A non-destructive method to detect pathogens in bumble bees for conservation research

doi:10.21203/rs.3.rs-2753591/v1

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A non-destructive method to detect pathogens in bumble bees for conservation research

https://doi.org/10.21203/rs.3.rs-2753591/v1

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Wild bees are declining worldwide, and many species are now threatened with extinction. Decline is caused by a combination of threats, including disease spillover from managed bees that reduces health of wild bees. An increasing number of studies thus aim to characterize bee health. The common approaches, however, require sacrificing tens to hundreds of individual bees per site or species, with reports of several thousand individuals collected per study. Considering the widespread need to assess bee health, this sampling approach is not sustainable, especially for endangered populations or species. Here, we propose a non-destructive method to assess parasite loads of wild-caught bumble bees. The standard protocol consists of net-capturing individual bumble bees and placing them in a 10 cm (diameter) petri dish to collect faeces. Although this approach is frequently used in laboratory settings, it is not in the field, because of the low success in collecting faeces. Placing bumble bees in a previously refrigerated cooler, we significantly improved faecal collection in the field from 76% with the standard protocol to 86% with the cooler protocol. We also successfully identified spores and cells of two common gut parasites Crithidia spp. and Vairimorpha spp. in faecal samples. The efficacy of the cooler protocol, combined to the low-cost and widespread availability of the equipment should promote its use in field studies.

Implication for insect conservation: As there are calls to reduce destructive sampling methods in bee research, using the updated cooler protocol will contribute to achieving this goal. This opens future avenues of combining this non-destructive approach to assess bee health with molecular tools.

intestinal parasites

faeces

bumblebee

conservation

non-lethal sampling

insect declines

Bombus

Crithidia

Vairimorpha

Nosema

Wild bees are declining worldwide from a combination of stressors, including habitat loss, pesticides, malnutrition, climate change, invasive species and disease spillover from managed bees (Aldercotte et al., 2022; Baron et al., 2017; Botías et al., 2021; Burnham et al., 2021; Cameron et al., 2011; Cameron & Sadd, 2020; Colla et al., 2012; Goulson et al., 2015; Graystock et al., 2013; Jackson et al., 2022). The impacts of environmental changes on bumble bees (Bombus: Apidae), an important group of wild bee pollinators, are increasingly documented (Botías et al., 2021; Cameron et al., 2011; Guzman et al., 2021; Jackson et al., 2022; Siviter et al., 2021; Su et al., 2022). A growing number of investigations are assessing health of wild bumble bee individuals, colonies, and populations (Cameron & Sadd, 2020; Garlin et al., 2022; Giacomini et al., 2018; McNeil et al., 2020; Pislak Ocepek et al., 2021; Trillo et al., 2021; Tsvetkov et al., 2021) and more than 50% of studies examining causes of bumble bee decline considered parasitic infections (Cameron & Sadd, 2020).

Conventional approaches to assess adult bee health use either morphological (e.g., wing morphology, ectoparasites, body mass or size) or physiological/molecular approaches (e.g., assessment of body fat, prevalence and loads of intestinal parasites or presence of viruses in body tissues) (Garlin et al., 2022; Giacomini et al., 2018; McNeil et al., 2020). Whereas morphological approaches provide information about general body condition and health status of individuals, physiological approaches can assess the identity, prevalence, and abundance of pathogens. These approaches, however, require killing a significant number of individual bees (N = 20 to 300/population or species) to accurately measure wing or body size/mass, to extract the gut or to grind bodies, before conducting microscopic or molecular analyses (e.g., RT-PCR or qPCR) (Babin et al., 2022; Blaker et al., 2014; Garlin et al., 2022; Giacomini et al., 2018; Graystock et al., 2020; McNeil et al., 2020; Tsvetkov et al., 2021). Considering the increasing number of studies assessing bee health, with around 400 conservation-based studies per year on bumble bees alone (Cameron & Sadd, 2020), collecting as many as several thousand individuals per study, this approach is not sustainable and raises conservation concerns (Miller et al., 2022; Montero-Castaño et al., 2022). The conservation impact of repeated and widespread lethal sampling is rarely studied and merits further investigation (Montero‐Castaño et al., 2022). The few studies on the topic report contrasting effects of repeated lethal sampling (Gezon et al., 2015; Gibbs et al., 2017). Although the impact of scientific research may be small compared to human-mediated impacts such as habitat loss, pesticides, pathogen spillover or climate change (Miller et al., 2022; Montero‐Castaño et al., 2022; Sánchez-Bayo & Wyckhuys, 2019; Wagner, 2020), the effects of such repeated destructive samplings are mostly unknown and likely to vary between populations, species or even castes of bees (Montero‐Castaño et al., 2022). Furthermore, sampling still remains an additional pressure on pollinators, which could be especially damaging to rare or endangered wild bee species and communities (Miller et al., 2022).

