APPLIED AND ENVIRONMENTAL MICROBIOLOGY, May 2010, p. 3032–3038 Vol. 76, No. 90099-2240/10/$12.00 doi:10.1128/AEM.03097-09Copyright © 2010, American Society for Microbiology. All Rights Reserved.

Five-Year Cohort Study of Nosema spp. in Germany: Does ClimateShape Virulence and Assertiveness of Nosema ceranae?�†

Sebastian Gisder,1 Kati Hedtke,1 Nadine Mockel,1 Marie-Charlotte Frielitz,1Andreas Linde,2 and Elke Genersch1*

Institute for Bee Research, Friedrich-Engels-Str. 32, D-16540 Hohen Neuendorf,1 and FH Eberswalde,Applied Ecology and Zoology, Alfred-Moller-Str. 1, D-16225 Eberswalde,2 Germany

Received 22 December 2009/Accepted 1 March 2010

Nosema ceranae and Nosema apis are two fungal pathogens belonging to the phylum Microsporidia andinfecting the European honeybee, Apis mellifera. Recent studies have suggested that N. ceranae is more virulentthan N. apis both at the individual insect level and at the colony level. Severe colony losses could be attributedto N. ceranae infections, and an unusual form of nosemosis is caused by this pathogen. In the present study,data from a 5-year cohort study of the prevalence of Nosema spp. in Germany, involving about 220 honeybeecolonies and a total of 1,997 samples collected from these colonies each spring and autumn and analyzed viaspecies-specific PCR-restriction fragment length polymorphism (RFLP), are described. Statistical analysis ofthe data revealed no relation between colony mortality and detectable levels of infection with N. ceranae or N.apis. In addition, N. apis is still more prevalent than N. ceranae in the cohort of the German bee population thatwas analyzed. A possible explanation for these findings could be the marked decrease in spore germination thatwas observed after even a short exposure to low temperatures (�4°C) for N. ceranae only. Reduced or inhibitedN. ceranae spore germination at low temperatures should hamper the infectivity and spread of this pathogenin climatic regions characterized by a rather cold winter season.

Microsporidia are highly evolved fungi with an obligatelyintracellular parasitic lifestyle (14, 34). They are common par-asites of insects and other invertebrates but are also known asparasites of vertebrates, including humans (8, 10, 47). To date,more than 160 genera and almost 1,300 species of microspo-ridia have been described in the literature (14), revealing agreat diversity of morphology and life cycle strategies withinthis phylum. The common characteristics that qualify an or-ganism as a microsporidian are that outside the host cell itexists only as metabolically inactive spores and that infection ofa host cell involves spore germination, i.e., extrusion of a spe-cialized structure, the polar tube, which pierces the host celland inoculates infective sporoplasm directly into the cytoplasmof the host cell (9).

For honeybees, two species of microsporidia have been de-scribed: Nosema apis and Nosema ceranae (23, 49). Originally,it was assumed that N. apis was a pathogen specific for theEuropean honeybee, Apis mellifera, causing nosemosis, whileN. ceranae was specific for the Asian honeybee, Apis cerana.However, early cross-infection experiments demonstrated thatN. apis can be infective for A. cerana and N. ceranae cansuccessfully infect A. mellifera (21). The fact that N. ceranaecan infect A. mellifera under natural conditions as well becameevident when in 2005 N. ceranae was isolated from diseasedhoneybees (A. mellifera) in Taiwan (31) and was found incollapsing A. mellifera colonies in Spain (30). Many studies on

the incidence of N. ceranae in A. mellifera were initiated due tothese findings; they revealed a worldwide distribution of N.ceranae in A. mellifera populations (11, 12, 24, 30, 32, 33, 40, 42,48). Experimental studies suggested that N. ceranae is highlyvirulent for A. mellifera (27), presumably due to immune sup-pression, which could be observed only after N. ceranae infec-tion, not after N. apis infection (3). In the field, N. ceranaecauses an unusual form of nosemosis, which led, and still leads,to severe colony losses in Spain (28, 37). One explanation forthe higher virulence of N. ceranae in the field could be thebetter adaptation of N. ceranae than of N. apis to elevatedtemperatures (18, 36), indicating that N. ceranae might be apathogen whose spread and assertiveness could be influencedby climate change.

Over the past decade, beekeepers in Europe and NorthAmerica reported dramatic increases in colony losses, bothduring the season and over the winter (1, 2, 43, 46). The mainculprits for these increases in colony mortality, among thepathogens affecting honeybee vitality, are viruses (6, 7, 15, 35)and Nosema ceranae (28, 37). While colony losses in the UnitedStates (termed colony collapse disorder [CCD] [45]) did notcorrelate with N. apis or N. ceranae infection (12, 15), colonymortality in Spain could clearly be attributed to N. ceranaeinfection (28, 37), and it was suggested that N. ceranae-inducedcolony collapse is not restricted to Spain but is at least aEurope-wide phenomenon. In Germany, beekeepers experi-enced dramatic overwintering losses in the winter of 2002 to2003, and they have reported an increase in overwinteringmortality since then. To evaluate whether or not N. ceranae canbe correlated with these losses, we conducted a cohort studyover 5 years involving 220 colonies in the northeastern part ofGermany, and we determined colony mortality and the inci-dences and prevalences of the two Nosema species. We also

* Corresponding author. Mailing address: Institute for Bee Re-search, Friedrich-Engels-Str. 32, D-16540 Hohen Neuendorf, Ger-many. Phone: 49 (0) 3303 293833. Fax: 49 (0) 3303 293840. E-mail:elke.genersch@rz.hu-berlin.de.

