an overdose of tricaine methanesulfate (MS-222) and grossly (98 fish) and/or histologically (37 fish). No other processed for histology. In addition, 2 moribund fish infected fish were detected in this group and there from the parasite exposed group were collected at 20 d have not been any subsequent cases detected in Lab 1. postexposure and examined by wet mount.

Case history: Lab 2 RESULTS A microsporidian infection consistent with Pleisto- Case history: Lab 1 phora hyphessobryconis was detected in 1 of 30 mori- bund fish submitted to the diagnostic laboratory of Zebrafish at this facility had been maintained as a ZIRC. The fish was a wild-type strain purchased from closed colony for several years. In an effort to add a commercial tropical fish vendor and was being held genetic diversity to this colony, a group of ~400 adult in quarantine at the time it became ill. The fish were zebrafish was purchased in June 2006 from a commer- hatched September 2008 and submitted to the diag- cial wholesale vendor that primarily supplies pet stores nostic laboratory March 2009. with various species of tropical fish. The new fish were held in quarantine for 90 d, and no disease problems were noted during this time. While in quarantine, the Case history: Lab 3 new fish were segregated from the other fish in a cir- cular 321.8 l tank with a continuous flow-through of The infection was initially detected in 2 of 13 mori- fresh, non-recycled water. The water source was the bund fish submitted to the ZIRC diagnostic service in same as that used for the established research colony: June 2009. The infected were 18 mo old fish a mixture of reverse-osmosis filtered municipal water from the CG1 strain: a clonal, homozygous strain of fish and water from a deep on-site well. The fish were pro- used exclusively for tissue transplantation studies and vided with the same food as that given to the adult fish developed using AB fish obtained from a laboratory 50 Dis Aquat Org 91: 47–56, 2010

colony and a brass mutant line obtained from a pet swimming in tanks. Some fish also displayed spinal store (Mizgireuv & Revskoy 2006). curvatures. Examination of infected fish with a dissect- The ‘founding’ CG1 fish in Lab 3 (i.e. grandparents ing microscope revealed multifocal to coalescing of the infected fish) were originally imported from white-gray, slightly raised regions where the skin another institution into Lab 3’s quarantine facility as appeared depigmented (Fig 1). Removal of the skin surface-disinfected eggs. This quarantine facility is showed that the underlying skeletal muscle was white, physically separated from the main fish holding room, soft, and edematous (Fig 1). The 1 positive AB fish from with isolated recirculating water systems, ultraviolet the Lab 3 sentinel group appeared normal when euth- treatment of effluent post-filtration, restricted access, anized. 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 and Microscopic examinations transferred to sterile water. After this discovery, more fish in this facility were Wet mount preparations of skeletal muscle from screened for the parasite. First, 10 live fish from the same infected fish revealed numerous sporophorous vesi- stock in which the initial infection was detected were cles, many of which contained fully developed spores. shipped live to Oregon State University for histology. All Spores in wet mounts were 7 by 4 µm (n = 20), and con- of these animals, many of which presented obvious clin- tained a prominent posterior vacuole measuring ~3 by ical signs of infection, tested positive for the parasite by 3 µm (Fig 2). Histopathologic changes were consistent histology. An additional 13 ind. from the subsequent (F3) from all 3 laboratories and thus are described to- generation of CG1 in this facility were examined and gether. As previously described by other investigators confirmed as positive for the infection by histology. Fi- (Schäperclaus 1941, Dyková & Lom 1980, Canning et nally, infections were detected in 3 more fish in August al. 1986), intramuscular infection by Pleistophora 2009, 2 more from the F3 generation of the CG1 strain hyphessobryconis microsporidian organisms caused and another fish from a separate population of WIK disruption of skeletal muscle comprising myomeric strain zebrafish. All 3 of these animals had been exposed units. However, in several of the fish examined as a to sublethal doses of gamma radiation to suppress the part of this study, the severity of infection was much immune system before tissue transplantation (Traver et more pronounced and was associated with severe al. 2004). The WIK fish in this facility were imported as chronic inflammation. In these fish, on average, 50 to described for the CGI line and had been maintained in- 80% of affected skeletal muscle myofibers displayed ternally for many generations, with no known history of extensive liquefactive necrosis and marked expansion exposure to fish outside of the colony. by the intramuscular microsporidial parasite. Skeletal In October 2009, 46 AB strain fish from Lab 3 were muscle degeneration, denoted by loss of cross-stria- submitted to ZIRC for histological analysis as part of tions as well as fragmented, hypereosinophilic myo- the parent institution’s health sentinel monitoring pro- fibers and centralized nuclei, was also observed gram. These fish had been directly imported into the facility from ZIRC as surface disinfected eggs and reared 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 from this group was positive for Pleistophora hyphesso- bryconis, and it had been housed in the same rack as the infected CG1 fish.

