Morphological, Molecular, and Phylogenetic Characterization of Nosema ceranae,a Microsporidian Parasite Isolated from the European Honey Bee, Apis mellifera1

YANPING P. CHEN,a JAY D. EVANS,a CHARLES MURPHY,b ROBIN GUTELL,c MICHAEL ZUKER,d

DAWN GUNDENSEN-RINDALe and JEFF S. PETTISa

aUSDA-ARS, Bee Research Laboratory, Beltsville, Maryland 20705, andbUSDA-ARS, Soybean Genomic & Improvement Laboratory, Beltsville, Maryland 20705, and

cInstitute for Cellular and Molecular Biology and Section of Integrative Biology, University of Texas, Austin, Texas 78712, anddRensselaer Polytechnic Institute, New York 12180, and

eUSDA-ARS, Invasive Insect Biocontrol and Behavior Laboratory, Beltsville, Maryland 20705

ABSTRACT. Nosema ceranae, a microsporidian parasite originally described from Apis cerana, has been found to infect Apis mellliferaand is highly pathogenic to its new host. In the present study, data on the ultrastructure of N. ceranae, presence of N. ceranae-specificnucleic acid in host tissues, and phylogenetic relationships with other microsporidia species are described. The ultrastructural featuresindicate that N. ceranae possesses all of the characteristics of the genus Nosema. Spores of N. ceranae measured approximately4.4 � 2.2 mm on fresh smears. The number of coils of the polar filament inside spores was 18–21. Polymerase chain reaction (PCR) signalsspecific for N. ceranae were detected not only in the primary infection site, the midgut, but also in the tissues of hypopharyngeal glands,salivary glands, Malpighian tubules, and fat body. The detection rate and intensity of PCR signals in the fat body were relatively lowcompared with other examined tissues. Maximum parsimony analysis of the small subunit rRNA gene sequences showed that N. ceranaeappeared to be more closely related to the wasp parasite, Nosema vespula, than to N. apis, a parasite infecting the same host.

Key Words. Disease, microorganism, parasitism, phylogeny.

S INCE their first recognition as pathogens in silkworms byNageli (1857), microsporidia have been identified as the

sources of many infectious diseases in vertebrates and inverte-brates, including human, fishes, and insects (Canning, Lom, andDykova 1986; Franzen and Muller 2001; Sprague and Vavra1977; Wasson and Peper 2000; Wittner and Weiss 1999). Of 143genera and over 1,200 microsporidian species described (Witt-ner and Weiss 1999), insects are frequently found to be their pri-mary host. Most of the entomopathogenic microsporidia occur inthe genus Nosema, which has 4150 described species and infectsnearly all taxonomic orders of insect, especially the orders of Le-pidoptera and Hymenoptera (Becnel and Andreadis 1999; Sprague1978).

Nosemosis is a serious disease of adult honey bees, caused byNosema species. Honey bee colonies are frequently infected, andall colony members, including adult worker bees, drones, andqueens, can be infected. Nosema infection occurs mostly throughingestion of spores with food or water. The physical and chemicalconditions of the midgut trigger the germination of spores and thevegetative stage of Nosema begins to grow and multiply insidemidgut cells. Bailey and Ball (1997) showed that 30–50 millionspores could be found inside a bee’s midgut within 2 wk afterinitial infection. Eventually the spores pass out of the bee in itsfeces, providing new sources of the infection through cleaning andfeeding activities in the colonies.

Nosema infections have significant negative impacts on honeybees, causing dysentery, shortened life spans of honey bees, su-persedure of infected queens, and decrease in colony size (Has-sanein 1953; Malone, Giacon, and Newton 1995; Rinderer andSylvester 1978). Nosema ceranae and Nosema apis are two spe-cies of Nosema that are reported to infect the European honey bee,Apis mellifera. For years, nosemosis of the European honey bee