† Supplemental material for this article may be found at http://aem.asm.org/.

� Published ahead of print on 12 March 2010.

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analyzed individual bees for Nosema infection and Nosemaspores for germination capacity. The implications of our re-sults are discussed below.

MATERIALS AND METHODS

Bee samples and field survey. A cohort of 220 colonies kept in 22 apiaries (10randomly selected colonies per apiary) and managed by hobbyist beekeepers inthe northeastern part of Germany (see Fig. S1 in the supplemental material)were monitored for Nosema infection between spring 2005 and spring 2009. Thecolonies selected for the survey were closely monitored by a professional beeinspector twice a year for the duration of the study without introducing anychanges in the beekeeping practices of the beekeepers. Each March, about 100dead bees were collected from the bottom board of each colony (representing thebees that had died over the winter), and live bees were collected from the wintercluster (representing the bees that had survived the winter). Qualitative Nosemadetection and differentiation did not differ between these two samples for a givencolony; therefore, the bees collected from the bottom boards were used in thecourse of the study. At the end of September/beginning of October, around 100live adult bees were collected from each colony (representing the bees that wereraised for overwintering). The collected bees were frozen at �20°C and storeduntil analysis. Overwintering success, survival during the summer season, and thepresence of symptoms of nosemosis were recorded individually for each colony.None of the monitored colonies showed clinical symptoms of nosemosis duringthe study. Lost colonies were replaced by colonies from the same apiary, pref-erentially by nuclei made from those colonies in the previous year. Since 220colonies were observed throughout the study, the sampling of the monitoredcolonies and of nuclei replacing lost colonies at nine different time points be-tween 2005 and 2009 resulted in a total of 1,997 samples analyzed (see Tables 1and 2).

Detection of Nosema spp. Qualitative microscopic diagnosis of Nosema sporeswas performed according to the method described in the Manual of DiagnosticTests and Vaccines for Terrestrial Animals, published by the Office Internationaldes Epizooties (OIE), the World Organization for Animal Health (38a). Briefly,20 abdomens from each sample were homogenized together in 2 ml double-distilled H2O (ddH2O) and were checked by light microscopy (magnification,�400) for the presence of microsporidian spores (41). The rather moderatesampling size does not allow the detection of the odd infected bee in the colonybut does allow the detection of an infection level above �15% at the 5%significance level (22), which can be considered biologically relevant (28).

Microscopically positive homogenates were transferred to a 1.5-ml reactiontube and were thoroughly homogenized with a 3-mm-diameter tungsten carbidebead (Qiagen) in a mixer mill (Retsch) for 30 s at 30 Hz. Subsequently, thehomogenate was centrifuged at 16,000 � g for 3 min, and DNA was extractedfrom the pellet using standard methods according to the manufacturer’s proto-cols (Plant DNA extraction kit; Qiagen). The extracted DNA was resuspended in50 �l elution buffer (Qiagen) and was frozen at �20°C until differentiation.

From each Nosema-positive bee sample, individual bees were dissected undera dissecting microscope (magnification, �10) under sterile conditions. The tis-sues (gut, malpighian tubules, fat body, hypopharyngeal glands, and brain) werecarefully isolated from individual bees by using fresh forceps for each organ toprevent contamination. The isolated organs were washed twice in 1� phosphate-buffered saline (PBS) and nuclease-free water to wash off any potentially con-taminating hemolymph. Subsequently, DNA was extracted from the tissues asdescribed above. The extracted DNA was resuspended in 50 �l elution buffer(Qiagen) and was frozen at �20°C until differentiation.

Differentiation of Nosema spp. via PCR-restriction fragment length polymor-phism (PCR-RFLP). A region of the 16S rRNA gene that is conserved for N. apisand N. ceranae (33) was selected for primer design using MacVector, version 6.5(Oxford Molecular). Primers nos-16S-fw (5�-CGTAGACGCTATTCCCTAAGATT-3�; positions 422 to 444 in GenBank accession no. U97150 [25a]) andnos-16S-rv (5�-CTCCCAACTATACAGTACACCTCATA-3�; positions 884 to909 in GenBank accession no. U97150 [25a]) were used to amplify ca. 486 bp ofthe partial 16S rRNA gene.

PCR analysis was performed in a final volume of 25 �l containing 5 �l oftemplate DNA (extracted from pelleted spores or individual organs), 2.5 �l of10� Qiagen PCR buffer, 2.5 mM MgCl2, 200 �M each deoxynucleoside triphos-phate (dNTP) (Qiagen), 0.625 U Taq polymerase (Qiagen), and 0.5 �M eachforward and reverse primer. PCR parameters for amplification were as follows:an initial DNA denaturation of 5 min at 95°C; 45 cycles of 1 min at 95°C, 1 minat 53°C, and 1 min at 72°C; and a final extension step at 72°C for 4 min.Amplification products (5 �l DNA) were electrophoresed on 1.1% agarose gels

(1� Tris-borate-EDTA [TBE]), stained with ethidium bromide, and visualizedunder UV light. A commercial 100-bp ladder (Peqlab) was used as a size marker.For each PCR, positive (reference N. apis and N. ceranae DNA extracts) andnegative (ddH2O) controls were run along with DNA extracts of isolates astemplates.

To differentiate between the species N. apis and N. ceranae, discriminatingrestriction endonuclease sites present in the PCR amplicon were used (33). Therestriction endonuclease PacI provides one unique digestion site for N. ceranae,while the enzyme NdeI digests only N. apis. MspI digests both N. apis and N.ceranae and was used as a control for successful restriction digestion of PCRproducts.