Macroscopic changes and clinical signs

The 1 infected fish from Lab 1 showed no obvious clinical or macroscopic changes. The 1 infected fish from Lab 2 was sluggish, appeared bloated, and exhib- ited a white area in the integument below the dorsal fin that was obvious in the swimming fish when exam- Fig. 1. Danio rerio. Zebrafish infected with Pleistophora hyphessobryconis. (A) Mottled appearance with light areas ined from above. (arrows) on flanks. (B) Same fish with skin removed, exhibit- In Lab 3, large, depigmented regions in the central ing opaque regions in muscle (arrows) representing massive dorsal fin area were observed in infected fish while still infection. Scale bar = 500 µm Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities 51

distinguishing spores. With the Accustain method, mature spores stained deep blue while immature spores appeared to take up less of the stain (Fig. 3D). With the Lillie-Twort method, fully formed spores stained deep blue to purple (Fig 3E). All 7 of the fish from Lab 3 examined by histology exhibited a mixed infection with Pseudoloma neu- rophilia (Fig 3C). This microsporidium was easily dis- tinguished from Pleistophora hyphessobryconis by its location in the central nervous system and lack of a sporophorous vesicle with a prominent wall.

Ribosomal DNA sequence

Of the small subunit rRNA gene, 1361 bp was se- quenced from Pleistophora hyphessobryconis obtained from an infected neon tetra and is available in the Gen- Fig. 2. Danio rerio. Wet mount of Pleistophora hyphesso- Bank database under accession number GU126672. bryconis spores. Note prominent posterior vacuole (arrow). The maximum likelihood and Bayesian analyses pro- Scale bar = 10 µm duced phylogenetic trees with identical topology (Fig. 4). The topology is also consistent with other phy- among the affected myofibers. In sections, muscle logenetic analyses of fish microsporidian parasites myofibers contained from 2 to 30 plus P. hyphesso- (Nilsen 2000). P. hyphessobryconis was placed in the bryconis organisms at varying developmental stages, clade containing Ovipleistophora ovariae and O. mi- ranging from rounded-up multinucleated meronts to randellae as well as Heterosporis anguillarum. sporophorous vesicles containing sporoblast mother A total of 1224 bp (position 1 to 1224) of the small cells and vesicles with mature spores (Fig. 3). The subunit rRNA gene was sequenced from Pleistophora affected skeletal muscle myofibers were frequently hyphessobryconis obtained from 3 zebrafish submitted surrounded by large numbers of intermixed histiocytes by Lab 3. All 3 of these sequences were identical, with and lymphocytes with fewer eosinophilic granular no insertions/deletions nor transitions/transversions. cells, which tracked along the endomysial connective Comparisons of the 1224 bp subset of the sequence tissue and were accompanied by marked endomysial obtained from the neon tetra and the 3 sequences edema that widely separated myofibers. Regenerating obtained from zebrafish showed no insertions/dele- myofibers characterized by enhanced sarcoplasmic tions and 2 transitions: at site 149 (T to C) and at site basophilia and nuclear rowing were frequently adja- 180 (G to A) for an overall paired distance of 0.002. cent to many of the necrotic myofibers (data not shown). Numerous liberated mature spores within the endomysial spaces were surrounded by dense aggre- Transmission gates of histiocytes and lymphocytes (Fig. 3B). Phago- cytized mature spores were occasionally seen within Both moribund zebrafish collected from the group activated histiocytes. exposed to infected neon tetra tissue exhibited massive The parasitic infection in the single fish from Lab 1 muscle infections based on wet mount observations at was limited to the skeletal muscles; however, in the 20 d postexposure. Histological examination of the fish from the other 2 laboratories, spores within phago- remaining fish collected at 30 d showed infections in 5 cytes were also detected in various other organs, of 8 fish. These fish had light infections exhibiting var- including the kidney interstitium, spleen, ovaries, ious stages of development. intestine, and mesenteries. Massive numbers of spores within phagocytes were frequently observed in the connective tissues of the ovary, but never within ova DISCUSSION (Fig. 3G). Aggregates of spores were observed in all layers of the intestine, extending through the serosal Identification and surface and into mesenteric connective tissue. Smooth and cardiac muscle were not affected in any of the fish Pleistophora hyphessobryconis infections in aquar- examined. Both Gram stain methods were effective for ium fishes have been documented for many decades 52 Dis Aquat Org 91: 47–56, 2010

