Pseudomonas contamination of a fungus-based biopesticide: Implications for honey bee (Hymenoptera: Apidae) health and Varroa mite (Acari: Varroidae) control

Biological Control 60 (2012) 312–320

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Pseudomonas contamination of a fungus-based biopesticide: Implications for honey bee (Hymenoptera: Apidae) health and Varroa mite (Acari: Varroidae) control William G. Meikle ⇑, Guy Mercadier, Fatiha Guermache, Marie-Claude Bon European Biological Control Laboratory, USDA-ARS, Campus International de Baillarguet, CS 90013 Montferrier sur Lez, 34988 St. Gely du Fesc, France

h i g h l i g h t s

g r a p h i c a l a b s t r a c t

" Fungal biopesticide used against Varroa had very negative effect on bees. " The biopesticide was found contaminated with Pseudomonas bacteria. " The bacteria interfered with fungal growth on nutritive media. " However, bacteria had little impact on bee health in cage studies.

a r t i c l e

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Article history: Received 14 October 2011 Accepted 9 December 2011 Available online 17 December 2011 Keywords: Pseudomonas fluorescens Apis mellifera Varroa destructor Beauveria bassiana Biological control Contamination 16S rRNA sequencing

a b s t r a c t The ectoparasitic mite Varroa destructor is a major honey bee pest, and its control using pathogen-based biopesticides would resolve many of the problems, such as contamination and pesticide resistance, experienced with chemical control. A biopesticide, formulated with commercially-prepared conidia of a strain of Beauveria bassiana isolated from V. destructor was tested against the mites in bee colonies in southern France. The impact of treatment on hive survivorship, weight and mite infestation levels were very different from those of previous experiments using laboratory-prepared conidia: bee hives treated with the biopesticide died at a higher rate, lost more weight, and had higher mite densities at the end of the study than control hives. The biopesticide was subsequently found to be contaminated with bacteria. Two strains of bacteria were identified, by biotyping and sequencing data of the 16S rRNA and rpoB regions, and while the strains were distinct both were Pseudomonas sp. belonging to the P. fluorescens group. In dual cultures B. bassiana growth was slowed or suppressed when bacterial cfu density was about equal or greater than that of B. bassiana. Experiments using caged adult bees showed that bees ingesting diet and sugar solution treated with B. bassiana and kept at 30 °C had significantly lower survival times than those treated with one of the bacterial strains, but the opposite was true at 33 °C. Because one arthropod (honey bees) was treated for infestation by another (V. destructor), the impact of bacterial contamination was likely more noticeable than in most uses of biopesticides, such as treating plants against phytophagous insects. To reduce such risk in biopesticide development, a systematic screening for bacterial contamination prior to field application is recommended. Published by Elsevier Inc.

1. Introduction Entomopathogenic fungi are being considered as alternatives to chemical varroacides (Chandler et al., 2001). Isolates of several species of entomopathogenic ascomycete fungi have been tested ⇑ Corresponding author. Current address: Kika de la Garza Subtropical Agricultural Research Center, USDA-ARS, 2413 E. Highway 83, Weslaco, TX 78596, USA. Fax: +1 956 969 5033. E-mail address: [email protected] (W.G. Meikle). 1049-9644/$ - see front matter Published by Elsevier Inc. doi:10.1016/j.biocontrol.2011.12.004

in laboratory and field experiments (e.g., Shaw et al., 2002; Kanga et al., 2003; James et al., 2006; Meikle et al., 2007) but of these only Beauveria bassiana (Balsamo) Vuillemin and B. varroae S.A. Rehner & Humber, sp.nov. (Rehner et al., 2011) (both Hypocreales: Cordycipitaceae) have been reported from field-collected Varroa mites, Varroa destructor Anderson and Truman (Acari: Varroidae) (Meikle et al., 2006; García-Fernández et al., 2008; Steenberg et al., 2010). B. bassiana isolates collected from Varroa mites have been shown to be virulent against Varroa mites in lab bioassays (Meikle et al.,

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2006; García-Fernández et al., 2008) and to significantly increase Varroa mite fall in field experiments (Meikle et al., 2007, 2008a). There are risks at different levels associated with the use of biopesticides against Varroa mites. Foremost is the risk to the bees themselves. Pathogens that attack mites often have a sufficiently broad host range so that they could, in theory, pose a health risk to bee colonies. B. bassiana, used by Meikle et al. (2007, 2008a, b), is known to have a particularly wide host range (Tanada and Kaya, 1993), can attack bee pupae in lab conditions (Meikle et al., 2006) and has been found naturally infecting honey bees, albeit in weakened hives (Calderón et al., 2004). It is for this reason that much emphasis has been placed in previous studies on monitoring bee colony health during and after application periods. No adverse health effects were observed from one, two or three field applications (Alves et al., 1996; Jaronski et al., 2003; Meikle et al., 2007, 2008a, b, 2009) but colony health should always be monitored to some extent, particularly when changes are made in the method or number of applications, the formulation, or the time between applications. A second risk concerns the formulation ingredients and their impact on bee health and contamination of honey, wax and propolis. The main formulation ingredient used here, carnauba wax, obtained from Copernicia cerifera Mart. or C. prunifera (Mill) (both Aracales: Aracaceae), is hydrophobic with no nutritive value for the conidia, and is permitted as a food additive in the US (US Code of Federal Regulations, Title 21, part 184, section 1978). This wax powder, mixed with small amounts of hydrated silica (0.05 g per application), has been used in previous studies with no observed side effects (Meikle et al., 2007, 2008a, b) apart from the immediate, small spike in Varroa mite fall expected from application of a powder (Fakhimzadeh, 2001; Macedo et al., 2002). A third risk is that of contamination of the biopesticide by either a chemical or a living organism. Fungal conidia are often produced in fermentation devices under conditions that are favorable to the proliferation of many undesirable micro-organisms, particularly fungi and bacteria, if strict equipment cleaning and sterilization procedures are not followed. In addition, laboratories where conidia are grown, formulated or stored often have samples of other micro-organisms on the premises. While the original objective of this study was to evaluate the impact of four successive applications of fungal conidia on honey bee colony health and on Varroa mite fall, the conidia used in the study were found to be contaminated with bacteria. The emphasis of the study was therefore shifted to identifying the bacteria, examining the field experiment results for evidence of impact of the bacteria on bee colony health, testing the bacteria for interaction with the biological control agent, and testing the impact of both the bacteria and the agent on bee health in laboratory experiments.