The predicted restriction fragments produced from the digestion of the PCRamplicons are illustrated in Fig. S2 in the supplemental material. Ampliconswere digested with MspI/PacI and with MspI/NdeI (New England Biolabs) in tworeactions at 37°C for 3 h in order to analyze and confirm the presence of eachNosema species in each sample. Digestions were performed in a 12.5-�l volumewith 7 �l of the amplified DNA and 1.5 U of each enzyme. For the reactions withMspI and NdeI, 1� NEBuffer 4 (provided by NEB with NdeI) was used as abuffer. For the reactions with MspI and PacI, 1� NEBuffer 1 plus bovine serumalbumin (BSA) (provided by NEB with PacI) was used. Fragments were sepa-rated in a 3% NuSieve agarose gel (Cambrex Bio Science) in 1� TBE buffer witha 20-bp ladder as a size marker at 110 V for 1 h 30 min. Gels were stained withethidium bromide and visualized under UV light.

Smear preparations for detection of developmental stages of Nosema spp. Forthe microscopic detection of developmental stages of Nosema spp. in smearpreparations of bee tissue, adult bees infected with Nosema ceranae or Nosemaapis were used. Brains, hypopharyngeal glands, malpighian tubules, fat bodies,and midguts were carefully isolated from individual bees as described above.Pieces of these tissues (area, 2 mm2) were placed on a microscopic slide for crushpreparations using coverslips and the back of a pencil. After removal of thecoverslips, the crushed tissue was air dried, fixed with 100% methanol, and airdried again prior to staining with Giemsa solution (1:10 Fluka Giemsa stain,modified solution) for 10 min. Stained tissues were rinsed with tap water, airdried, and embedded with Entellan (Merck). Microscopy analysis was performedat 1,000-fold magnification using a stereomicroscope (Leica).

In vitro germination of Nosema spores. Bee samples that tested positive foreither N. apis or N. ceranae were used for isolation of the respective spores. Theprocedure for spore accumulation and purification was modified from themethod of Chen et al. (13). Alimentary tracts of 20 individual bees were removedby snatching the sting with forceps and gently pulling the hindgut and the midgutout of the abdomen. The sting was cut with a scalpel, and the alimentary tract wastransferred to a 1.5-ml reaction tube and crushed in a final volume of 1.5 mlsterile double-distilled water in a mixer mill (Retsch) with a 3-mm-diametertungsten carbide bead (Qiagen) for 30 s at 30 Hz. The homogenate was filteredthrough a nylon cell strainer (Falcon) with a 100-�m mesh diameter. The filtratewas gently overlaid on a 90% Percoll (Sigma-Aldrich) cushion in a 15-ml reactiontube and was centrifuged twice at 15,000 � g for 45 min at 20°C in a Eppendorf5810 R centrifuge using an F34-6-38 rotor. The small but dense band just abovethe bottom of the tube, which contained purified spores, was aspirated with asyringe and a Sterican 0.8-mm by 120-mm needle (Braun) and was transferred toa 15-ml reaction tube. Spores were washed three times with 8 volumes of double-distilled water and were centrifuged at 6,500 � g for 10 min at 20°C. After a finalcentrifugation step, the supernatant was removed, the pellet was resuspended in500 �l AE buffer (Qiagen), and the spore concentration was determined using ahemocytometer. The identity of the spores was verified by PCR-RFLP as de-scribed above. Aliquots of purified N. apis or N. ceranae spores (20 �l; 2E�8spores per ml) were spotted onto glass slides, air dried, and kept at differenttemperatures for different periods of time. Subsequently, germination was trig-gered by the addition of 30 �l of 0.1 M sucrose in PBS buffer to the air-driedspores (39), a procedure that mimics the natural conditions for the germinationof environmental spores. Representative results obtained with freshly isolated,dried, and germinated spores (�22°C) or after storage of the dried spores at 4°Cfor 4 days are shown in Fig. 4. These experiments were repeated 10 times withdifferent spore preparations (n � 10).

Data evaluation and statistical analysis. We compared surviving and collapsedcolonies by using nonparametric chi-square tests, because the basal assumptionsof parametric tests (i.e., normality and constant variance) were not satisfied. Thedistribution of colony losses differed between the years and between the seasons(see Tables 1 and 2); therefore, the data sets were analyzed separately. Chi-square tests were performed by comparing the infection statuses of the survivingcolonies with those of the collapsed colonies. A P value of �0.05 was consideredsignificant.

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RESULTS

Incidences of Nosema apis and Nosema ceranae. During theentire period of our study, we observed the seasonality ofNosema-positive colonies that was described previously (20)(Fig. 1). The proportion of samples positive for Nosema spp. inthe cohort monitored was always higher in the spring than inthe autumn. In the spring, the proportion of Nosema-positivesamples ranged from 22.4% (2007) to 35.4% (2008). In theautumn, the highest prevalence of Nosema spp. was detected in2005 (12.7%) and the lowest in 2008 (5.2%).

Since recent studies had suggested the replacement of N.apis by N. ceranae in Europe (33, 40), we expected to findhardly any N. apis-positive samples. Surprisingly, this was notthe case. With the exception of spring 2007, N. apis was alwaysmore prevalent than N. ceranae. The proportion of N. apis-positive colonies in the spring ranged from 15.7% (2009) to3.7% (2007), and that in the autumn ranged from 8.0% (2005)to 2.9% (2008) (Fig. 1). The proportion of colonies that testedpositive for N. ceranae ranged from 14.9% (2007) to 4.1%(2005) in the spring and from 4.2% (2005) to 1.3% (2006) inthe autumn (Fig. 1). In spring 2007, 14.9% N. ceranae-positivecolonies and only 3.1% N. apis-positive colonies were detectedin our cohort. A low prevalence of mixed infections with thesame seasonal pattern (more positive colonies in the spring

than in the autumn) was also detected throughout the studyperiod (Fig. 1).