Two pathogens that are commonly monitored to assess bee health are Crithidia spp. (Trypanosomatidae) and Vairimorpha spp. (Nosematidae) (Cameron & Sadd, 2020; Graystock et al., 2020; Grupe & Quandt, 2020). Cells or spores of these parasites are horizontally transmitted (i.e., via an oral-faecal route), either through contamination of flowers during foraging or through contamination of the nest (Graystock et al., 2015; Graystock et al., 2020; Grupe & Quandt, 2020). They have been identified in a large variety of bee species and genera (Cameron & Sadd, 2020; Figueroa et al., 2021; Gillespie, 2010; Grupe & Quandt, 2020). In bumble bees, depending on infection intensity, the parasites reduce gyne production, queen fitness and colony growth (Brown et al., 2000; Goulson et al., 2018), impair learning and foraging ability of workers (Gegear et al., 2005; Goulson et al., 2018), increase mortality of both males and workers (Grupe & Quandt, 2020; Otti & Schmid-Hempel, 2007) and ultimately lead to colony decline under field-realistic stressful conditions (Brown et al., 2000). In addition, Crithidia and Vairimorpha are common parasites found in managed bees (i.e., commercial bumble bees and domestic honey bees) that raise important concerns regarding pathogen spillover to native and/or wild bees (Grupe & Quandt, 2020; Strange et al., 2023), in which they are able to replicate (Ngor et al., 2020), though their pathogenicity remain to be determined for several species. Finally, rare and endangered bumble bees are at higher risk of pathogen spillover and show greater prevalence (i.e., rate of infection) of Vairimorpha than common and non-threatened species (Averill et al., 2021; Cameron et al., 2011; Gillespie, 2010), increasing the need to monitor parasites without causing harm threatened populations or endangered bumble bee species, such as B. affinis or B. terricola (Colla & Packer, 2008; Jacobson et al., 2018). It is thus paramount to develop and implement non-destructive, widely accessible, and reproducible practices that work across species and castes, to monitor bee health in ecology and conservation research.

A common non-destructive approach to monitor vertebrates’ health in ecology and conservation consists of collecting and screening their faeces to assess the presence of parasites or other pathogens (Biswas et al., 2019; Darimont et al., 2008). Here we present the step-by-step development of a non-destructive protocol to i) collect adult bumble bee faeces in the field, ii) increase the rate of faeces collection, and iii) confirm the presence of the gut parasites Crithidia spp. and Vairimorpha spp., both commonly found in bumble bee faecal samples (Cameron & Sadd, 2020; Graystock et al., 2020; Grupe & Quandt, 2020). We adapted a laboratory-validated faecal screening protocol, which consists of placing individual bees inside petri dishes until they defecate (Chen et al., 2006; Gomez-Moracho et al., 2021), to field conditions. This method is not commonly deployed in the field due to the low success in collecting faeces, leading to increases in sampling effort. We hypothesized that reduced stress during handling could increase faecal collection success, by placing individuals in a dark and cool low-sensory environment. Since bees often defecate in the nest (Adler et al., 2021; Bodden et al., 2019; McArt et al., 2014), i.e., when in a low-stress state, we hypothesized that a low-sensory environment would improve collection success, determined as defecation probability. This methodological paper is a step in the development of a non-destructive approach to assess bumble bee health through faeces collection.