2. Materials and methods 2.1. Biopesticide preparation B. bassiana isolate EBCL 05002 (NRRL no. 30976; ARSEF no. 8254) conidia were obtained from a European supplier (hereafter ‘‘supplier A’’) on two occasions: April (about 12.0 g conidia) and September (about 20.0 g conidia), 2008 and stored in a refrigerator at 4 °C at the European Biological Control Laboratory (EBCL), USDA-ARS near Prades-le-Lez, Hérault department, France. A third shipment of conidia was received from another European supplier (hereafter ‘‘supplier B’’), but these conidia were not used in the biopesticide. The two batches of conidia from supplier A were mixed just prior to the first application. The dose per colony of biopesticide consisted of 0.88 g conidia mixed with 9.12 g carnauba wax powder (Strahl & Pitsch Inc., West Babylon, NY, USA) and 0.05 g hydrated silica (Hi-Sil-233, Pittsburgh Plate Glass, Pittsburgh, PA,

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USA) as a flow agent. The dose per colony of blank powder consisted of 10.0 g carnauba wax powder and 0.05 g silica. The formulations were mixed using a food processor (RoboPro 1800, SEB, Dijon, France). The density of colony-forming units (cfu) per g was determined at the time of first application by plating three sub-samples of biopesticide at three concentrations (7-, 8- and 9-fold dilution) in distilled water and Tween 80 (Merck, Munich, Germany) on potato-dextrose agar (PDA) with chloramphenicol (0.4 g/l), then counting the cfu 96 h later. The cfu analysis was repeated during the treatment period, using PDA media without antibiotic to evaluate the number of bacterial cfu in the biopesticide. Other samples also tested without antibiotic included: (1) unformulated conidia remaining from the first commercial shipment in April 2008; (2) the powder-only treatment (using same technique, with a 0.1 g sample); (3) biopesticide produced in September, 2007 (see Meikle et al., 2009); and 4) conidia sent from supplier B. All formulated and unformulated conidia were stored at 4 °C. 2.2. Field experiment On 7 October 2008, 23 honey bee colonies were transported about 90 km from Narbonne (Aude department) to a site near St. Gély du Fesc (Hérault department), Southern France. The bee colonies were kept in painted, 10-frame, wooden Langstroth brood boxes (56 l capacity) with telescoping lids, screens underneath the frames, and queen excluders on top of the brood box. Wooden pallets, approximately 1.5 m on each side, were placed under each hive. The hives were arranged into four groups of 4–7 hives each, with about 2 m between neighboring pallets and at least 15 m between groups. Sticky boards (31  42 cm, Mann Lake Ltd., Hackensack, MN, USA) were placed under the hives on 7 October and replaced on 13 and 17 October, then replaced and processed weekly until 5 December. All mites adhering to the used boards were counted. In all samples from the control group, and in pre- and post-treatment samples from the biopesticide group, 40-mite samples were taken from each board and plated on water agar (6.0 g per liter) with chloramphenicol (0.4 g per liter), incubated at 23 °C, and examined for sporulation after 15 days. If the board had fewer than 40 mites, then all the mites were plated. On 13 October, 2008, 350 to 450 live adult bees were shaken into glass jars containing 70% ethanol and the jars were sealed tightly. The jars were agitated and poured into plastic trays partially filled with water. Varroa mites were removed with forceps, the liquid was filtered to collect any remaining Varroa, and the mites and bees were counted for each sample. Also on 13 October each hive was weighed using a portable electronic balance with a precision of 50 g (OHaus Corporation model Champ CQ100L, Pine Brook, NJ, USA). Surviving hives were weighed again on 28 November. Nine colonies were randomly selected for treatment with biopesticide and 14 colonies for treatment with carnauba wax powder only. Each application of biopesticide and wax powder was loaded into a 500 ml plastic laboratory wash bottle (Nalgen Nunc International, Rochester, NY). At the moment of application, the hive lid and super were removed, the preparation blown between all brood box frames by squeezing the wash bottle, and the super and lid replaced. All the wash bottles were re-used after being thoroughly washed with soap solution and 90% ethanol. The biopesticide and the blank powder were each applied four times: on 17, 24 and 31 October, and on 6 November. Data were analyzed using SAS (SAS Institute, Inc., Cary NC, USA) software. Repeated measure analyses of variance were conducted for a linear mixed model using Proc Mixed of SAS with mite fall and the number of mites per 100 bees (both log transformed) as response variables, colony number as a random effect, and three fixed effects: treatment, date, and their interaction (a = 0.05). The

