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Applied and Environmental Microbiology, January 2006, p. 606-611, Vol. 72, No. 1
0099-2240/06/$08.00+0     doi:10.1128/AEM.72.1.606-611.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Prevalence and Transmission of Honeybee Viruses

Y. P. Chen,^* J. S. Pettis, A. Collins, and M. F. Feldlaufer

Bee Research Laboratory, USDA-ARS, Beltsville, Maryland 20705

Received 12 August 2005/ Accepted 24 October 2005

Top Abstract Introduction Materials and Methods Results Discussion References

 
Transmission mechanisms of six honeybee viruses, including acute bee paralysis virus (ABPV), black queen cell virus (BQCV), chronic bee paralysis virus (CBPV), deformed wing virus (DWV), Kashmir bee virus (KBV), and sacbrood bee virus (SBV), in honey bee colonies were investigated by reverse transcription-PCR (RT-PCR) methods. The virus status of individual queens was evaluated by examining the presence of viruses in the queens' feces and tissues, including hemolymph, gut, ovaries, spermatheca, head, and eviscerated body. Except for head tissue, all five tissues as well as queen feces were found to be positive for virus infections. When queens in bee colonies were identified as positive for BQCV, DWV, CBPV, KBV, and SBV, the same viruses were detected in their offspring, including eggs, larvae, and adult workers. On the other hand, when queens were found positive for only two viruses, BQCV and DWV, only these two viruses were detected in their offspring. The presence of viruses in the tissue of ovaries and the detection of the same viruses in queens' eggs and young larvae suggest vertical transmission of viruses from queens to offspring. To our knowledge, this is the first evidence of vertical transmission of viruses in honeybee colonies.

Top Abstract Introduction Materials and Methods Results Discussion References

 
The honeybee, Apis mellifera L., plays a vital role in agriculture by assisting in the pollination of a wide variety of crops, with an annual added market value exceeding 15 billion dollars (13). However, the health and vigor of honeybee colonies are threatened by numerous parasites and pathogens, including viruses, bacteria, protozoa, and mites. Among pathogens attacking honeybees, viruses are probably the least understood because of the lack of information about the dynamics underlying viral disease outbreaks.

A most crucial stage in the dynamics of virus infections is the mode of virus transmission. In general, transmission of viruses can occur through two pathways: horizontal and vertical transmission. In horizontal transmission, viruses are transmitted among individuals of the same generation, while vertical transmission occurs from adults to their offspring. Transmission can occur through multiple routes in social organisms. Over the past several years, horizontal transmission of honeybee viruses has been documented in bee colonies. Bowen-Walker et al. (3) experimentally demonstrated that the parasitic mite Varroa destructor obtained deformed wing virus (DWV) from infected bees and acted as a vector to transmit the virus to uninfected bees, which developed morphological deformities or died after mites fed on them for certain periods of time. Our recent study with Kashmir bee virus (KBV) established the role of V. destructor in virus transmission and provided evidence of mite-to-brood transmission and mite-to-mite acquisition of viruses in bee colonies (5). Although these results demonstrated horizontal transmission of viruses, virus transmission in honeybees is still not completely understood. For instance, in our subsequent studies with DWV (4, 6), we detected virus in honeybee eggs and young larvae, life stages not parasitized by Varroa mites. These results suggested an alternative route of transmission and led us to investigate the issue of whether or not viruses can be transmitted vertically from the queen bee to her offspring.

For the present study, the transmission of viruses from the queen to the next generation was investigated in honeybee colonies. Using reverse transcription-PCR (RT-PCR), we examined various tissues of queen bees for presence of viruses. Furthermore, virus titers in different queen tissues were compared by semiquantitative RT-PCR to identify potential tissue reservoirs of virus replication in the body of the queen. Additionally, we examined the eggs, larvae, and adult worker bees associated with each queen for the presence of viruses in an attempt to prove vertical transmission.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Sample collection.
Ten honeybee colonies maintained in the USDA-ARS Bee Research Laboratory apiaries, Beltsville, MD, were chosen randomly for vertical virus transmission studies. Queens were collected from each colony, isolated in a small cage individually with several attending workers, and supplied with honey. A representative brood frame containing eggs and brood of various ages was selected from each colony. Larvae were collected by uncapping brood cells and transferring them to individual microcentrifuge tubes with fine forceps. Adult bees were collected by shaking the frame and gently scraping worker bees into 50-ml conical tubes. About 50 eggs from each colony were collected with a tapered brush. Eggs were washed in 5% Clorox solution for 5 min to minimize possible surface contamination and then rinsed in DNase- and RNase-free water (Invitrogen, Carlsbad, CA) three times. Eggs were subjected to RNA extraction immediately. All samples were stored at –80°C for further analysis.