Colony losses during the study period. In order to evaluatethe impact of Nosema infection on the mortality of honeybeecolonies, we differentiated between colony losses in the sum-mer season (weeks 15 to 35) and overwintering losses (weeks36 to week 14 of the following year), and we related these tothe detection of Nosema spp., N. apis, and N. ceranae in springand autumn samples (Tables 1 and 2). The winter losses weobserved in our cohort during the study period ranged from22.4% (2005/2006) to 4.8% (2008/2009), showing that ourstudy period covered winters with low to normal colony mor-tality rates as well as winters with rather high colony mortalityrates (Table 2). We observed a similar variation, although on amuch lower level (1.8% for 2007; 6.7% for 2008), for colonylosses during the season (Table 1). Our data did not reveal asignificant relation between the detection of Nosema infectionin the spring and colony losses in the following season (Table1, P values) or between detectable Nosema infections in theautumn and colony collapses during the following winter (Ta-ble 2, P values). In addition, no significant differences betweenthe mortality rates of uninfected colonies and those of coloniesinfected by N. apis, N. ceranae, or both (mixed infections)could be established (P, 0.05).

FIG. 1. Epidemiological survey of 22 apiaries in the northeastern part of Germany for Nosema prevalence between spring 2005 and spring 2009.The prevalences of samples positive for Nosema spp. (filled diamonds), Nosema apis (open squares), Nosema ceranae (open triangles), and bothspecies (mixed infections) (shaded circles) are shown at each time point. Diagnosis of Nosema spp. was performed microscopically. Nosemadifferentiation was performed via RFLP analysis of a PCR-amplified partial sequence of the 16S rRNA gene.

TABLE 1. Effects of Nosema infection in the spring on honeybee colony losses in the following season

Summerseason

Total no. ofcolonies

analyzed inspring

No. of colonies between wk 15 and 35

Pb%

Colonylosses

Surviving Collapsed

Total Nosemapositivea

Nosemanegative Total Nosema

positiveNosemanegative

2005 220 213 51 (36, 9, 6) 162 7 1 (1, 0, 0) 6 0.554 3.22006 238 223 65 (41, 10, 14) 158 15 2 (2, 0, 0) 13 0.187 6.32007 228 224 51 (17, 34, 10) 173 4 0 (0, 0, 0) 4 0.279 1.82008 209 195 67 (33, 13, 21) 128 14 7 (2, 5, 0) 7 0.237 6.72009 210 206 68 (33, 23, 12) 138 4 0 (0, 0, 0) 4 0.162 1.9

a The numbers of colonies positive for N. apis, for N. ceranae, and for both (mixed infections) are given in parentheses.b Determined by the 2 test.

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Tissue tropism of N. apis and N. ceranae. While N. apisinfections are reported to be restricted to the gut (16, 19, 21),N. ceranae could also be detected in hypopharyngeal glands,salivary glands, malpighian tubules, and fat bodies, indicating amore generalized infection (13), which could possibly be linkedto the reported increased lethality of N. ceranae (27, 40). Sinceour results did not reveal the previously reported, and there-fore expected, correlation between N. ceranae infection andcolony losses (28, 37), we asked whether a less virulent form ofN. ceranae, which did not cause such generalized infections,might be present in Germany. We therefore analyzed the tis-sue tropism of N. apis and N. ceranae in the infected beescollected in our study in detail but did not find any substantialdifferences from what had already been described (13). By theuse of PCR analysis and RFLP differentiation, N. apis wasdetected only in the guts of infected bees, supporting the tissuetropism reported for N. apis (16, 17, 19). In contrast, using thesame method, we detected N. ceranae in hypopharyngealglands, brains, guts, malpighian tubules, and fat bodies (Fig. 2).For the majority of the bees, infection of the gut could bedemonstrated. In addition to the detection of N. ceranae in thetissues, as reported by Chen and coworkers (13), we coulddetect N. ceranae in the brain as well. All N. ceranae-positivebees analyzed showed a rather generalized infection, with atleast two different tissues testing positive for N. ceranae. How-

ever, using the less sensitive method of Giemsa-stained smearpreparations, we could detect both N. apis and N. ceranae onlyin gut tissue (Fig. 3).

Germination of N. apis and N. ceranae. Previous studies haveshown that N. ceranae spores, but not N. apis spores, aresensitive to a temperature of �20°C (18). Since germination isthe first step in the infection process, we analyzed the germi-nation capacities of N. apis and N. ceranae spores exposed tolow temperatures versus room temperature. We found that thegermination of both Nosema species was affected after 4 daysat 4°C. Estimation of the numbers of germinated spores andextruded polar tubes revealed that about 80% of the N. apisspores were still capable of germination, while N. ceranae sporegermination was reduced to less than 10% (Fig. 4). In addition,the polar tubes extruded from the few N. ceranae spores stillcapable of germination were unusually short, most likely notrepresenting true germination (Fig. 4).