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covariance matrix of both response variables was inspected for patterns and residual plots were assessed visually for variance homogeneity. The AR(1) autoregressive model was fit to the data, and degrees of freedom were calculated using the Satterthwaite method. 2.3. Bacterial isolation The formulated B. bassiana was plated on Sabouraud Dextrose Agar (SDA) with 3% yeast extract without chloramphenicol and incubated at 35 °C for 96 h to favor the development of bacteria but not the fungus; some plates were also incubated at 37 °C. Bacterial colonies were streaked for isolation on Tryptic Soy Agar (TSA) medium (Fluka) and two types of colonies were obtained. Each colony was serially diluted to a dilution of 106 and 100 ll of the final dilution were plated onto TSA medium in Petri dishes for incubation at 26 °C for 48 h. Bacteria were also grown on King B agar medium for fluorescein production. The plates were incubated at 26 °C for 48 h. Both bacteria strains were stored in glycerol at 80 °C at EBCL. 2.4. Bacterial biotyping Single bacterial colonies were biotyped with the commercial API 20 NE miniaturized gallery (bioMérieux, Marcy l’Etoile, France) following the manufacturer’s recommendations. 2.5. Bacterial genotyping A single bacterial colony was suspended in 50 ll of sterile TE buffer (10 mM Tris [pH 8.0], 1 mM EDTA) and boiled at 99 °C for 10 min. The lysed bacterial suspension was diluted 100 fold and 2 ll was used as DNA template for the PCR. PCR amplification of 16SRNA and rpoB genes was carried out in a final volume of 25 ll containing 1 Qiagen PCR buffer (final concentration of 1.5 mM MgCl2), 0.2 lM of each dNTP, 0.3 lM of each primer, and 1 U of Taq DNA polymerase (Qiagen, Hilden, Germany). All amplifications were performed with a Perkin Elmer thermocycler 2720. The 16S rRNA gene was amplified with universal primers 27F (50 -AGAGTTTGATCCTGGCTCAG-30 ) and 1495R (50 -CTACGGCTACCTTGTTACGA-30 ) (Lane, 1991). PCR conditions were as follows: denaturing step at 95 °C for 5 min, 35 cycles of 30s denaturing at 95 °C, 30s annealing at 55 °C and 1 min extension at 72 °C, followed by a final 7-min extension step at 72 °C. The rpoB gene was amplified with primers LAPS (50 -TGGCCGAGAACCAGTTCCGCGT-30 ) and LAPS27 (50 -CGGCTTCGTCCAGCTTGTTCAG30 ) (Ait Tayeb et al., 2005). The PCR program used was an initial denaturation at 94 °C for 90 s, 35 cycles of denaturation at 94 °C for 10 s, annealing at 52 °C for 20 s, and extension at 72 °C for 1 min, plus one additional cycle with a final 5-min extension step at 72 °C. Negative controls were used in which no DNA template was added. PCR products were electrophoresed on 1% agarose gels for 15 min at 100 V and stained with SYBr SafeÒ diluted 1/10 under 254 nm UV light. PCR fragments were gel extracted using the QIAquick Gel Extraction Kit (Qiagen). Both strands of each PCR product were sequenced at Genoscreen (Lille, France) on an ABI 3730 XL TM automated sequencer with the same primer sets used for amplification. Consensus sequences were obtained after alignment of both strands using BioEdit v7.0.9 and aligned using Clustal X program version 1.81 (Thompson et al., 1997). Similarity search with existing sequence entries in NCBI public database was done using the BLASTN algorithm (BLASTN 2.2.20) (Altschul et al., 1990). 2.6. Bacteria/fungus dual cultures Cultures of B. bassiana isolate 05002 were grown on SDAY at 23 °C for 20 days. Conidia were harvested by scraping the culture surface into glass petri dishes with a metal spatula, and placing

the dishes in a crystallizing dish with silica gel for 20–24 h at room temperature to dry. Conidia were suspended in a 0.1% sterile aqueous solution of Tween 80 and distilled water to a density of 1  106 cfu per ml. Hundred microliter of the suspension was used to inoculate each of 48 plastic petri dishes (10 cm diam.). In a separate lab, bacterial cultures of each strain, hereafter PSP1 and PSP2, were grown as described above and used to prepare stock suspensions (hereafter ‘‘stock’’). The PSP1 stock contained 4  107 cfu per ml and that of PSP2 contained 1  108 cfu per ml. Each stock was diluted 10 and 100 fold, resulting in PSP1 suspensions of 4  106 and 4  105 cfu per ml, and PSP2 suspensions of 1  107 and 1  106 cfu per ml. To test the effect of any water-soluble metabolites produced by the bacteria, each strain was cultured in liquid King B medium for 48 h, on an orbital shaker (New Brunswick C24 incubator shaker) at 200 rpm. The supernatant was collected by centrifuging 20 ml of liquid medium at 4000 rpm for 10 min (Hettich centrifuge –Universal 34 R). Each supernatant was filter sterilized through an hydrophilic syringe filter 0.2 lm pore size (Sartorius Minisart High-Flow). About 4 h after inoculation of plates with B. bassiana, the same plates were then inoculated with of one of the following (four replicates per treatment): (1) 100 ll of either PSP1 or PSP2 stock; (2) 100 ll of 10-fold dilution of either PSP1 or PSP2 stock; (3) 100 fold dilution of either PSP1 or PSP2 stock; (4) 400 ll of either PSP1 or PSP2 filtrate; or (5) 100 ll of a 50:50 mix of PSP1 + PSP2 stock. As controls, four plates were inoculated with B. bassiana alone, and one plate each was inoculated with 100 ll of either the PSP1 or PSP2 stock or the PSP1 + PSP2 mix. Plates were examined after 1, 4, 10 and 12 days and the fungal growth was recorded as good, intermediate, slow, or no growth. The experiment was repeated with fresh stock of 1.2  107 cfu per ml for each strain. These new stock suspensions were then diluted to obtain suspensions of 1.2  106, 1.2  105, 1.2  104 and 1.2  103 cfu per ml, which were used to inoculate plates previously inoculated with B. bassiana using the method and concentration described above. Plates were examined after 1, 4, 6 and 12 days.