was exclusively attributed to N. apis. Nosema ceranae, a speciesoriginally found in the Asian honey bee, Apis cerana (Fries et al.1996), is now a common infection of European honey bees andis highly pathogenic to its new host (Cox-Foster et al. 2007; Frieset al. 2006; Higes, Martin, and Meana 2006; Higes et al. 2007;Huang et al. 2007; Klee et al. 2007). Chen et al. (2008) demon-strated that N. ceranae was transferred from A. cerana to A. mell-ifera at least a decade ago and is now replacing N. apis as thepredominant microsporidian infection in A. mellifera of the U.S.populations. Although widespread infection by N. ceranae in theU.S. population of A. mellifera has been identified, many biolog-ical features of this parasite in the host A. mellifera remain to beelucidated. To remedy this deficiency, we describe key morpho-logical features of N. ceranae based on light and electron micros-copy, and we use PCR to determine the presence of N. ceranae-specific nucleic acid in different tissues of the infected hosts. Bycomparing the sequences of small subunit rRNA (SSUrRNA)genes of different microsporidia, we construct a phylogenetic treeto determine the genetic relationship of N. ceranae with otherspecies of microsporidia infecting insects.

MATERIALS AND METHODS

Honey bee sample collection. Honey bees were collectedfrom colonies maintained in Beltsville, MD. The abdomens of10 honey bees from each colony were ground up in 2 ml of steriledistilled water. One drop of the homogenate was examined by alight microscope for presence of Nosema spores. When spores ofNosema were observed under the microscope, the remaining por-tion of the homogenized abdomens was used for DNA extractionand PCR assays to determine the species status of Nosema infec-tion. Once the Nosema infection of bee colonies was identified,additional adult bees were collected from those heavily infectedcolonies and stored at � 20 1C for subsequent morphological andmolecular analyses.

Purification of Nosema spores. To obtain purified Nosemaspores, the alimentary tracts of honey bees from Nosema-infectedcolonies were removed individually by grasping the sting withforceps and gently pulling the alimentary tract away from the ab-domen. The midgut was separated from the hindgut and 25 piecesof midgut were crushed in 2 ml of sterile distilled water and fil-

Corresponding Author: Y. Chen, USDA-ARS, Bee Research Labo-ratory, Building 476, BARC-East Beltsville, MD 20705—Telephonenumber: 11 301 504 8749; FAX number: 11 301 504 8736; e-mail:judy.chen@ars.usda.gov

1Disclaimer: Mention of trade names or commercial products in thisarticle is solely for the purpose of providing specific information anddoes not imply recommendation or endorsement by the U.S. Departmentof Agriculture.

142

J. Eukaryot. Microbiol., 56(2), 2009 pp. 142–147r 2009 The Author(s)Journal compilation r 2009 by the International Society of ProtistologistsDOI: 10.1111/j.1550-7408.2008.00374.x

tered through a Corning Netwell (Corning Incorporated Life Sci-ence, Lowell, MA) insert (24 mm diam., 74 mm mesh size) to re-move tissue debris. The filtered suspension was centrifuged at1,500 g for 5 min and the supernatant was discarded. The pelletwas resuspended in 1 ml of sterile water and overlayed very gentlyon a discontinuous 25%, 50%, 75%, and 100% of Percoll (Sigma-Aldrich, St. Louis, MO) gradient from top to the bottom and cen-trifuged twice at 8,000 g for 20 min at 4 1C using a Beckman rotor(SW 28) in a Beckman L8-70M ultracentrifuge (Beckman Instru-ments, Inc., Palo Alto, CA) to collect spores having the same size,shape, and density. The supernatant was discarded and the sporepellet was resuspended in distilled sterile water and collected bycentrifugation. After a final centrifugation at 8,000 g for 10 min at4 1C, the spore pellet was resuspended in distilled sterile water andstored at 4 1C until used. Spore sizes were measured under anEclipse TE 300 light microscope (Nikon, Melville, NY) and pho-tographed with a Nikon Digital Camera (DXM 1200).