Queen feces collection.
Queens were removed from each colony and initially isolated individually in small queen cages, and then each queen was transferred to a 100- by 15-mm petri dish to allow it to defecate. The clear fecal material was collected with a micropipette; about 20 to 25 µl of feces could be collected from each queen after 30 min of isolation. Collected feces were stored in –80°C freezer until used.

Tissue dissection.
After feces collection, the queens were sacrificed for tissue dissection. Each live queen was fixed on the wax top of a dissecting dish with insect pins. Hemolymph was collected with a micropipette tip by making a small hole on the roof of the queen's thorax with a needle to make it bleed. About 15 to 20 µl of hemolymph was collected from each queen. Following hemolymph collection, the abdomen of the queen was opened with scissors. Tissues of spermatheca, gut, and ovaries were carefully separated and pulled out with forceps under a dissecting microscope. The head was cut off from the eviscerated body, and both the head and the remaining body of the queen were saved in different tubes. To prevent possible contamination with hemolymph, all tissues were rinsed with 1x phosphate-buffered saline once and nuclease-free water twice. All of the tissue samples were subjected to subsequent RNA extraction.

RNA extraction.
For each colony, total RNA was extracted from 10 samples of larvae, 10 samples of adult bees, 1 sample of 50 eggs, 1 sample of queen feces, and 6 samples of queen tissues, including hemolymph, gut, spermatheca, ovaries, head, and eviscerated body, for a total of 28 samples per colony. Individual samples were homogenized in TRIzol reagent (RNA extraction kit; Invitrogen, Carlsbad, CA), and total RNA was extracted following the manufacturer's standard protocol. RNA samples were dissolved in diethyl pyrocarbonate-treated water in the presence of RNase inhibitor (Invitrogen, Carlsbad, CA). The concentration of total RNA was determined by measuring the absorption at 260 nm, and the purity of RNA was estimated by the absorbance ratio of 260 nm/280 nm using a spectrophotometer with a 50-µl ultramicrovolume cell holder (Ultrospec 3300 pro; Amersham Biosciences). RNA samples were stored at –80°C prior to molecular detection for viruses.

RT-PCR.
All RNA samples were tested for the presence of six bee viruses, namely, acute bee paralysis virus (ABPV), black queen cell virus (BQCV), chronic bee paralysis virus (CBPV), DWV, KBV, and sacbrood bee virus (SBV) by RT-PCR. The primers used in the study were synthesized by Invitrogen and are shown in Table 1. The GenBank accession numbers for each virus and ß-actin are also included in Table 1. The Access RT-PCR system (Promega, Madison, WI) was used for RT-PCR according to the manufacturer's instructions. The reaction was performed in a total volume of 25 µl with a final concentration of 1x avian myeloblastosis virus/Tfl reaction buffer, 0.2 mM of deoxynucleoside triphosphate (dNTP), 1 µM of sense primer, 1 µM of antisense primer, 2 mM of MgSO₄, 0.1 unit of avian myeloblastosis virus reverse transcriptase, 0.1 unit of Tfl DNA polymerase, and 500 ng of total RNA. Amplification was undertaken using the PTC-100 DNA Engine (MJ Research, Waltham, MA) with the following thermal cycling profiles: one cycle at 48°C for 45 min for reverse transcription; 1 cycle of 95°C for 2 min; 40 cycles at 95°C for 30 s, 55°C for 1 min, and 68°C for 2 min; and 1 cycle of 68°C for 7 min. Negative (H₂O) and positive (previously identified positive sample) controls were included in each run of the RT-PCR. PCR products were electrophoresed in 1% agarose gel containing 0.5 µg/ml ethidium bromide and visualized by UV transillumination. A 100-bp DNA ladder (Invitrogen) was included as a standard on each gel.