2.7. Laboratory tests on bees In order to test the effects of exposure to either bacteria strain or to B. bassiana compared to a non-exposed control, groups of bees (30–50) were placed in small cages and kept in a constant temperature and humidity (CTH) chamber at 30 °C and 50 r.h. (Percival models I36VL and I30BLL, Perry IA, USA). The back, top and sides were made of wood and the front of glass. The cages had interior dimensions of 12 cm high  9 cm wide  8.5 cm deep. The top of each cage had two holes covered with 4 mm metal mesh onto which bottles (20 ml) containing 10 g sucrose (Imperial Sugar Company, Sugar Land, TX) dissolved in 15 ml water, were placed to feed the bees. The bottom of the cage consisted of 4 mm mesh on top of a piece of cardboard, which was replaced when dirty. A piece of honey comb was suspended in the cage on a nail driven through the top. In addition to the sugar solution, each cage was provided with 5 g protein diet, consisting of granulated sugar (44.2% by weight); Bee-Pro artificial pollen (Mann Lake Ltd., Hackensack MN) (32.2% by weight), bee-collected pollen (5.4% by weight) and water (18.3% by weight), placed in a dish at the bottom of the cage. Bacteria cells were produced as described above for the 2nd and 3rd experiments. Fungal conidia were produced as described above for the 2nd experiment; for the 3rd experiment conidia were obtained from ARSEF (USDA-ARS, Ithaca, NY). Conidia were dried and kept refrigerated in a tightly-sealed container. Bacteria cells were suspended in a phosphate buffer consisting of 8.75 g K2HPO4 and 6.75 g KH2PO4 dissolved in 1 l distilled water and kept refrigerated; each strain was kept separately in its own

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suspension. Prior to the start of the experiments, the cfu densities of the conidia and both bacteria strains were measured as above. To collect the bees, the same protocol was followed for each experiment: a frame of sealed brood was obtained from experimental hives and placed in a wood and mesh cage in an incubator at 30 °C. Emerged bees were collected over 48 h and placed immediately in cages in the CTH chamber. During the course of the experiment, dead bees were removed 4–5 times per week. Three experiments were conducted. The 1st experiment consisted of 19 cages of untreated bees, conducted in order to determine the expected longevity of bees under these conditions. In the 2nd and 3rd experiments, bee cages were divided into four treatment groups: PSP1, PSP2, B. bassiana and control. In the PSP1 and PSP2 groups, 10 ll of the respective bacterial suspension was added to 1 g protein diet and to 20 ml sugar solution one week after the addition of the bees for each cage. For the B. bassiana group, 1 mg of conidia was added to the diet and sugar solution one week after adding the bees. Nothing was added to the control group food. As the bees consumed the sugar solution and the diet, it was replaced with food treated in an identical fashion. The 2nd experiment was conducted at 30 °C, with six replicate cages per treatment group. The 3rd experiment was conducted at 33 °C, to examine possible temperature interactions, with seven replicate cages per treatment group and nine control replicate cages. To avoid contamination between experiments, new cages with about the same interior dimensions were built for the 3rd experiment using Plexiglas. The growth rate of the bacteria and fungus in the sugar solution used in the cage experiments was evaluated to improve the estimate of the inoculum exposure to the bees. Four sets of three 20 ml glass vials were sterilized and filled with the same sucrose and water solution (10 g sucrose in 15 ml water) used in the cage experiments. Each set of vials was treated the same as for the cage experiments, that is inoculated with either 10 ll of the PSP1 stock solution used in the cage studies, 10 ll of the PSP2 stock solution, 1 mg of the B. bassiana conidia, or nothing (control). After 7–10 d, 10 ll samples from the fungal treatments were examined at 1000 (Meiji Techno, Santa Clara, CA, USA) for hyphae and spores. Bacterial cell densities were estimated using the techniques described above for cfu estimation and the intrinsic rate of increase of the bacterial population per d was calculated:

r¼

lnðNt Þ  lnðN 0 Þ t

where r is the intrinsic rate of increase per d, N0 is the initial cell population, Nt is the final cell population and t is time in d (after Gutierrez, 1996). The rate of increase per d was used to adjust the estimated exposure of the bees to the bacteria. The experiment was conducted four times at both 30 and 33 °C. 2.8. Statistics Data were analyzed using SAS software (SAS Institute, Inc., Cary NC, USA). Multiple regression analyses and repeated measures analyses (a = 0.05) were conducted for linear mixed models using PROC MIXED (Littell et al., 1996). All response variables expressed as proportions were arcsine square-root transformed, as is recommended for percentages that cover a large range of values (Steel and Torrie, 1960). In analyses of field data, the experimental units were bee hives and were evaluated using repeated measures with treatment as a fixed effect. Degrees of freedom were calculated using the Satterthwaite method, type III sums of squares were used where applicable and residual plots were assessed visually for variance homogeneity. Post hoc contrasts of the least squares means differences were conducted for all significant factors, using the Bonferroni adjustment for the t-value probability. Test-of-effect