Experimental design: bee captures and faeces collection

We gathered data on faecal collection success using two protocols (standard protocol or updated cooler protocol, two samplings per protocol) implemented during four independent studies conducted on bumble bee health. Although it would have been ideal to compare the success of both methods within the same study, each of these studies was conducted for a separate additional purpose, constraining us to use a single protocol per study for consistency. The standard protocol consisted of placing bees in a shady environment during faeces collection to prevent thermic stress and limit risk of death (Fig. 1; studies 1 and 2). The updated cooler protocol consisted of placing bees in a dark and cool low-sensory environment (studies 3 and 4; see Fig. 2). In all four studies, adult bumble bees were net-captured by hand in the field (see details below). Bees were placed individually in transparent, 10 cm diameter, petri dishes for faeces collection and species identification. Net capturing is an efficient and minimally invasive approach to catch bumble bees (Montero-Castaño et al., 2022; Tronstad et al., 2022). For each bumble bee and in each study, we recorded the site and date of captures (Fig. 1). Each bumble bee was released from the petri dish after all faecal samples were collected. Faecal samples were immediately placed in a standard, low-cost and widely available portable hard case cooler (12–17 liters, Coleman, USA) with ice packs or on ice (either work for short-term storage of samples) until screening under the microscope (see details for each study). If bees did not defecate after one hour, they were considered as not having defecated. These bees were released or transferred into a vial and placed on ice until completion of the faecal sampling session at each site, after which all captured bees were released.

Studies and sites

a) Standard protocol

Study 1

Captures were conducted once a week at five agricultural sites in the vicinity of Lac Brome (Southern Québec, Canada; 45° 12' 18.4752'' N and 72° 33' 32.8320'' W) from August 6–17, 2020, between 10:00 and 17:00. We conducted three capture events in total: one capture per site per week, and the order of the sites was randomized each week. Ambient temperatures ranged from 17–34°C during sampling sessions. A total of 48 bumble bees were captured during this period. Upon capturing, each bee was immediately placed in an individual petri dish, identified to species, and placed in a shady environment, avoiding direct exposure to the sun to prevent thermic stress and risk of death. At an ambient temperature of 30°C, bumble bees already have difficulty maintaining their body temperature below the lethal threshold of 45°C (Heinrich, 1977). Placing a bee inside a petri dish in the sun when ambient temperatures are above 30°C, without ventilation, exposes the individual to extreme temperatures, likely to be lethal. Bees were maintained in the petri dish in this shady environment until faeces were collected or for up to 1 hour. None of the bees placed in the shade died during the study, but one bee that had been forgotten in the sun for several minutes by the experimented died, likely because of overheating. We inspected petri dishes for faeces every 10 min. When faeces were confirmed, we placed the side with faeces on top by gently turning the petri dish over, to limit destroying the sample due to the bumble bee walking inside. Of the 48 bees captured, we successfully collected faecal samples from 30 bees (62.5% success). Samples were screened for parasites under the microscope within 24 hours after collection.

Study 2

Captures were conducted every other week from 08:00 to 16:00 inside and surrounding commercial greenhouses around Compton (Southern Québec, Canada, 33° 53' 45.0492'' N and 118° 13' 12.2520'' W) from April 22nd to August 13th, 2021. Upon being placed in a petri dish, bumble bees were placed in a shaded environment outside of the greenhouses (standard protocol described in study 1). Two bees died in the petri-dishes during the experiment. Ambient temperatures ranged from 15 to 32°C during sampling sessions. A total of 398 bumble bees were captured, from which we collected 288 faecal samples (72.4% success). Faecal samples were screened for parasites under the microscope within 24 hours after collection using a Neubauer hemacytometer.