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TABLE 1. Primers used in this study

Densitometry of the RT-PCR products.
PCR products of the queen tissues were analyzed by densitometry to provide a semiquantification of virus titers in different tissues. To normalize the results, the quality of all samples was checked by RT-PCR amplification of an internal control, ß-actin. The intensities of the PCR bands on the gel were quantified by densitometry (LabWorks 4.0, Image Acquisition and Analysis Software; UVP). The densitometry result of each sample was divided by the densitometry result of the ß-actin. The results are expressed as the ratio of the band intensity of the target PCR product to that of ß-actin.

Sequencing.
The specificity of the RT-PCR assay was confirmed by sequence analysis. RT-PCR bands specific for ABPV, BQCV, CBPV, DWV, KBV, and SBV were excised from the low-melting-temperature agarose gel (Invitrogen, Carlsbad, CA) and purified using Wizard PCR Prep DNA purification system (Promega, Madison, WI). Purified RT-PCR fragments were individually ligated into a TOPO cloning vector (Invitrogen, Carlsbad, CA). Recombinant plasmid DNAs were purified using a Plasmid Mini Prep kit (Bio-Rad, Hercules, CA). The nucleotide sequences of the cloned RT-PCR fragments were determined from both forward and reverse directions. The sequence data of each virus fragment were analyzed using the BLAST server at the National Center for Biotechnology Information, NIH.

Statistical analysis.
Statistical analysis was performed using Statistix7 statistical software (Analytic Software). The Fisher's least significant difference comparison of means test was used to analyze for significant differences of virus titers among different tissues of the queens. The results are expressed as the mean ± standard deviation. Differences were considered statistically significant if P was <0.05.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Detection of multiple viruses from queen feces.
RT-PCR was performed to screen for presence of bee viruses from samples of queen feces. Of six viruses screened, BQCV and DWV were detected in the samples of queen feces. One hundred percent of feces samples (10/10) collected from individual queens tested positive for the presence of BQCV, and 90% of feces samples (9/10) tested positive for the presence of DWV (Table 2).

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TABLE 2. Presence of bee viruses in feces and tissues of queens by RT-PCR assay

Distribution of viruses in the body of queens.
The results of RT-PCR analysis on tissues of hemolymph, gut, ovaries, spermatheca, head, and eviscerated body from 10 queens for the presence of six viruses are shown in Table 1. Among the six viruses screened, ABPV was not found in any materials. BQCV, CBPV, DWV, KBV, and SBV were detected in one or more queen tissues. Except for tissues of the head, which were negative for all six viruses, the other five tissue samples were found to be virus positive. The presence of DWV was found in 100% of hemolymph samples, 80% of gut samples, 100% of ovary samples, 80% of spermatheca samples, and 100% of eviscerated body samples. The presence of BQCV was found in 100% of gut samples and 70% of ovary samples. The presence of SBV was found in 40% of ovary samples, 20% of hemolymph samples, and 60% of eviscerated body samples. The presence of CBPV was found in 30% of hemolymph and 40% of eviscerated body samples. The presence of KBV was found in 20% of the eviscerated body samples.

Semiquantification of virus titers in tissues of queens.
RT-PCR using the ß-actin primers resulted in the amplification of the ß-actin fragment from all tissue samples tested, indicating that approximately equal amounts of initial RNAs were used for RT-PCR analysis. Semiquantification of DWV levels in different tissues of queens showed that virus titer in the guts was significantly higher than in any other tissues tested. There was no significant difference in DWV titer between tissues of spermatheca and eviscerated body samples. The lowest relative DWV titers were observed in the tissues of ovaries. The ratios of the band intensity of DWV-specific PCR product relative to that of ß-actin were 2.63 ± 0.68, 0.08 ± 0.25, 2.0 ± 0.45, and 2.16 ± 0.49 for gut, ovaries, spermatheca, and eviscerated body, respectively (Fig. 1A).

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FIG. 1. Semiquantification of bee viruses in tissues of queens. (A to E, top panels) RT-PCR results on tissues of queens for the presence of bee viruses. Six tissues including hemolymph, gut, ovaries, spermatheca, head, and eviscerated body (E. body) were examined for the presence of viruses. Primer pairs specific for five viruses including BQCV, CBPV, DWV, KBV, SBV, and an internal control, ß-actin, were used separately to amplify RT-PCR products of 700 bp, 455 bp, 702 bp, 415 bp, 824 bp, and 357 bp, respectively. Negative (H2O [N]) and positive (P) controls (previously identified positive sample) were included in each run of the RT-PCR. (A to E, bottom panels) Densitometric analyses. The y axis depicts the ratio of band intensity of the virus-specific RT-PCR products to that of ß-actin RT-PCR products. Results are expressed as mean ± standard deviation (n = 3). Different letters indicate a significant difference among tissues (P = 0.05). (A) Semiquantification of DWV in tissues of queens. (B) Semiquantification of BQCV in tissues of queens. (C) Semiquantification of SBV in tissues of queens. (D) Semiquantification of CBPV bee viruses in tissues of queens. (E) Semiquantification of KBV bee viruses in tissues of queens.