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slices were used to evaluate significant interaction effects. Adult bee longevity for cage trials was evaluated using Kaplan–Meier method for estimating the continuous-time survivor function (Singer and Willett, 2003), using a log rank analysis with pairwise comparisons (SigmaPlot 11.0). Each cage was considered as an experimental unit, and median survivorship per cage was used to generate the survival functions. 3. Results 3.1. Field experiments The B. bassiana cfu density was estimated as 2.0  109 per g biopesticide, which was low but similar to that measured in previous experiments (see Meikle et al., 2008b). Bacterial density for that sample was estimated at 8.3  107 cfu per g biopesticide. In the conidia sent by supplier B bacteria were also detected at 1.4  104 cfu per g, compared to B. bassiana cfu of 2.3  109 per g; these conidia were not used in the field experiment and the bacteria were not identified. No bacteria were found in conidia sent from supplier A in April 2008, nor were any detected in samples of the powder-only treatment (carnauba wax + hydrated silica alone) or the biopesticide samples from previous field experiments. The powder-only treatment was prepared using the same equipment just prior to preparation of the biopesticide itself, this result suggests that no bacteria were present in the wax powder, the hydrated silica powder, or the equipment itself. Overall there was no difference between treated and control hives in terms of mite fall. Mite fall changed over time (F1,167 = 4.17, p < 0.0001) but neither treatment (p = 0.50) nor treatment  date (p = 0.11) were significant (Fig. 1A), in spite of a spike in mite fall observed among control hives on one sampling occasion two weeks after the end of the application period. Infected mites were found in most hives (n = 13) before biopesticide application, but the proportion infected mites in a given hive was low: from 1 to 9 mites were found infected out of 40. During the application period the average (s.e.) proportion of infected mites among control colonies ranged over time from 0.091 (0.038) to 0.389 (0.101), probably due to robbing of the treated hives. After the application period, infection levels ranged over time from 0.918 (0.041) to 0.975 (0.016) among treated hives and from 0.188 (0.049) to 0.441 (0.070) among control hives. Treatment was associated with a problem not observed in previous experiments using the same fungal isolate and formulation: high hive mortality. Just four of the nine treated hives survived until the end of the experiment, compared to 13 of the 14 control hives (Fig. 1B). Treated hives had significantly higher phoretic mite densities than untreated hives after application period (F1,21 = 10.03, p = 0.0047); date was also significant (F2,37 = 17.15, p < 0.0001) but not the interaction (p = 0.11) (Fig. 1C). Treated hives also lost more weight: surviving treated hives lost an average (s.e.) of 4.38 kg (0.38), which was significantly more (t15 = 2.84, p = 0.0123) than the 2.71 kg (0.39) lost by surviving control colonies. 3.2. Bacterial biotyping Two strains of bacteria were found, PSP1 and PSP2, the colonies of which differed from each other morphologically: PSP2 colonies were larger and highly mucilaginous compared to those of PSP1. Both strains produced a fluorescent pigment on King’s medium and neither grew at 37 °C. Biotyping strain PSP2 with the commercial API 20NE Kit produced the numerical code 0157455 (positive reactions: arginine dihydrolase, gelatinase, assimilation of glucose, arabinose, mannose, mannitol, gluconate, caprate, malate, citrate, and oxidase; negative reactions: nitrate reduction, indole, glucose fermentation, urease, esculin hydrolysis, b-galactosidase, assimilation of N-acetylglucosamine, maltose, adipate, and phenylacetate), which

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between the two strain sequences after alignment. Similarity searches were conducted with all Pseudomonas rpoB sequence entries available in the NCBI GenBank. The three best matches for both sequences were observed for P. fluorescens strain LMG 5825 (98% rpoB similarity score; accession no. AJ748146), P. fluorescens strain LMG 5830 (98% rpoB similarity score; accession no. AJ748152) and P. marginalis strain LMG 5170 (97% rpoB similarity score for PSP1 and 96% for PSP2). Ait Tayeb et al. (2005) recently re-classified the first two strains as unnamed species and the third one as P. veronii. The next closest phylogenetic taxa (96% rpoB similarity score) were P. azotoformans type strain CIP 106744T, P veronii type strain CIP 104663T and P. fluorescens type strain CIP 69.13, all belonging to the P. fluorescens subgroup of Palleroni rRNA group I (Anzai et al., 2000; Mullet et al., 2010). We identified the two strains as Pseudomonas sp, belonging to the P. fluorescens group in reference to Anzai et al. (2000) and more precisely to the P. fluorescens subgroup in reference to Mulet et al. (2010). 3.4. Bacteria/fungus interaction

Fig. 1. Results of a field experiment involving four applications (represented by vertical dashed lines) of either biopesticide (n = 9; triangle with dotted line) or blank wax powder (n = 13; circles with solid line) conducted in fall, 2008, near St. Gely du Fesc (34), France. (A) Average (with s.e. bars) varroa mite fall onto sticky boards over time; (B) Proportion hive survivorship; (C) Phoretic mite density (no. mites per 100 bees).

corresponded to P. fluorescens with %id = 99.9 and T = 0.85 (%id 6100 as relative proximity to the different taxa of bioMérieux data base and T 61 as proximity to the most typical profile in each of the taxa). Biotyping strain PSP1 with the same kit produced the code 0047555 (positive reactions: assimilation of glucose, arabinose, mannose, mannitol, N-acetylglucosamine, gluconate, caprate, malate, citrate, and oxidase; negative reactions: nitrate reduction, indole, glucose fermentation, arginine dihydrolase, urease, esculin, gelatinase, bgalactosidase, assimilation of maltose, adipate, and phenylacetate) corresponding to P. fluorescens with %id = 99.8 and T = 0.9.