Fig. 3. Danio rerio. Histological sections of zebrafish infected with Pleistophora hyphessobryconis. (A) Numerous sporophorous vesicles with spores and developmental stages within myocytes. (B) Spores in phagocytes associated with chronic infections and myolysis (arrows). (C) Mixed infection with P. hyphessobryconis in muscle and xenomas of Pseudoloma neurophilia in spinal cord (arrows). (D) Meronts (m) and developing (sp) and mature spores in sporophorous vesicles. Some fully developed spores stain 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 53

hyphessobryconis amongst various fishes (Canning et al. 1986). Here we confirmed 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 in small subunit rRNA gene sequence be- tween parasites from the type host (the neon tetra) and from zebrafish. This is consistent with intraspecific variation in this region of the small subunit rRNA for other Pleistophora spp. and related microporidia. The intraspecific pair- wise distance between other available sequences from multiple individuals in GenBank 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, we found an average pairwise interspe- cific distance of 0.198 with all members of the genera Pleistophora, Oviplei- stophora, and Heterosporis with se- quences published in GenBank. The minor, but consistent, differences seen between the 3 zebrafish sequences compared to that of neon tetra might suggest different strains of the parasite. Regardless, the parasite from neon tetra was easily transmitted to ze- Fig. 4. Phylogenetic tree of Pleistophora hyphessobryconis obtained from neon brafish. Our findings agree with previ- tetra and related microsporidia. 1361 bp small subunit rRNA gene sequences ous transmission studies, confirming from 20 microsporidia infecting fish were used to reconstruct phylogeny. Maxi- the broad host specificity of this mi- mum likelihood tree is shown. Branch numbers: maximum likelihood bootstrap crosporidium (Canning et al. 1986). support based on 100 replicates/Bayesian posterior probabilities. names: as recorded in GenBank with new genus designations in parentheses. Mi- It was noted previously that the crosporidian parasite Pseudoloma neurophilia was selected as an outgroup taxa. genus Pleistophora does not form a Scale bar represents number of base substitutions per site monophyletic clade (Nilsen et al. 1998). In fact, ultrastructural charac- terization and molecular analyses of since the original description in 1941 by Schäperclaus the small subunit rRNA gene sequences have resulted (Schäperclaus 1941). Unusually for microsporidia, the in the movement of P. mirandellae (Pekkarinen et al. parasite shows remarkably broad host specificity, 2002) and P. anguillarum (Lom et al. 2000) to new gen- infecting some 20 species of freshwater fishes in 4 era (Ovipleistophora and Heterosporis, respectively). orders (Steffens 1962, Schäperclaus 1991, Lom & Prior ultrastructural descriptions of P. hyphessobryco- Dyková 1992). Among those are zebrafish (Opitz 1942) nis (Lom & Corliss 1967) confirm the placement of this and the dwarf danio Danio nigrofasciatus (Spence et species in the genus Pleistophora as redescribed by al. 2008). Here we found the infection in 3 separate Canning & Nicholas (1980). However, phylogenetic zebrafish research colonies with no known movement analyses of P. hyphessobryconis place it more closely of fish between the facilities. Confirmation of host to both O. mirandellae and H. anguillarum than P. typ- ranges of parasites with limited morphological charac- icalis, the type species described for the genus (Fig. 4). ters (such as Microsporidia) often requires cross trans- Interestingly, unlike the members of Ovipleistophora, mission studies or sequence comparisons. There are this species does not infect oocytes but rather myocytes reports of experimental transmission experiments of P. as does the type species of this genus, P. typicalis. 54 Dis Aquat Org 91: 47–56, 2010