Light and electron microscopy. Midguts of adult bees from aNosema-infected colony were dissected out as described aboveand midgut tissue was fixed for 2 h at room temperature by im-mersion in 3% (v/v) glutaraldehyde in 0.05 M sodium cacodylatebuffer, pH 7.0. After overnight incubation in a refrigerator at 4 1C,tissues were washed in a sodium cacodylate buffer rinse, 6 timesover 1 h, postfixed in 2% (w/v) osmium tetroxide in 0.05 M so-dium cacodylate buffer, pH 7.0, for 2 h, dehydrated in ethanol andimbedded in Spurr’s low-viscosity embedding resin. One-micrometer-thick sections were cut on a Reichert/AO Ultracutmicrotome (Leica Inc., Deerfield, IL) with a Diatome (Hatfield,PA) diamond knife. The sections for light microscopy weremounted onto slides, stained with 0.5% (w/v) toluidine blue andphotographed with the same system for spores. Sections for elec-tron microscopy were mounted onto 200-mesh Ni grids, stainedwith 4% (w/v) uranyl acetate and 3% (w/v) lead citrate, andviewed in a H-7000 Hitachi (Tokyo, Japan) microscope at 75 kV.

Tissue dissection. Fifteen adult worker bees collected fromNosema-infected colonies were used for tissue dissection. Tissuesof alimentary canal, Malpighian tubules, muscle, fat body, hypo-pharyngeal gland, and salivary gland were carefully separatedunder a Zeiss SV11 Stereomicroscope (Thornwood, NY) and pho-tographed with a Zeiss AxioCam digital camera. All tissues wererinsed once with 1 � PBS and twice with nuclease-free water toprevent possible contamination and then subjected to subsequentmolecular analysis to determine the presence of N. ceranae-spe-cific nucleic acid in tissues.

DNA isolation, PCR amplification, and DNA sequencing.Dissected tissues were frozen in liquid nitrogen individually andground to a fine powder using a mortar and pestle. The genomicDNA was extracted using the DNAzol DNA purification kit (Inv-itrogen, Carlsbad, CA) following the manufacturer’s protocol.Two pairs of primers specific for N. apis (N-apis-F: 50-ccattgccggataagagagt-30; N-apis-R: 50-cacgcattgctgcatcattgac-30)and N. ceranae (N-ceranae-F: 50-cggataaaagagtccgttacc-30; N-ceranae-R: 50-tgagcagggttctagggat-30) were used in the study asdescribed previously (Chen et al. 2008). The specificity of ampli-fication was confirmed by cloning the purified PCR fragmentsfrom 1.5% low-melting-point agarose gel using Wizard PCR PrepDNA Purification System (Promega, Madison, WI) in pCR 2.1vector (Invitrogen), and sequencing the PCR fragments from bothdirections using M13-forward and M13-reverse primers. The se-quence data was analyzed using the BLAST server at the NationalCenter for Biotechnology Information.

Phylogenetic analysis. The 21 species of microsporidianSSUrRNA sequences with the highest BLAST similarity scoreagainst the complete sequence of the N. ceranae SSUrRNA wereretrieved from GenBank database. The hosts of microsporidianspecies used for phylogenetic analysis were all insects from the

Orders Hymenoptera, Lepidoptera, and Coleoptera. Trachipleis-tophora hominis infecting Homo sapiens was used as an outgroupto root the phylogenetic tree. Sequences were aligned using Meg-Align (DNASTAR Lasergene software program, Madison, WI)and sequences that could not be aligned unambiguously at both30- and 50-ends were truncated. The percentage identity anddivergence of sequences between equivalent microsporidianSSUrRNA was generated by the MegAlign. Aligned sequencesof 20 microsporidia species and the outgroup were imported intothe phylogenetic analysis program PAUP 4.03 (Sinauer Associ-ates, Sunderland, MA). Maximum Parsimony under a heuristicsearch with random stepwise addition and TBR branch swappingwas used to construct the phylogenetic trees. Phylogenies wereassessed by a 1,000 bootstrap replication.