In the 1st experiment, no B. bassiana growth was observed on any plate treated with bacteria until day 10. On day 10, slow and intermediate growth was observed on two plates treated with the stock suspension (1  108 cfu per ml) of PSP1; the remaining plates in that group showed no growth. No further change was noted by day 12. In all other plates with both fungal and bacterial suspensions only bacterial colonies were observed. B. bassiana growth was healthy in plates treated with bacterial filtrate. Growth was vigorous in all control plates with no signs of cross-contamination. In plates treated with PSP1 + PSP2 dense colonies of both strains were observed. In the 2nd experiment, some growth was observed in the low density treatment (1.2  103 cfu per ml) on day 1. By day 4, either slow or intermediate growth was observed in all plates treated with the PSP1 strain, and in all plates except one (at 1.2  106 cfu per ml) treated with PSP2. By day 6 intermediate or good growth was observed in all plates except those treated with high density of either strain, in which slow or no growth was observed with PSP2. By day 12, good growth was observed in all plates treated at 1.2  105 cfu per ml or lower, while in the high density treatment only slow fungal growth was observed (Fig. 2). No crosscontamination was observed in control plates. In summary, in the 1st experiment with the PSP1 strain, some fungal growth was observed at 4  107 cfu per ml but none at densities either one or two magnitudes lower, while no fungal growth was observed in any plates treated with PSP2 at densities of 1  106 to 1  104 cfu per ml. In the 2nd experiment some fungal growth was observed with both bacterial strains at 1.2  106 cfu per ml, and good growth at all bacterial cfu densities lower than that. We observed on several occasions that fungal growth among plates was inconsistent, even for plates treated with the same suspensions of conidia and bacteria. Strain PSP1 seemed to have marginally less impact on B. bassiana growth than strain PSP2.

3.3. Bacterial genotyping 3.5. Laboratory tests on bees The 854-bp 16S rRNA sequences of strains PSP1 and PSP2 were deposited in the GenBank database under accession nos. FJ948082 and FJ948083, respectively. No difference between the two strains was found in those 854 bp. The unique sequence was compared with all of the eubacterial 16S rRNA sequences available in the NCBI GenBank database. The best similarity matches were observed for Pseudomonas strains. The 16S rRNA sequence similarity score of 100% indicates both strains are Pseudomonas species. The 1155-bp rpoB sequences of strains PSP1 and PSP2 were deposited in the GenBank database under accession nos. FJ948084 and FJ948085, respectively. Only two mismatches were observed

In the 1st trial, with untreated bees, data were excluded for the first 6 d because of high mortality due to feeding problems which were resolved. Average bee survival time per cage was 27.1 d and average maximum survival time per cage was 47.0 d. For the 2nd experiment, conducted at 30 °C, the exposure per bee cage was 1.2  105 cfu for the PSP1 treatment, 4.0  105 cfu for the PSP2 treatment, and 9.2  107 cfu for the B. bassiana treatment. Analysis of survival times showed that treatment had a significant effect (log rank statistic = 10.897, d.f. = 3, P = 0.012) (Fig. 3). Pairwise contrasts showed that bees fed PSP1-contaminated food lived

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Fig. 2. Photographs of petri dish cultures after incubation for 12 days. Top photo (upper left) shows control plates treated with 100 ll of a 1  106 cfu per ml suspension of Beauveria bassiana conidia only. White indicates B. bassiana growth, yellow indicates area covered by bacterial colonies. Photos in the middle row show, from left to right, plates treated with B. bassiana (same concentration) and with 100 ll of a suspension of bacterial strain PSP2 at concentrations of 1.2  103, 1.2  104, 1.2  105 and 1.2  106 cfu per ml; the central plate in each photo shows a control plate treated only with bacterial suspension at the respective concentration. Similarly, photos in the bottom row show, from left to right, plates treated with B. bassiana and with a suspension of strain PSP1 at concentrations of 1.2  103, 1.2  104, 1.2  105 and 1.2  106 cfu per ml, along with control plates.

significantly longer (37.9 d) than those fed B. bassiana-contaminated food (27.1 d). No other contrasts were significant. Bees in the control treatment had a mean survival time of 35.7 d and an average maximum survival of 53 d. For the 3rd experiment, conducted at 33 °C, the exposure per cage of bees was 5.6  104 cfu for the PSP1 treatment, 6.0  104 cfu for the PSP2 treatment, and 3.8  106 cfu for the B. bassiana treatment. Analysis of survival times showed that treatment had a significant effect (log rank statistic = 10.235, d.f. = 3, P = 0.017). Pairwise contrasts showed that bees fed PSP1-contaminated food lived significantly shorter (20.1 d) than those fed B. bassiana-contaminated food (25.9 d) and no other contrasts were significant. Bees in the control treatment had a mean survival time of 21.9 d and an average maximum survival of 33 d. Sugar solution treated with B. bassiana showed hyphal growth on SDAY agar plates, and microscopic examination of solution samples revealed hyphal fragments as well as blastospores and submerged conidia, which are typically produced in rich-broth submerged cultures and nutrient-poor submerged cultures, respectively (Holder et al., 2007). Spore density was estimated at <103 blastospores or submerged spores per ml. Bacterial grew in the sugar solution but growth was inconsistent. PSP1 had a high rate of increase, 0.51 ± 0.01, in one experiment at 30 °C, and would thus be expected to be 39 times higher after 7 d, but populations failed entirely in three other experiments and the strain did not last for more than 2–3 d at 33 °C. PSP2 was more consistent: r = 0.50 ± 0.15 across two experiments at 30 °C and r = 0.41 ± 0.13 across two experiments at 33 °C, but PSP2 populations also failed in one experiment at 33 °C. Based on these data, PSP2 densities would be expected to be 33 times higher at 30 °C and 18 times higher at 33 °C after 7 d.