Comprehensive analysis of the genus is beyond the that the fish was subclinically infected for more than scope of this study, but further investigation appears to 1 yr. All of the fish purchased from the vendor had be needed. been removed from Lab 1 more than 7 mo before the infected fish was detected. Even if the fish had not been infected as an embryo but instead at a later time Transmission through accidental cross-contamination with an infected fish from the group purchased from the ven- The 3 facilities had no history of sharing fish and are dor, it indicates that this fish was subclinically infected located in different areas of the United States. Thus, for more than 7 mo. we presume that the infections arose in 3 independent As with other microsporidian infections, it is likely instances by exposure to other species of infected that immune status influences susceptibility of zebra- aquarium fishes. Indeed, the CG1 line was actually fish to Pleistophora hyphessobryconis. Recently, Ram- derived from zebrafish purchased from a pet store say et al. (2009) showed that stress enhances infections (Mizgireuv & Revskoy 2006). The affected fish from of Pseudoloma neurophilia in zebrafish. Infections by Lab 2 had also been purchased from a pet store and the P. hyphessobryconis were widespread in only one maternal parent of the infected fish at Lab 1 had been strain (CG1) in one of the labs with the infection sug- purchased from a vendor that supplied fish to the gesting that this line may be particularly susceptible to commercial pet trade. Transmission of Pleistophora the parasite. The microsporidium was also found in fish hyphessobryconis, ostensibly per os, was achieved by (CG1 and WIK) that were irradiated and thus were placing large numbers of spores in water with fish, a immune compromised. Although immune status likely method that has been used by others (Canning et al. affects the susceptibility and progression of the disease 1986). As with other microsporidia, it is assumed that in zebrafish, apparently immunocompetent fish can infection is initiated by ingestion of spores. Spores become infected as demonstrated by reports of P. released from dead fish are a likely source of infection, hyphessobryconis in zebrafish from aquaria (Opitz but could also be released from the intestines. 1942, Steffens 1962), and experimental infections in Schäperclaus (1941) suggested that spores may be presumably healthy zebrafish reported here. The latter released from the skin or urinary tract of infected fish. were exposed at 15 d old, a time at which fish have Zebrafish spawn frequently within aquaria, and tank innate immunity but adaptive immunity has not fully mates quickly eat available eggs (Lawrence 2007, developed (Lam et al. 2004, Trede et al. 2004). Spence et al. 2008). Although spores were not found within eggs, the massive numbers seen within ovaries of some infected zebrafish suggest that infectious Pathology spores could be released during spawning and would thus be available to fish feeding on eggs or to infect the The infection in zebrafish was consistent with reports next generation of fish. Maternal transmission, includ- from neon tetras and other species (Dyková & Lom 1980, ing true vertical transmission within eggs, has been Canning et al. 1986, Schäperclaus 1991). The most verified or implicated for other microsporidia of fishes remarkable pathological feature of Pleistophora hyphes- (Docker et al. 1997, Kent & Bishop-Stewart 2003, sobryconis is the severe intensity of infection in the Phelps & Goodwin 2008). Schäperclaus (1941) sug- skeletal muscle, with well over half of the myofibers con- gested the possibility of maternal transmission as he taining numerous spores and developmental stages in found infections in 8 d old neon tetras derived from some fish. Parasites within myocytes were not associated infected parents. Once established in zebrafish, it is with inflammation; however, mature spores released conceivable that the infection could be maternally from degenerate myofibers into interstitial spaces were transferred to the next generation. This could even consistently associated with inflammation, and these lib- occur with spores outside the egg as microsporidian erated spores were often engulfed by phagocytes. This is spores are very resistant to chlorine (Ferguson et al. similar to that seen with other intramuscular parasites of 2007). This provides one explanation for the occur- fishes, such as Kudoa thyrsites of salmon (Moran et al. rence of the infection in fish derived from surface- 1999). Likewise, other microsporidia show a similar disinfected eggs. Alternatively, these fish may have sequelae of infection in which spores elicit significant become infected by an unrecognized breach in biose- inflammation only after they are released from their curity protocols. intracellular environment within xenomas (Dyková & The infected fish from Lab 1 was the 13 mo old off- Lom 1980, Kent & Speare 2005). Microsporidian spores spring of a female fish that had been purchased from remain intact within phagocytes, and thus may be trans- an outside vendor and brought into the laboratory. If ported to sites beyond where they originally developed this fish was infected as an embryo, this would mean (Kent et al. 1999). The occurrence of large numbers of Sanders et al.: Pleistophora hyphessobryconis infecting zebrafish in research facilities 55