Semiquantification of BQCV levels showed that virus titer in the guts was significantly higher than that in the ovaries and that the ratios of the band intensity of BQCV-specific PCR product relative to that of ß-actin were 1.45 ± 0.98 and 0.25 ± 0.35 for gut and ovaries, respectively (Fig. 1B).

Semiquantification of SBV levels showed that virus titer in the ovaries was significantly lower than that SBV titer in the hemolymph and eviscerated body and that SBV titer in the eviscerated body was significantly lower than in hemolymph. The ratios of the band intensity of SBV-specific PCR product relative to that of ß-actin were 1.03 ± 0.60, 0.33 ± 0.30, and 0.52 ± 0.69 for hemolymph, ovaries, and eviscerated body, respectively (Fig. 1C). Infection of CBPV was detected in tissues of the hemolymph and the eviscerated body. The ratios of the band intensity of CBPV PCR product relative to that of ß-actin were 0.20 ± 0.38 and 0.18 ± 0.93 for hemolymph and the eviscerated body, respectively. The band intensity of CBPV PCR product varied significantly among different samples of eviscerated body, which caused a high value for the standard deviation in the ratio of the band intensity. There was no significant difference in the titer of CBPV between tissues of hemolymph and eviscerated body (Fig. 1D). The presence of KBV was found only in eviscerated body samples, and the ratio of the band intensity of KBV specific PCR product to that of ß-actin was about 0.42 ± 0.69 (Fig. 1E).

Evidence of the vertical transmission of viruses from queens to their offspring.
Individual queens that were detected with virus or viruses in their feces or in any one of the dissected tissues were considered to be virus positive, although all of the queens in this study showed no pathological signs of virus infections. Among 10 queens examined, 100% were RT-PCR positive for DWV and BQCV, 60% were SBV positive, 40% were CBPV positive, and 20% were KBV positive. Based on the virus status of the queens, 10 experimental colonies were divided into two groups. In group A, queens were identified to be positive for three to five viruses, while in group B, queens were infected with only two viruses, BQCV and DWV.

The possibility of vertical transmission in the colonies was investigated by examining the virus status of the queens' offspring, including eggs, larvae, and adults in both groups. As shown in Table 3, all of the queens (n = 6) in group A were positive for BQCV, DWV, and SBV; the presence of BQCV, DWV, and SBV was found in 100% of egg samples (6/6), in 27%, 92%, and 25% of larvae, and in 3%, 37%, and 10% of adult bees, respectively. When 67% of the queens (4/6) in group A were positive for CBPV, 50% of egg samples, 17% of larvae, and 17% of adult bees were infected with CBPV. When 33% of the queens in group A were positive for KBV, 33% of egg samples were infected. No KBV was detected in larvae and adult bees. In group B, all of the queens were positive for BQCV and DWV, 100% of egg samples (4/4) were identified to be infected with the same two viruses, 22% of larvae and 5% adult bees were infected with BQCV, and 65% of larvae and13% of adult bees were infected with DWV.

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TABLE 3. Vertical transmission of viruses from virus-infected queens to their offspring

Sequence confirmation of RT-PCR specificity.
Sequence analysis showed that sequences of selected RT-PCR fragments representing individual viruses, including BQCV, CBPV, DWV, KBV, and SBV, and ß-actin matched the corresponding virus sequences published in GenBank, indicating that amplification bands in this study were virus specific.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Vertical transmission is thought to play an important role in the long-term maintenance of viruses in the wild. The occurrence of vertical transmission of viruses has long been known to occur in mammals, vertebrates, arthropods, and plants (12). Vertical transmission of viruses in honeybees has been considered to be possible and has been discussed in a honeybee disease model (9), but no data based on experimental observations were presented to support this mechanism. The present study, using the sensitive RT-PCR method, demonstrated the vertical transmission of multiple viruses from mother queens to their offspring by two findings: first, the presence of viruses in queen excretion and queen tissues, particularly in the tissue of ovaries; and second, detection of the same viruses in queens' eggs and young larvae that are not normally associated with V. destructor, which is an important vector of bee viruses.