DISCUSSION

A recent study (33) is frequently cited to substantiate thenotion that N. ceranae is predominant in Europe, although thepublished data rather show that the patterns of incidence of N.apis and N. ceranae differ for different regions in Europe. N.ceranae seems to be more prevalent than N. apis in Denmark,

TABLE 2. Effects of Nosema infection in the autumn on honeybee colony losses in the following winter

Winter season

Total no. ofcolonies

analyzed inautumn

No. of colonies between wk 36 and 14

Pb%

Colonylosses

Surviving Collapsed

Total Nosemapositivea

Nosemanegative Total Nosema

positiveNosemanegative

2005–2006 237 184 21 (16, 4, 1) 163 53 9 (3, 6, 0) 44 0.283 22.42006–2007 226 212 19 (14, 3, 2) 193 14 1 (1, 0, 0) 13 0.816 6.22007–2008 219 181 11 (7, 3, 1) 170 38 4 (3, 1, 0) 34 0.324 17.42008–2009 210 200 11 (6, 5, 0) 189 10 0 (0, 0, 0) 10 0.446 4.8

a The numbers of colonies positive for N. apis, for N. ceranae, and for both (mixed infections) are given in parentheses.b Determined by the 2 test.

FIG. 2. Detection of Nosema spp. and differentiation via species-specific 16S rRNA gene RFLP in different tissues of infected bees sampledfrom colonies that tested positive for Nosema ceranae only. The tissues analyzed were hypopharyngeal glands (h. glands), brains, guts, malpighiantubules (m. tubules), and fat bodies.

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Greece, Italy, Serbia, and Spain, while N. apis could be de-tected more often than N. ceranae in Sweden and the UnitedKingdom. In the German samples analyzed (14 random sam-ples from spring 2006), the prevalence of both Nosema specieswas balanced. Although in that study a meaningful overviewwas given for Europe, the limited data for each country lowerthe representative character of the study. The present cohortstudy, performed with about 220 colonies over 5 years, revealsthat N. apis is still more prevalent in Germany, as in the UnitedKingdom and Sweden (33), than N. ceranae, although the prev-alence of N. ceranae might be on the rise. Two- and 3-foldincreases in the proportion of N. ceranae positive coloniescould be seen in spring 2008 and spring 2009, respectively, overthat at the beginning of the study (spring 2005 and spring2006), suggesting a current process of increasing N. ceranaeprevalence. We will continue our study with the same cohort inorder to see whether or not we are observing the replacementof N. apis by N. ceranae in Germany, a process that seems likelyaccording to data from other European countries (33) and theUnited States (12) and that will be most interesting to watch.

One of the hallmarks of the unusual form of nosemosiscaused by N. ceranae is reported to be a “loss of seasonality.”For N. apis it has been reported that in spring more colonieswill have detectable infection levels, i.e., more individuals willbe infected, and infected individuals will exhibit a higher sporeload. Typically, pathological symptoms of nosemosis (dysen-tery accompanied by defecation within the hive; crawling bees)will be evident in early spring, and colonies will collapse beforethe season starts (4, 5, 20). N. ceranae-infected European hon-eybees do not show this pathology. Instead, it is reported thatcolonies eventually collapse from the disease without showingany obvious symptoms of nosemosis (24). In addition, for N.ceranae infections of European honeybees in Spain, a changein seasonality has been reported, with an increase in Nosema-positive samples throughout the year, finally leading to a total

absence of seasonality in infection prevalence (37). Our datado not support such a situation for the prevalence of Nosema-positive colonies in Germany. The proportion of colonies withdetectable levels of Nosema infection was always higher inspring than in autumn. For colonies infected by N. ceranae,seasonality was not really pronounced in the beginning of thestudy, when the proportion of infected colonies was quite low(about 4%), but this changed in 2007, when 14.9% of thespring samples tested positive for N. ceranae. Since then, N.ceranae has followed the seasonality that is described for N.apis and could be verified in our study except for one season,spring 2007, when as few as 3.1% of the colonies were N. apispositive. Hence, the notion that the prevalence of colonies withdetectable levels of N. ceranae infection over the year does notshow the “normal” seasonality could not be substantiated inour study.

Studies performed with experimental infection of caged beeshave recently suggested that in addition to a different pathol-ogy, N. ceranae has a higher individual virulence than N. apis(27, 40), although this effect could be overcome by feeding thecaged bees ad libitum (38). Reports on the colony-level viru-lence of N. ceranae are contradictory. Several studies fromSpain suggest that N. ceranae is highly virulent at the colonylevel and that hence, infected colonies inevitably die from theinfection if left untreated. These studies also imply that N.ceranae is the cause for the unusual colony losses reportedfrom several regions in Europe and in the United States (28,29, 37). Other studies rather question a link between N. cera-nae infections and increased colony mortality or identify othercauses for unusual colony losses (11, 12, 15, 32, 44, 45). Theresults of our study also failed to reveal a relation between N.ceranae infection of colonies and colony mortality, even inseasons with unusually high colony loss rates. Likewise, mon-

FIG. 4. In vitro germination of N. apis and N. ceranae spores.Spores were isolated, air dried on glass slides, and kept at differenttemperatures for different periods. Representative results obtainedafter storage of the dried spores at �4°C for 4 days are shown incomparison with freshly isolated, dried, and germinated spores. Ex-truded polar tubes can be seen as curved lines in the images. Whitearrowheads point to extremely short, extruded polar tubes seen onlyfor N. ceranae after 4 days at �4°C.

FIG. 3. Detection of Nosema spp. via smear preparations in in-fected bees. Different organs of infected bees were analyzed for thepresence of spores and vegetative stages of Nosema spp. by usingGiemsa-stained smear preparations. A representative image from thegut of an N. apis-infected bee is shown. n, nucleus. Nuclei are stainedred, and spores are stained blue-white.