RESULTS

Nosema ceranae infection was found in adult bees of A. mell-ifera collected in Beltsville, MD. When the abdomens of infectedbees were crushed in water, a large numbers of mature sporeswere released, although most infected bees did not exhibit overtbehavior and morphological signs of infection. The samples ex-amined in this study were exclusively N. ceranae-positive: noneof the PCR reactions using N. apis-specific primers yielded anyproduct (data not shown).

Light microscopy revealed that fresh N. ceranae spores wereoval or rod shaped, varied in size with a length of 3.9–5.3 mm(mean � SE 5 4.4 � 0.41 mm) and a width of 2.0–2.5 mm (mean-SE 5 2.2 � 0.09 mm) (N 5 50) (Fig. 1). Observation of Nosema-infected midgut tissue showed that mature spores not only accu-mulate in midgut epithelial cells but also are released into the gutlumen (Fig. 2, 3).

Ultrastructural studies showed that different developmentalstages, including meronts, sporonts, sporoblasts, and maturespores, are found in the midgut epithelial cells. Meronts, the ear-liest developmental stage, had two nuclei in diplokarytic arrange-ment and were bound by a plasma membrane in direct contactwith host cytoplasm (Fig. 4). Sporonts were elongated and oval inshape with dense cytoplasm and no discernible internal structures(Fig. 5). Sporoblasts were generally smaller than sporonts with amore clearly defined cell wall and two nuclei (Fig. 4). Electronmicrographs of longitudinal sections of mature spores showed thatthe spore wall consisted of a dense exospore, 48–53 nm thick, anda lucent layer endospore, and that the sporoplasm was enclosed by

Fig. 1. Nosema ceranae spores. Light micrograph of oval- to rod-shaped spores of N. ceranae after Percoll purification. Scale bar 5 10 mM.

143CHEN ET AL.—CHARACTERIZATION OF NOSEMA CERANAE

a plasma membrane (Fig. 6). The anchoring disk was located inthe anterior pole of the spore. The lamellate polaroplast occupiedthe anterior part of the spore adjacent to the anchoring disk (Fig.6). A vacuole was located in the posterior end of the spore and notprominent. Two nuclei in diplokaryotic arrangement were closelyapposed in the central region of the spore between the polaroplastand the posterior vacuole and the polar filaments were arranged in18–21 isofilar coils in two rows (Fig. 7). When a spore had anextruded polar tube, the posterior vacuole swelled and becamevery prominent inside the spore (Fig. 8).

The PCR assays revealed that N. ceranae-specific nucleic acidwas detected in 100% of the alimentary canals, Malpighian tu-bules, and hypopharyngeal glands, in 87% of the salivary glands,and in 20% of the fat bodies (N 5 15). No N. ceranae-specificPCR signal was detected in the muscle tissue examined (lane 5,Fig. 9).

The complete DNA sequences of the rRNA gene is 4,475 bp.The G1C content of the SSUrRNA cistron at positions 1–1,259was 36.46%. The internal transcribed spacer (ITS) region con-sisted of a 39-bp sequence and was located between nucleotides1260 and 1298. The DNA sequence of LSUrRNA, located at the30-end between nucleotides 1299 and 3828, contained 2,530 bpand was 32.86% G1C.

The percent of SSUrRNA sequence identity revealed that N.ceranae shared the highest degree of sequence identity (97.5%)with N. vespula and was the most distantly related to Nosemaplutellae with 19.3% sequence divergence among all micros-poridia included in this study. Our phylogenetic tree of 20 mi-crosporidian taxa contains two distinct clades (Fig. 10). One cladeincludes Vairimorpha imperfecta and some species of the ‘‘true’’Nosema group, a group of lepidopteran Nosema species closelyrelated to Nosema bombycis (Baker et al. 1994). Nosema ceranae,along with several non-lepidoteran Nosema species and true Nose-ma species forms another clade (Fig. 10). Within this latter clade,

Fig. 2, 3. Cross-section of the midgut showing spores. 2. Spores ac-cumulated in midgut lumen. 3. Epithelial cells of the midgut infected withNosema ceranae. Arrows indicate infected epithelial cells with tightlypacked parasites.