In this study, we didn't make an attempt to inoculate virus-negative queens with purified viruses and then to estimate the filial infection rates or to recover the viruses from the queens' progeny due to the following three limitations. (i) Honey bees are often attacked by multiple viral infections (7); therefore, it is difficult to purify virus particles that contain only a single virus. (ii) Most of the queens among honeybees are virus carriers (4). (iii) It is difficult to identify the virus status of queens without killing them. Although this study is not a definite experimental proof for transovarian transmission, the presence of viruses in tissues of ovaries and surface-sterilized eggs is perhaps the best evidence of transovarian transmission, in which viruses infect ovarian tissues of the queen and disseminate in developing eggs before oviposition. Semiquantitative RT-PCR analysis showed that three viruses, BQCV, DWV, and SBV, were detected in tissues of ovaries and that virus titers in ovaries were significantly lower than those in tissues of the gut. However, two other viruses, CBPV and KBV, which were found in tissues of hemolymph and eviscerated body as well as queens' offspring, were not detected in the ovaries. Previous studies with a mosquito (Culex pipiens) that transmitted West Nile virus indicated that the amount of viruses in mosquito ovaries tended to be very low (8). Low titers of viruses in ovaries can keep infection in a latent stage so that viruses would not be propagated to the level that will have deleterious effects on embryos. However, this benign state of virus infection could be transformed into a replicative and infective state when the host is stressed (10). No positive signal of the presence of CBPV and KBV in ovaries may imply that the titer of viruses was below the level of detection.

Our results indicate that viruses found in queens were also present in their eggs. One group of queens were found to be positive for three to five viruses, and the same viruses were detected in their eggs, larvae, and adult worker bees. Meanwhile, another group of queens were positive for only two viruses (BQCV and DWV), and the same two viruses were detected in their eggs, larvae, and adult worker bees. The viruses (CBPV, KBV, and SBV) that didn't present in the queens were also not detected in the queens' eggs. These data clearly demonstrated that viruses were transmitted from queens to their progeny. Detection of viruses in surface-sterilized eggs excludes the possibility of transovum transmission and indicates that viruses were transmitted from the infected queens to eggs. Although detection of viruses by RT-PCR has been substantiated as highly sensitive and specific, it is very difficult for us to extract enough RNA from a single bee egg for RT-PCR analysis. As a result, eggs from the same colony were combined into one group and the transmission rate of viruses from queen to eggs could not be estimated in this study.

Our previous study, which detected DWV in adult drones, indicated that drones are more resistant to DWV infection than other castes in the colony (4). Detection of viruses in spermathecae, together with the fact that DWV was detected in adult drones, suggests that another vertical transmission pathway, venereal transmission, may exist within the bee colony. Venereal transmission is a type of horizontal transmission in which viruses are transmitted from infected males to females during mating. Drones in a bee colony can be infected vertically from their parental female and can also transmit viruses venereally to queens during fertilization. Further studies are necessary to verify this possibility.

The detection of viruses in feces of queens suggests a role for feeding in virus transmission. Evidence of detection of viruses in honeybee feces has been reported previously (1, 11). This study is consistent with previous findings and showed that two viruses, BQCV and DWV, were found in queen feces. The detection of viruses in feces of queens suggests a role for feeding in virus transmission. Positive RT-PCR signals for viruses together with the fact that BQCV and DWV were detected in tissues of the gut suggest that viruses could be ingested by queens from contaminated foods and passed into the digestive tract. The high titers of BQCV and DWV detected in gut among six examined tissues suggest that tissue of the gut may be a major reservoir for replication of BQCV and DWV. Detection of BQCV and DWV in other tissues such as hemolymph, ovaries, and spermatheca with relatively lower virus titers suggests the possibility that infected viruses in the gut might penetrate the wall of the gut and move into the insect hemocoel to spread infections to other tissues. However, further studies are necessary to identity the precise route(s) of virus infection in the body of the queen.

Our work has provided substantial evidence for the vertical transmission of viruses in honeybees, but a number of factors that may play important roles in the efficiency of virus transmission are far from being understood. For example, the host immune responses and virus pathological features that facilitate the vertical transmission of individual viruses are not known. The roles of vertical transmission of viruses in bee disease epidemiology need to be determined. This will be especially relevant for honeybees, where viruses normally persist as latent infections and group living can possibly drive high levels of horizontal transmission or amplification of existing infections. Further studies of host-virus interactions might give some insight into these issues.