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itoring of the fate of individual N. ceranae-infected coloniesover several years (data not shown) did not show a mandatorylink between this infection and failure of the colony. Instead,infestation by Varroa destructor and infection with deformedwing virus (DWV) and acute bee paralysis virus (ABPV) couldbe identified as pathogenic processes related with high signif-icance to increased colony mortality in Germany (26).

Nevertheless, the facts remain that (i) in Spain and Italy N.ceranae nearly replaced N. apis over the past decade (33) and(ii) N. ceranae infections cause severe honeybee colony losses,at least in Spain (28, 30, 37). Possible reasons for this asser-tiveness and virulence of N. ceranae are the exceptional bioticpotential of N. ceranae at higher temperatures and the spores’tolerance to temperatures as high as 60°C, combined withresistance to desiccation (18, 36). On the other hand, N. cera-nae spores have been reported to be sensitive to freezing tem-peratures (18, 20). Since in our study, unexpectedly, neitherpredominance nor a noticeable colony-level virulence of N.ceranae could be verified, we searched for an explanation.Differences in virulence between genetically different isolates(25) were possible and will be addressed in further studies.However, in view of the studies of the influence of temperatureon N. ceranae spore viability and infectivity, discussed above,we rather suspected that climatic factors play a role in theoutcome of N. ceranae infections (20). The northeastern partof Germany is characterized by moderately warm but not hotsummers and rather cold and long winters (December to Feb-ruary) with a mean upper temperature threshold of about�4°C. We therefore analyzed and compared the germinationcapacities of N. apis and N. ceranae spores at different temper-atures. After as few as 4 days at �4°C, we found a markeddecrease in the level of germination of N. ceranae spores butnot in that of N. apis spores. Since germination is the first stepin the infection process of ingested environmental spores, adecrease in the germination of N. ceranae spores after such ashort time at moderately cold temperatures will reduce theinfectivity and virulence of N. ceranae at low temperatures.This cold sensitivity of N. ceranae spores, not seen with N. apisspores, could pose a clear disadvantage for N. ceranae in com-peting with N. apis in climatic regions with comparable or evencolder temperatures in the winter season. Environmentalspores (e.g., those defecated inside the hive and “waiting” to beingested by another bee) of N. ceranae will have a limitedchance of staying infective during any cold period in winter,while N. apis spores might stay unaffected even at �20°C (20).Within-colony transmission of N. ceranae will, therefore, behampered at lower temperatures, preventing N. ceranae frombuilding up in the colony over the winter as N. apis does. Thedisadvantage posed by the cold sensitivity of N. ceranae sporesmight even neutralize any advantageous effect of thermotoler-ance and resistance to desiccation, especially in regions wherethe summer temperature rarely rises above 33°C. This expla-nation for the limited assertiveness and virulence of N. ceranaein Germany so far is consistent with earlier results showing apredominance of N. apis in countries such as Sweden (withrather cold and long winters) but a prevalence of N. ceranae incountries such as Italy and Spain (with hot summers and mod-erate winters) (33). Further laboratory and field studies areneeded to substantiate the impact of climate on the colony-level virulence of the two Nosema species and on the success of

N. ceranae in replacing N. apis in the European honeybeepopulation.

In summary, N. ceranae is still not predominant in Germanyand has not yet replaced N. apis in the European honeybeepopulation in Germany. No link between N. ceranae infectionand an increased risk of colony mortality could be established.We presented evidence that the germination capacity of N.ceranae spores is affected by moderately low temperatures(�4°C), which may hamper the infectivity and transmission ofN. ceranae during the winter season. Hence, we suggest thatthe virulence and assertiveness of N. ceranae in the Europeanhoneybee population is influenced by climate and mightchange with climatic changes.

ACKNOWLEDGMENTS

This work was supported by grants from the Ministries for Agricul-ture from the Federal States of Brandenburg (MIL) and Sachsen-Anhalt (MLU), Germany, and through grants from Bayer AnimalHealth AG. S.G. and N.M. were supported by a grant from the Ger-man Ministry of Nutrition, Agriculture and Consumer Protection(BMELV).

REFERENCES

1. Aizen, M. A., L. A. Garibaldi, S. A. Cunningham, and A. M. Klein. 2008.Long-term global trends in crop yield and production reveal no currentpollination shortage but increasing pollinator dependency. Curr. Biol. 18:1572–1575.

2. Aizen, M. A., and L. D. Harder. 2009. The global stock of domesticatedhoney bees is growing slower than agricultural demand for pollination. Curr.Biol. 19:915–918.

3. Antunez, K., R. Martin-Hernandez, L. Prieto, A. Meana, P. Zunino, and M.Higes. 2009. Immune suppression in the honey bee (Apis mellifera) followinginfection by Nosema ceranae (Microsporidia). Environ. Microbiol. 11:2284–2290.

4. Bailey, L. 1955. The epidemiology and control of Nosema disease of thehoney bee. Ann. Appl. Biol. 43:379–389.

5. Bailey, L. 1967. Nosema apis and dysentery of the honey bee. J. Apicult. Res.6:121–125.

6. Ball, B. V. 1983. The association of Varroa jacobsoni with virus diseases ofhoney bees. Exp. Appl. Acarol. 19:607–613.

7. Ball, B. V. 1996. Honey bee viruses: a cause for concern? Bee World 77:117–119.

8. Becnel, J. J., and T. G. Andreadis. 1999. Microsporidia in insects, p. 447–501.In M. Wittner and L. M. Weiss (ed.), The microsporidia and microsporidi-osis. ASM Press, Washington, DC.