spores of P. hyphessobryconis within phagocytes in M. Schuster for assistance in translation of German refer- organs other than the skeletal muscle, as seen in the pre- ences, and K. Berkenkamp for histology slide preparation and sent study, has been previously observed (Lom & Dyková technical support. 1992). Spores of microsporidia are Gram positive in his- tological sections (Gardiner et al. 1998, Bruno et al. LITERATURE CITED 2006). Both Lilly-Twort and Accustain Gram stains were very effective in demonstrating spores and particularly Ackermann G, Paw BH (2003) Zebrafish: a genetic model for vertebrate organogenesis and human disorders. Front useful for visualizing individual spores within visceral Biosci 8:d1227–d1253 organs. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Some infected zebrafish exhibited concurrent in- Basic local alignment search tool. 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In: An atlas of protozoan parasites in animal tissues. Armed As suggested for (Kent et al. Pseudoloma neurophilia Forces Institute of Pathology, Washington, DC, p 12–13 2009), the most feasible strategy would be to hold brood Grunwald DJ, Eisen JS (2002) Headwaters of the zebrafish— fish in quarantine and screen them and their progeny emergence of a new model vertebrate. Nat Rev Genet 3: for the infection using a PCR test specific to the para- 717–724 site. Sentinel fish programs and sampling of moribund Guindon S, Delsuc F, Dufayard J, Gascuel O (2009) Estimat- ing maximum likelihood phylogenies with PhyML. Meth- fish are also recommended as a means of surveillance ods Mol Biol 537:113–137 of colony health. Notably, 2 of these cases were from Hill AJ, Teraoka H, Heideman W, Peterson RE (2005) fish that had been in contact with commercial pet store Zebrafish as a model vertebrate for investigating chemical fish. It would be prudent to not mix zebrafish used in re- toxicity. 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Editorial responsibility: Dieter Steinhagen, Submitted: March 8, 2010; Accepted: May 19, 2010 Hannover, Germany Proofs received from author(s): July 3, 2010