 
The authors wish to express their sincere gratitude to Michele Hamilton, Virginia Williams, Nathan Rice, Andrew Ulsamer, and Dave Vincent for providing excellent technical assistance.

This work was supported in part by the 2004 California State Beekeepers’ Association (CSBA) research fund.

Mention of trade names or commercial products in this article is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture.

 
* Corresponding author. Mailing address: Bee Research Laboratory, USDA-ARS, Bldg. 476, BARC East, Beltsville, MD 20705. Phone: (301) 504-8749. Fax: (301) 504-8736. E-mail: chenj{at}ba.ars.usda.gov

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Bailey, L., and A. J. Gibbs. 1964. Acute infection of bees with paralysis virus. J. Insect Pathol. 6:395-407.
2
2. Benjeddou, M., N. Leat, M. Allsopp, and S. Davison. 2001. Detection of acute bee paralysis virus and black queen cell virus from honeybees by reverse transcriptase PCR. Appl. Environ. Microbiol. 67:2384-2387.[Abstract/Free Full Text]
3
3. Bowen-Walker, P. L., S. J. Martin, and A. Gunn. 1999. The transmission of deformed wing virus between honey bees (Apis mellifera L.) by the ectoparasitic mite Varroa jacobsoni Oud. J. Invertebr. Pathol. 73:101-106.[CrossRef][Medline]
4
4. Chen, Y. P., J. A. Higgins, and M. F. Feldlaufer. 2005. Quantitative analysis by real-time reverse transcription-PCR of deformed wing virus infection in the honeybee (Apis mellifera L.). Appl. Environ. Microbiol. 71:436-441.[Abstract/Free Full Text]
5
5. Chen, Y. P., J. S. Pettis, J. D. Evans, M. Kramer, and M. F. Feldlaufer. 2004. Molecular evidence for transmission of Kashmir bee virus in honey bee colonies by ectoparasitic mite, Varroa destructor. Apidologie 35:441-448.[CrossRef]
6
6. Chen, Y. P., B. Smith, A. M. Collins, J. S. Pettis, and M. F. Feldlaufer. 2004. Detection of deformed wing virus infection in honey bees, Apis mellifera L., in the United States. Am. Bee J. 144:557-559.
7
7. Chen, Y. P., Y. Zhao, J. Hammond, H.-T. Hsu, J. D. Evans, and M. F. Feldlaufer. 2004. Multiple virus infections in the honey bee and genome divergence of honey bee viruses. J. Invertebr. Pathol. 87:84-93.[Medline]
8
8. Dohm, D. J., M. R. Sardelis, and M. J. Turell. 2002. Experimental vertical transmission of West Nile virus by Culex pipiens (Diptera: Culicidae). J. Med. Entomol. 39:640-644.[Medline]
9
9. Fries, I., and S. Camazine. 2001. Implications of horizontal and vertical pathogen transmission for honey bee epidemiology. Apidologie 32:1-16.
10
10. Fusa, J. R., and A. R. Richter. 1992. Virulence and multigeneration passage of a nuclear polyhedrosis virus selected for an increased rate of vertical transmission. Biol. Control 2:171-175.
11
11. Hung, A. C. F. 2000. PCR detection of Kashmir bee virus in honey bee excreta. J. Apic. Res. 39:103-106.
12
12. Mims, C. A. 1981. Vertical transmission of viruses. Microbiol. Rev. 45:267-286.[Free Full Text]
13
13. Morse, R. A., and N. W. Calderone. 2000. The value of honey bee pollination in the United States. Bee Cult. 128:1-15.
14
14. Ribiere, M., C. Triboulot, L Mathieu, C. Aurieres, J. P. Faucon, and M. Pepin. 2002. Molecular diagnosis of chronic bee paralysis virus infection. Apidologie 33:339-351.
15
15. Stoltz, D., X.-R. Shen, C. Boggis, and G. Sisson. 1995. Molecular diagnosis of Kashmir bee virus infection. J. Apic. Res. 34:153-160.

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Applied and Environmental Microbiology, January 2006, p. 606-611, Vol. 72, No. 1
0099-2240/06/$08.00+0     doi:10.1128/AEM.72.1.606-611.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

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