9. Bigliardi, E., and L. Sacchi. 2001. Cell biology and invasion of the micro-sporidia. Microbes Infect. 3:373–379.

10. Canning, E. U., and J. Lom. 1986. The microsporidia of vertebrates, p. 1–16.Academic Press, New York, NY.

11. Chauzat, M. P., M. Higes, R. Martín-Hernandez, A. Meana, N. Cougoule,and J. P. Faucon. 2007. Presence of Nosema ceranae in French honey beecolonies. J. Apicult. Res. 46:127–128.

12. Chen, Y., J. D. Evans, I. B. Smith, and J. S. Pettis. 2008. Nosema ceranae isa long-present and wide-spread microsporidian infection of the Europeanhoney bee (Apis mellifera) in the United States. J. Invertebr. Pathol. 97:186–188.

13. Chen, Y. P., J. D. Evans, C. Murphy, R. Gutell, M. Zuker, D. Gundensen-Rindal, and J. S. Pettis. 2009. Morphological, molecular, and phylogeneticcharacterization of Nosema ceranae, a microsporidian parasite isolated fromthe European honey bee, Apis mellifera. J. Eukaryot. Microbiol. 56:142–147.

14. Corradi, N., and P. J. Keeling. 2009. Microsporidia: a journey through radicaltaxonomical revisions. Fungal Biol. Rev. 23:1–8.

15. Cox-Foster, D. L., S. Conlan, E. C. Holmes, G. Palacios, J. D. Evans, N. A.Moran, P.-L. Quan, S. Briese, M. Hornig, D. M. Geiser, V. Martinson, D.vanEngelsdorp, A. L. Kalkseitn, L. Drysdale, J. Hui, J. Zhai, L. Cui, S.Hutchison, J. F. Simons, M. Egholm, J. S. Pettis, and W. I. Lipkin. 2007. Ametagenomic survey of microbes in honey bee colony collapse disorder.Science 318:283–287.

16. de Graaf, D., and F. J. Jacobs. 1991. Tissue specificity of Nosema apis.J. Invertebr. Pathol. 58:277–278.

17. de Graaf, D., H. Raes, G. Sabbe, P. H. Rycke, and F. J. Jacobs. 1994. Earlydevelopment of Nosema apis (Microspora: Nosematidae) in the midgutepithelium of the honeybee (Apis mellifera). J. Invertebr. Pathol. 63:74–81.

18. Fenoy, S., C. Rueda, M. Higes, R. Martín-Hernandez, and C. del Aguila.2009. High-level resistance of Nosema ceranae, a parasite of the honeybee, totemperature and desiccation. Appl. Environ. Microbiol. 75:6886–6889.

VOL. 76, 2010 NOSEMA INFECTION IN HONEYBEES 3037

on March 7, 2021 by guest

http://aem.asm

.org/D

ownloaded from

19. Fries, I. 1988. Infectivity and multiplication of Nosema apis Z. in the ven-triculus of the honey bee. Apidologie 19:319–328.

20. Fries, I. 2010. Nosema ceranae in European honey bees (Apis mellifera).J. Invertebr. Pathol. 103:573–579.

21. Fries, I. 1997. Protozoa, p. 59–76. In R. A. Morse and K. Flottum (ed.),Honey bee pests, predators, and diseases. A. I. Root Company, Medina, OH.

22. Fries, I., G. Ekbohm, and E. Villumstad. 1984. Nosema apis, sampling tech-niques and honey yield. J. Apicult. Res. 23:102–105.

23. Fries, I., F. Feng, A. daSilva, S. B. Slemenda, and N. J. Pieniazek. 1996.Nosema ceranae n. sp. (Microspora, Nosematidae), morphological and mo-lecular characterization of a microsporidian parasite of the Asian honey beeApis cerana (Hymenoptera, Apidae). Eur. J. Protistol. 32:356–365.

24. Fries, I., R. Martin, A. Meana, P. Garcia-Palencia, and M. Higes. 2006.Natural infections of Nosema ceranae in European honey bees. J. Apicult.Res. 45:230–233.

25. Gatehouse, H. S., and L. A. Malone. 1999. Genetic variability among Nosemaapis isolates. J. Apicult. Res. 38:79–85.

25a.Gatehouse, H. S., and L. A. Malone. 1998. The ribosomal RNA gene regionof Nosema apis (Microspora): DNA sequence for small and large subunitrRNA genes and evidence of a large tandem repeat unit size. J. Invertebr.Pathol. 71:97–105.

26. Genersch, E., W. von der Ohe, H. Kaatz, A. Schroeder, C. Otten, R. Buchler,S. Berg, W. Ritter, W. Muhlen, S. Gisder, M. Meixner, G. Liebig, and P.Rosenkranz. The German bee monitoring project: a long term study tounderstand periodically high winter losses of honey bee colonies. Apidologie,in press. doi:10.1051/apido/2010014.

27. Higes, M., P. Garcia-Palencia, R. Martín-Hernandez, and A. Meana. 2007.Experimental infection of Apis mellifera honeybees with Nosema ceranae(Microsporidia). J. Invertebr. Pathol. 94:211–217.

28. Higes, M., R. Martín-Hernandez, C. Botías, E. Garrido Bailon, A. V. Gonza-lez-Porto, L. Barrios, M. J. del Nozal, J. L. Bernal, J. J. Jimenez, P. GarcíaPalencia, and A. Meana. 2008. How natural infection by Nosema ceranaecauses honeybee colony collapse. Environ. Microbiol. 10:2659–2669.