4. Discussion A biopesticide formulated with commercially-prepared conidia of a strain of B. bassiana, itself isolated from V. destructor, was tested against the mites in bee colonies. The effects of biopesticide application were very different from those of previous experiments with laboratory-prepared conidia: bee hives treated with the biopesticide died at a high rate, lost more weight, and had higher mite densities than control hives. The biopesticide made with commercially-prepared conidia was subsequently found to be contaminated with bacteria in the P. fluorescens group. Subsequent laboratory experiments showed that that role of the bacteria in the bee/mite/fungus system was likely complex. While this field experiment was intended to be an extension of previous work done by our group on multiple applications of biopesticide to control V. destructor, the biopesticide itself was for the first time strongly linked with weakening and killing hives. That hives occasionally die in experiments of this nature is not surprising since they usually have moderate to heavy V. destructor infestations at the start, but losses in previous experiments have been minor: no hives were lost among the 22 hives used in a field experiment using the same kind of biopesticide with one application (Meikle et al., 2008b) and only one hive lost, in the untreated control group, in a field experiment involving two applications (Meikle et al., 2008a). In an experiment with 35 hives involving three consecutive applications of biopesticide, three hives were lost: one each in the biopesticide, powder-only, and untreated control groups (Meikle et al., 2009). In the experiment presented here, most treated hives were lost while only one from the control group was lost. Surviving treated hives also lost more weight than other hives, probably due to robbing and bee mortality. The impact on

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Fig. 3. Kaplan–Meier survival functions for laboratory experiments conducted with caged bees fed diet and sugar solution treated with either P. fluorescens strain PSP1, P. fluorescens strain PSP2, B. bassiana conidia, or nothing (control). (A) Survival functions for 2nd cage experiment conducted at 30 °C (see text); points show censored data for cages in which bees (5 or fewer) were alive at the end of the experiment after 64 d. (B) Survival functions for 3rd cage experiment conducted at 33 °C (see text); there were no censored data.

the V. destructor population is difficult to quantify, since V. destructor density depends largely on bee colony size and health. While mite fall was not different between groups, phoretic mite densities were significantly higher in treated hives. Bacteria were found only in the biopesticide used in the field experiment described here; powder samples from the blank treatment as well as biopesticide samples from a previous experiment (described in Meikle et al., 2009) tested clean. No bacteria were found in the wax powder, silica, or in the equipment used to formulate the biopesticide. In all our previous experiments B. bassiana conidia were produced at EBCL, under strict axenic conditions and with antibiotic in the growth media (Meikle et al., 2006, 2007, 2008a, b, 2009), but in the present study the conidia were commercially-produced. Unformulated samples from another supplier were also contaminated with bacteria, albeit at much lower levels, indicating this problem may be widespread and represent a genuine quality control issue. Because we were treating one arthropod (honey bees) for infestation by another (V. destructor), the impact of bacterial contamination was likely more noticeable than in most uses of biopesticides, such as treating plants against phytophagous insects. As an explanation for the poor field results the question remained: what kind of bacteria were they, and could they have caused the problems we observed? We used two approaches to identify the bacteria: biotyping with commercial API 20 NE miniaturized gallery; and genotyping of the 16S rDNA and rpoB genes. Molecular tools, including primers specific for universally conserved bacterial 16S rDNA, are being used successfully to identify bacteria from different environments, such as food, clinics, and industrial water systems (Weisburg et al., 1991; Iwamoto et al., 2000). Anzai et al. (2000) recognized that, using the 16S rRNA, at