Fig. 4, 5. Epithelial cells infected with different developmental stagesof Nosema ceranae. The developmental stages include meront (M), spo-ront (ST), sporoblast (SB), and mature spore (MS). MB, membrane of theinfected host cell; ES, empty shell of the hatched spore.

144 J. EUKARYOT. MICROBIOL., 56, NO. 2, MARCH–APRIL 2009

N. ceranae is most closely related to N. vespula with 80% boot-strap support, and was distantly related to N. apis.

DISCUSSION

The transfer of N. ceranae from its described original host,A. cerana, to a possible novel host, A. mellifera, adds a new di-mension to the biological and epidemiological aspects of this par-asite. Experimental infection of A. mellifera by N. ceranaeconducted by Higes et al. (2007) clearly showed that this para-site is highly pathogenic to its new host and poses a serious threatto the beekeeping industry.

The morphological and molecular characterization of N. cer-anae in Asian honey bees was conducted by Fries et al. (1996).Later, Fries et al. (2006) reported the natural infection N. ceranaein European honey bees. However, many morphological details ofspores such as types and sizes of spores in a dense spore purifi-cation and the morphology at the different developmental stagesof spores in midgut epithelium cells in naturally infected hostsremained to be demonstrated. Our observation with light micros-copy showed that spores of N. ceranae from European honey beesare oval shaped and rather uniform in shape. The electron mi-croscopy indicates that N. ceranae contains all of the ultrastruc-tural characteristics of the genus Nosema (Larsson 1986):diplokaryotic nuclei present in all developmental stages; a longflexible polar filament that appears in the mature spores; meronts,the earliest stages in the life cycle of the parasite, which are indirect contact with host cell cytoplasm; mature spores that arebounded by a thickened wall consisting of electron-dense exo-spore and electron-lucent endospore layers; and the thickness ofexospore is 48–52 nm. The number of polar filament coils is animportant taxonomic criterion to differentiate different species ofNosema (Burges, Canning, and Hulls 1974). The number of coilsof polar filament inside N. ceranae spores measured by us was18–21, overlapping with the range of 20–23 coils reported byFries et al. (1996), which is much smaller than the 430 coils re-corded for N. apis (Fries 1989; Liu 1984).

Although not all examined tissues showed visible signs ofpathological changes, PCR assay followed by sequencing analy-sis showed that N. ceranae-specific PCR signals are not restrictedto the midgut tissue but spread to other tissues, including theMalpighian tubules, hypopharyngeal glands, salivary glands, andfat bodies. The presence of the signal suggests that these tissuesmay be infected, as was determined microscopically for Nosemabombi in a bumble bee (Fries et al. 2001). However, microscopicstudies of N. ceranae in A. mellifera tissues, which would verifythe infections, remain to be conducted. The detection of N. cer-anae-specific PCR signals in both hypopharyngeal and salivaryglands suggests that royal jelly, the secretion of hypopharyngealand salivary glands of worker bees used to feed the queen andlarvae, could be another vehicle for horizontal fecal–oral andfood-borne transmission of the parasite in the bee colonies. Aweak PCR signal specific for N. ceranae detected in the fat bodytissue suggests a low parasite load, arguing that fat body tissue isnot a primary target for N. ceranae infection even though fat-bodytissues are one of the primary sites for microsporidian infection.Infection of fat bodies causes formation of whitish cysts and theinfected gut becomes swollen and whitish as a result of impairedfat metabolism in many other insects (Sokolova et al. 2006).

Although honey bee colonies with reduced longevity, de-creased population size, higher autumn/winter colony loss, and/or reduced honey production are often reported to be associatedwith the presence of N. ceranae, the disease signs such as dysen-tery or crawling behavior or milky white coloration of gut, that areusually associated with N. apis infection, has never been describedin N. ceranae-infected bees (Fries et al. 2006). The absence ofthese disease symptoms in N. ceranae-infected A. mellifera mightreflect the absence or low intensity of N. ceranae specific PCRsignals in the muscles and fat bodies of infected bees, respec-tively. It is not clear why N. ceranae has different pathologicaleffects on the host A. mellifera compared with N. apis. Furtherstudies are warranted to ascertain the pathogenesis of both para-sites in the A. mellifera.