29. Higes, M., R. Martín-Hernandez, C. Botias, and A. Meana. 2009. The pres-ence of Nosema ceranae (Microsporidia) in African honey bees (Apis mel-lifera intermissa). J. Apicult. Res. 48:217–219.

30. Higes, M., R. Martin, and A. Meana. 2006. Nosema ceranae, a new micro-sporidian parasite in honeybees in Europe. J. Invertebr. Pathol. 92:93–95.

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

32. Invernizzi, C., C. Abud, I. H. Tomasco, J. Harriet, G. Ramallo, J. Campa, H.Katz, G. Gardiol, and Y. Mendoza. 2009. Presence of Nosema ceranae inhoneybees (Apis mellifera) in Uruguay. J. Invertebr. Pathol. 101:150–153.

33. Klee, J., A. M. Besana, E. Genersch, S. Gisder, A. Nanetti, D. Q. Tam, T. X.Chinh, F. Puerta, J. M. Ruz, P. Kryger, D. Message, F. Hatjina, S. Korpela,I. Fries, and R. J. Paxton. 2007. Widespread dispersal of the microsporidianNosema ceranae, an emergent pathogen of the western honey bee, Apismellifera. J. Invertebr. Pathol. 96:1–10.

34. Lee, S. C., N. Corradi, E. J. Byrnes III, S. Torres-Martinez, F. S. Dietrich,

P. J. Keeling, and J. Heitman. 2008. Microsporidia evolved from ancestralsexual fungi. Curr. Biol. 18:1675–1679.

35. Martin, S. J. 2001. The role of Varroa and viral pathogens in the collapse ofhoneybee colonies: a modelling approach. J. Appl. Ecol. 38:1082–1093.

36. Martín-Hernandez, R., A. Meana, P. Garcia-Palencia, P. Marin, C. Botias,E. Garrido-Bailon, L. Barrios, and M. Higes. 2009. Effect of temperature onthe biotic potential of honeybee microsporidia. Appl. Environ. Microbiol.75:2554–2557.

37. Martín-Hernandez, R., A. Meana, L. Prieto, A. M. Salvador, E. Garrido-Bailon, and M. Higes. 2007. Outcome of colonization of Apis mellifera byNosema ceranae. Appl. Environ. Microbiol. 73:6331–6338.

38. Mayack, C., and D. Naug. 2009. Energetic stress in the honeybee Apismellifera from Nosema ceranae infection. J. Invertebr. Pathol. 100:185–188.

38a.Office Internationale des Epizooties—World Organization for AnimalHealth. 2008. Manual of diagnostic tests and vaccines for terrestrial animals,p. 410–414. OIE, Paris, France.

39. Olsen, P. E., W. A. Rice, and T. P. Liu. 1986. In vitro germination of Nosemaapis spores under conditions favorable for the generation and maintenanceof sporoplasms. J. Invertebr. Pathol. 47:65–73.

40. Paxton, R. J., J. Klee, S. Korpela, and I. Fries. 2007. Nosema ceranae hasinfected Apis mellifera in Europe since at least 1998 and may be morevirulent than Nosema apis. Apidologie 38:558–565.

41. Ritter, W. 1996. Diagnostik und Bekampfung der Bienenkrankheiten. Fi-scher, Stuttgart, Germany.

42. Tapaszti, Z., P. Forgach, C. Kovago, L. Bekesi, T. Bakonyi, and M. Rusvai.2009. First detection and dominance of Nosema ceranae in Hungarian hon-eybee colonies. Acta Vet. Hung. 57:383–388.

43. vanEngelsdorp, D. 2008. A survey of honey bee colony losses in the U.S., fall2007 to spring 2008. PLoS One 3:e4071.

44. vanEngelsdorp, D., J. D. Evans, L. Donovall, C. Mullin, M. Frazier, J.Frazier, D. R. Tarpy, J. Hayes, and J. S. Pettis. 2009. “Entombed pollen”: anew condition in honey bee colonies associated with increased risk of colonymortality. J. Invertebr. Pathol. 101:147–149.

45. vanEngelsdorp, D., J. D. Evans, C. Saegerman, C. Mullin, E. Haubruge,B. K. Nguyen, M. Frazier, J. Frazier, D. Cox-Foster, Y. Chen, R. Underwood,D. R. Tarpy, and J. S. Pettis. 2009. Colony collapse disorder: a descriptivestudy. PLoS One 4:e6481.

46. vanEngelsdorp, D., and M. D. Meixner. 2010. A historical review of managedhoney bee populations in Europe and the United States and the factors thatmay affect them. J. Invertebr. Pathol. 103(Suppl. 1):S80–S95.

47. Weber, R., R. T. Bryan, D. A. Schwartz, and R. L. Owen. 1994. Humanmicrosporidial infections. Clin. Microbiol. Rev. 7:426–461.

48. Williams, G. R., A. B. A. Shafer, R. E. L. Rogers, D. Shutler, and D. T.Stewart. 2008. First detection of Nosema ceranae, a microsporidian parasiteof European honey bees (Apis mellifera), in Canada and central USA. J. In-vertebr. Pathol. 97:189–192.

49. Zander, E. 1909. Tierische Parasiten als Krankheitserreger bei der Biene.Munch. Bienenzeitung 31:196–204.

3038 GISDER ET AL. APPL. ENVIRON. MICROBIOL.

on March 7, 2021 by guest

http://aem.asm

.org/D

ownloaded from