least 57 species belong to Pseudomonas (sensu stricto), including P. aeruginosa, the type species of Pseudomonas (a list of species and sub-species is available at www.bacterio.cict.fr/p/pseudomonas.html). Although the 16S rRNA gene is a powerful tool for genus assignments, it does not discriminate sufficiently at the inter-species level within the Pseudomonas genus (Ait Tayeb et al., 2005; Mullet et al., 2010). Ait Tayeb et al. (2005) analyzed Pseudomonas phylogenetic relationships using rpoB gene sequences, since it codes for the RNA polymerase beta subunit, is a highly conserved, and has been used as a signature for identifying bacteria, including Pseudomonas. Ait Tayeb et al. (2005) found that taxonomic resolution within Pseudomonas was improved by a factor of 3 when rpoB is used rather than genes such as rrs, although some lineages and intragenic clusters in P. fluorescens groups remain confused in the rpoB phylogenetic tree. Sequencing rpoB did not permit identification to species, but allowed identification to group and showed that the two strains are slightly different. A more reliable species signature in this particular group is expected as more genes are sequenced (Mullet et al., 2010). The bacteria were thus identified as two strains of the P. fluorescens group, belonging to the Palleroni rRNA group I. Assuming that the environment and the baseline hive health in this field experiment were similar to those in previous experiments conducted by our group, the most likely explanations for the high mortality among treated hives are: (1) the impact of four applications (compared to fewer applications); (2) bacterial contamination or (3) an interaction between the two. In other words, were the bacteria the problem? Pseudomonas fluorescens is known as a food spoilage agent in milk, fish and vegetable preparations (Dogan and Boor, 2003; Liao, 2009), and has been found contaminating wastewater (Sacchetti et al., 2007). Strains of P. fluorescens have been found to grow on most kinds of honey (Mundo et al., 2004) so the bacteria may have attacked the food stores in the hive, interfered with feeding and brood provisioning, and/or produced insect toxins (Péchy-Tarr et al., 2008). Strains in the P. fluorescens group have been isolated from suppressive soils and found to inhibit growth and activity of fungal plant pathogens (Hass and Dèfago, 2005). The two Pseudomonas strains were shown to slow or suppress B. bassiana growth on nutritive media when the cfu densities of the bacteria and the fungus were about equal. While the bacterial cfu density in the biopesticide formulation was only about 4% that of the fungal cfu, it should be noted that the lab tests were designed slightly in the favor of the fungus, as the fungus was plated 4–6 h before the bacteria, and that the bacteria were found to grow much better than the fungus at 33–36 °C, temperatures likely to be encountered in the brood area (Winston, 1987). While field trials would have been ideal for studying the interaction of B. bassiana, the bacteria and honey bees, such trials were not possible owing to regulatory and human health issues. Bee cage experiments were conducted under level II quarantine conditions at the Subtropical Agricultural Research Center, USDA-ARS, in Weslaco, Texas, and dust applications of either the fungus or the bacteria were not permitted. Bee cage experiments did not indicate a clear impact of either P. fluorescens or B. bassiana in the bee diet. At 30 °C, bees fed diet and sugar solution treated with PSP1 lived longer than those with food treated with B. bassiana, but the opposite was true at 33 °C. In neither test did the survival times of the treated bees differ from that of the control group, so the cage results did not fully explain the colony death rate in the field experiment. Based on cfu calculations from the biopesticide application and assuming 10,000 bees per hive, a group of 40 bees would have been exposed to about 3.3  106 bacterial cfu on average and about 8.0  107 fungal cfu in the field trial. Total exposure levels (diet and sugar solution) in the cage experiments were intentionally lower than those estimated for the field, on average 2–12% of the estimated field levels for bacteria and 4–12% of the field levels for B. bassiana, in order to

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simulate exposure through ingestion since much of the biopesticide in the field landed on hive parts or on bee bodies where it either fell off or was removed. Experiments to improve the estimate of inoculum exposure to the bees, at least with respect to the sugar solution, showed that while both bacterial strains and B. bassiana grew at 30 °C, PSP1 growth was inconsistent and it did not grow at all at 33 °C. A simple concentrated sugar solution is likely not their ideal medium. However, both bacterial strains grew well on nutritive agar at 33 °C, and open comb cells of nectar and pollen may have provided substrate for bacteria and fungi growth in treated hives. B. bassiana may attack bees through the cuticle, so in that regard the cage experiments may underestimate B. bassiana impact. In the concentrated sugar solution, B. bassiana was observed to produce low densities of blastospores and submerged spores, which can cause higher insect mortality than conidia when applied topically or injected into the hemocoel (Holder et al., 2007) although that was not done here. There are also reasons to suspect that B. bassiana impact on bee health in the field would be overestimated by the cage studies. As Evans and Spivak (2010) pointed out, honey bee colonies have different kinds of resistance, ranging from individual immunological and physiological responses, to auto- and allo-grooming, to the colony level, which includes specialized hygienic behaviors. Niu et al. (2010) reported that propolis (bee-collected plant resins) helped bees break down the mycotoxins found in bee bread by enhancing the activity of enzymes involved in detoxification. Some of these features of colony-level resistance were not present in the laboratory experimental units. In this study unusual results from a field experiment led to the discovery of contaminating bacteria. However, subsequent testing in bee cages indicated that the mere presence of the bacteria was unlikely to be the sole cause of the unusual results and any role the bacteria had probably involved interactions with environmental factors not present in the cage studies. An interaction was demonstrated between cultures of P. fluorescens and B. bassiana, so the effect of P. fluorescens may be more complex than that of simply killing bees. Pathogens intended for biopesticides should be checked for bacterial and fungal contamination, whether they were produced under small-scale conditions in a research laboratory or under commercial conditions. Bacteria such as P. fluorescens, which are common contaminants of wastewater and stored products and which interact with fungal pathogens, pose particular risks to biopesticide production and application. Acknowledgments The authors would like to thank F. Annas and V. Girod at the Association de Développement de l’Apiculture Professionnelle in Languedoc Roussillon (ADAPRO-LR) and R. Humber (USDA-ARS) for their support, R. Diaz, C. Gracia, R. Medrano, and C.S. Patt for their help in the laboratory, J. Patt and K.D. Murray (both USDAARS) for helpful comments on the manuscript, and two anonymous reviewers for their help in improving the manuscript. References Ait Tayeb, L., Ageron, E., Grimont, F., Grimont, P.A.D., 2005. Molecular phylogeny of the genus Pseudomonas based on rpoB sequences and application for the identification of isolates. Research in Microbiology 156, 763–773. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. Journal of Molecular Biology 215, 403–410. Alves, S.B., Marchini, L.C., Pereira, R.M., Baumgratz, L.L., 1996. Effects of some insect pathogens on the Africanized honey bee, Apis mellifera L. (Hym.: Apidae). Journal of Applied Entomology 120, 559–564. Anzai, Y., Kim, H., Park, J.Y., Wakabayashi, H., Oyaizu, H., 2000. Phylogenetic affiliation of the pseudomonads based on 16S rRNA sequence. International Journal of Systematic and Evolutionary Microbiology 50, 1563–1589. Calderón, R.A., Rivera, G., Sanchez, L.A., Zamora, L.G., 2004. Chalkbrood (Ascosphaera apis) and some other fungi associated with Africanized honey bees (Apis mellifera) in Costa Rica. Journal of Apicultural Research 43, 187–188.

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