The sequences of the rRNA operon have been widely used as amolecular marker for detection of microsporidian infection,differentiation of closely related species, and estimation of phylo-genetic relationship among microsporidia. The organization of therRNA gene of N. ceranae contains one SSUrRNA gene at the

Fig. 6–8. Electron-micrographs of longitudinal sections of spores ofNosema ceranae. 6. Micrograph showing anchoring disk (AD), polaroplast(P), posterior vacuole (PV), polar filament (PF). 7. Micrograph showingendospore (EN), exospore (EX), plasma membrane (PM), nucleus (N), 20–22 isofilar coils of the PFs. 8. A spore with an extruded polar filament(EPF). Note the more conspicuous PV.

145CHEN ET AL.—CHARACTERIZATION OF NOSEMA CERANAE

50 end, one LSUrRNA gene at 30 end, and an ITS located betweenthe SSUrRNA and LSUrRNA genes. Parallel comparison of therRNA gene sequences of N. ceranae and N. apis showed a se-quence identity of 92.7% for SSUrRNA, 91.9% for LSUrRNA,and 48.5% for ITS. Although N. apis and N. ceranae infect the

same host and share similarities in sequences of rRNA gene, ourphylogenetic analysis based on sequences of SSUrRNA showedthat N. apis is not the closest relative of N. ceranae. The samephylogenetic relationshop of N. apis and N. ceranae was also il-lustrated in earlier works for characterization of the microsporidian

Fig. 9. Detection of Nosema ceranae by polymerase chain reaction (PCR) amplification of nucleic acids from different tissues. DNA was extractedfrom tissues and examined for the presence of N. ceranae-signal by PCR method and electrophoresis. The gel numbers 1–6 indicate the hypopharyngealgland, salivary gland, alimentary canal, Malpighian tubules, muscle, and fat body, respectively; N indicates negative control and P indicates positivecontrol. The size of PCR fragments is indicated on the right of the gel.

Fig. 10. Phylogenetic tree of microsporidia. Phylogenetic tree of microsporidia infecting insects based on the sequences of the small subunit rRNAgene and constructed by maximum parsimony analysis under a heuristic search. Trachipleistophora hominis-infecting Homo sapiens was used as anoutgroup. The non-lepidopteran Nosema species are indicated by an asterisk. The reliability of the tree topology is shown by the bootstrap values locatedon the tree branches.

146 J. EUKARYOT. MICROBIOL., 56, NO. 2, MARCH–APRIL 2009

species from mosquitoes and spider mites (Becnel et al. 2002;Muller et al. 2000). Within the same clade, N. ceranae appears tobe more closely related to N. vespula, a parasite infecting wasps,with 80% bootstrap support. Nosema apis seems to have branchedoff earlier and is most closely linked to N. bombi, a parasite in-fecting bumble bees.

The comparative analysis of rRNA sequences indicated thatribosomal RNA is conserved and maintains a similar secondaryand tertiary structure for all types of organisms (Gutell, Noller,and Woese 1986a; Gutell et al. 1986b). While the microsporidianrRNAs contain some of the characteristic features found in thevast majority of the eukaryotic rRNAs, the 16S-like and 23S-likerRNAs of N. ceranae are very unusual. They lack many of thestructural elements present in other nuclear-encoded eukaryoticrRNAs, and they are significantly shorter in length. For examplethe Saccharomyces cerevisiae 16S-like and 23S-like rRNAs areapproximately 1,800 and 3,550 nucleotides in length, the N. cer-anae 16S-like and 23S-like rRNAs are 1,259 and 2,530 nucleo-tides in length, respectively. To determine how the reduction insize of rRNA contributes to the life cycle of the intracellular par-asite in the host, further studies are needed.

As with many other new and emerging pathogens, we are justbeginning to scratch the surface of understanding how N. ceranaeadopt and establish infection in the new host. Genomic and bio-chemical characterizations of N. ceranae are currently in progressto study the roles of parasite genetic variability, host physiologicalconditions, and host immune status in the course of infection anddisease.

ACKNOWLEDGMENTS

We would like to thank Michele Hamilton, Bart Smith, andAndrew Ulsamer for their excellent technical assistance. Thework was supported in part by the 2006 California State Bee-keepers’ Association (CSBA) research fund. R. Gutell was sup-ported by the National Institutes of Health (GM067317) and theWelch Foundation (F-1427).

LITERATURE CITED

Bailey, L. & Ball, B. V. 1997. Honey Bee Pathology. 3rd ed. AcademicPress, London. p. 59–76.

Baker, M. D., Vossbrinck, C. R., Maddox, J. V. & Undeen, A. H. 1994.Phylogenetic relationships among Vairimorpha and Nosema species(Microspora) based on ribosomal RNA sequences. J. Invertebr. Pathol.,61:100–106.

Becnel, J. J. & Andreadis, T. G. 1999. Microsporidia in insect. In: Wittner,M. & Weiss, L. M. (ed.), The Microsporidia and Microsporidiosis.ASM Press, Washington, DC. p. 447–501.

Becnel, J. J., Jeyaprakash, A., Hoy, M. A. & Shapiro, A. 2002. Morpho-logical and molecular characterization of a new microsporidian speciesfrom the predatory mite Metaseiulus occidentalis (Nesbitt) (Acari,Phytoseiidae). J. Invertebr. Pathol., 79:163–172.

Burges, H. D., Canning, E. U. & Hulls, J. K. 1974. Ultrastructure of Nose-ma oryzaephili and the taxonomic value of the polar filament. J. In-vertebr. Pathol., 23:135–139.

Canning, E. U., Lom, J. & Dykova, I. 1986. The Microsporidia of Ver-tebrates. Academic, New York. 289 p.

Chen, Y. P., Evans, J. D., Smith, J. B. & Pettis, J. S. 2008. Nosema ceranaeis a long-present and wide-spread microsporidian infection of the Eu-ropean honey bee (Apis mellifera) in the United States. J. Invertebr.Pathol., 97:186–188.

Cox-Foster, D. L., Conlan, S., Holmes, E., Palacios, G., Evans, J. D., Mo-ran, N. A., Quan, P. L., Briese, T., Hornig, M., Geiser, D. M., Martin-son, V., van Engelsdorp, D., Kalkstein, A. L., Drysdale, A., Hui, J.,Zhai, J., Cui, L., Hutchison, S. K., Simons, J. F., Egholm, M., Pettis, J.S. & Lipkin, W. I. 2007. A metagenomic survey of microbes in honey-bee colony collapse disorder. Science, 318:283–287.

Franzen, C. & Muller, A. 2001. Microsporidiosis: human diseases and di-agnosis. Microbes Infect., 3:389–400.

Fries, I. 1989. Observation on the development and transmission ofNosema apis Z. in the ventriculus of the honey bee. J. Api. Res., 28:107–117.

Fries, I., Feng, F., Silva, A. D., Slemenda, S. B. & Pieniazek, N. J. 1996.Nosema ceranae n. sp. (Microspora, Nosematidae), morphological andmolecular characterization of a microsporidian parasite of the Asianhoney bee Apis cerana (Hymenoptera, Apidae). Eur. J. Protistol.,32:356–365.

Fries, I., Martın, R., Meana, A., Garcıa-Palencia, P. & Higes, M. 2006.Natural infections of Nosema ceranae in European honey bees. J. Api.Res., 45:230–233.

Fries, I., Ruijter, A. de, Paxton, R. J., da Silva, A. J., Slemeda, S. B. &Pieniazek, N. J. 2001. Molecular characterization of Nosema bombi(Microsporidia: Nosematidae) and a note on its sites of infection inBombus terrestris (Hymenoptera: Apidae). J. Api. Res., 40:91–96.

Gutell, R. R., Noller, H. F. & Woese, C. R. 1986a. Higher order structurein ribosomal RNA. EMBO J., 5:1111–1113.

Gutell, R. R., Weiser, B., Woese, C. R. & Noller, H. F. 1986b. Compar-ative anatomy of 16S-like ribosomal RNA. Prog. Nucleic Acid Res.Mol. Biol., 32:155–216.

Hassanein, M. H. 1953. The influence of Nosema apis on the larval hon-eybee. Ann. Appl. Biol., 38:844–846.

Higes, M., Martın, R. & Meana, A. 2006. Nosema ceranae, a new mi-crosporidian parasite in honey bees in Europe. J. Invert. Pathol., 92:93–95.

Higes, M., Garcıa-Palencıa, P., Martın-Hernandez, R. & Meana, A. 2007.Experimental infection of Apis mellifera honeybees with Nosema cer-anae (Microsporidia). J. Invertebr. Pathol., 94:211–217.

Huang, W. F., Jiang, J. H., Chen, Y. W. & Wang, C. H. 2007. A Nosemaceranae isolate from the honeybee Apis mellifera. Apidologie, 38:30–37.

Klee, J., Besana, A. M., Genersch, E., Gisder, S., Nanetti, A., Tam, D. Q.,Chinh, T. X., Puerta, F., Ruz, J. M., Kryger, P., Message, D., Hatjina, F.,Korpela, S., Fries, I. & Paxton, R. J. 2007. Widespread dispersal of themicrosporidian Nosema ceranae, an emergent pathogen of the westernhoney bee, Apis mellifera. J. Invertebr. Pathol., 96:1–10.

Larsson, R. 1986. Ultrastructure, function, and classification of micros-poridia. In: Corliss, J. O. & Patterson, D. J. (ed.), Progress in Protisto-logy. vol. 1. Biopress, Bristol, U.K., 325–390.

Liu, T. P. 1984. Ultrastructure of the midgut of the worker honey Apismellifera heavily infected with Nosema apis. J. Invertebr. Pathol.,44:103–105.

Malone, L. A., Giacon, H. A. & Newton, M. R. 1995. Comparison of theresponses of some New Zealand and Australian honey bees (Apis mell-ifera L) to Nosema apis Z. Apidologie, 26:495–502.

Muller, A., Trammer, T., Chioralia, G., Seitz, H. M., Diehl, V. & Franzen,C. 2000. Ribosomal RNA of Nosema algerae and phylogenetic rela-tionship to other microsporidia. Parasitol. Res., 86:18–23.

Nageli, K. W. 1857. Uber die neue Krankheit der Seidenraupe und ve-rwandte Organismen. Bot. Z., 15:760–761.

Rinderer, T. E. & Sylvester, H. A. 1978. Variation in response to Nosemaapis, longevity, and hoarding behavior in a free-mating population ofthe honey bee. Ann. Entomol. Soc. Am., 71:372–374.

Sokolova, Y. Y., Kryukova, N. A., Glupov, V. V. & Fuxa, J. R. 2006.Systenostrema alba Larsson 1988 (Microsporidia, Thelohaniidae) in thedragonfly Aeshna viridis (Odonata, Aeshnidae) from South Siberia:morphology and molecular characterization. J. Eukaryot. Microbiol.,53:49–57.

Sprague, V. 1978. Characterization and composition of the genus Nosema.Misc. Publ. Entomol. Soc. Am., 11:5–16.

Sprague, V. & Vavra, J. 1977. Systematics of the microsporidia. In: Bulla,L. A. & Cheng, T. C. (ed.), Comparative Pathobiology. Plenum, NewYork. p. 31–335.

Wasson, K. & Peper, R. L. 2000. Mammalian microsporidiosis. Vet. Pa-thol., 37:113–128.

Wittner, M. & Weiss, L. M. 1999. The Microsporidia and Microsporidosis.ASM Press, Washington, DC. 553 p.

Received: 12/21/07, 04/02/08, 05/22/08, 07/08/08; accepted: 10/08/08

147CHEN ET AL.—CHARACTERIZATION OF NOSEMA CERANAE