b) Updated cooler protocol: optimizing faecal collection success

Study 3

Captures were conducted once every other week at four agricultural sites in the vicinity of Alma (Québec, Canada; 45° 12' 18.4752'' N and 72° 33' 32.8320'' W) from June 1st to September 9th, 2021, between 10 am and 5 pm. Ambient temperatures ranged from 13 to 32°C during sampling sessions. In study 1, faeces collection seemed especially limited in the afternoon, when ambient temperatures could exceed 30°C and some bees displayed signs of thermic stress. We thus developed an updated protocol to optimize faecal screening success (i.e., the probability that a bee will defecate), using two coolers (Fig. 2) to provide the bumble bees with a cool and dark environment. We first placed two icepacks in one of the coolers for at least 1 hour (Fig. 2A, B). We then started net-capturing bumble bees in the field. As soon as the first bumble bee was captured, we removed the two icepacks from the cooler and placed the petri dish with the bumble bee inside (Fig. 2C). Removing the icepacks is especially important; if bumble bees were placed on the icepacks, no faeces would be obtained due to a major decrease in metabolism. At that step, the icepacks were placed in the second cooler, subsequently used to store faecal samples upon collection. We checked the cooler for faces approximately every 10 min, or every time we placed a new petri dish in the cooler (Fig. 2C). As soon as we observed faeces in a petri dish, we placed the section of the petri dish with the faeces on top by gently turning the petri dish, since we could not release bees immediately to avoid re-capturing them. Once faecal collection was completed, all bees were released. A total of 298 bumble bees were captured in this study, on which we collected 256 faecal samples (85.9% success; 6 samples with unknown information). No bees died during this experiment. Faecal samples were frozen at -80°C within 24 hours after collection and subsequently analyzed within 1 hour after being defrosted using a Neubauer hemacytometer.

Study 4

Captures were conducted following the updated cooler protocol (Fig. 2) at 13 sites (one survey per site) around Toronto (Southern Ontario, Canada; 43° 39' 3.8520'' N and 79° 20' 49.2540'' W) from May 1st to June 27th, 2022. Ambient temperatures ranged from 10 to 26°C during sampling sessions, which lasted up to three hours. No bees died during this experiment. A total of 194 bumble bees were net captured, from which we collected 163 individual faecal samples (84.0% success). Samples were analyzed under the microscope on the day of collection using a Neubauer hemacytometer.

Identification of Crithidia and Vairimorpha parasites

Faecal samples were preserved on icepacks or in the fridge at + 4°C until being processed (see details for each study above). When processing a sample, we added 100 µL of distilled water into the petri dish with a micropipette and homogenized (stirring with the pipette). We then collected the faecal solution, placed it in a 0.5 ml Eppendorf tube on ice, and then shacked this sample with a Vortex mixer for 15 seconds. We immediately conducted faecal screening by taking 4 µL of the faecal solution and mounting it on a slide with a coverslip. We analyzed the sample under a compound microscope (phase-contrast, x40 magnification). We recorded the presence/absence of cells and spores of Crithidia and Vairimorpha, respectively, to assess parasite presence by looking at the entire sample mounted. Methods for measuring number of spores/cells varied between studies so it is not possible to compare intensity, whereas presence/absence was analyzed similarly between studies and can be used to validate that cells and spores of these parasites can be detected with this protocol.

Statistical analyses

This paper intended to validate a non-destructive protocol and assess the efficacy of the updated cooler protocol to collect adult bumble bee faeces in the field, increase the rate of faeces collection, and validate the presence of Crithidia and Vairimorpha in faecal samples. For those reasons, we did not perform analyses of protocol vs species or caste, or compare parasite loads or prevalence. Using a Generalized Linear Mixed Model (GLMM) with a binomial distribution, we investigated the effect of the protocol (standard versus cooler) on the faeces collection success (binary response). Sampling protocol and Julian date (accounting for ambient temperature) were included as fixed effects, while the site and the species were included as random effects. As data presented here were collected as part of four independent studies, compositions of species and castes vary between each study (Table S1). Including the species and sites as a random effect in our model allowed us to control for potential species or landscape effects on faecal collection success. In addition, we ran an alternative model looking at caste effect on faces collection success, to ensure that results of the model testing for the method were not driven by a different proportion of queens and workers in each study. This model showed no effect of caste on faces collection success (Table S2) and had a greater AIC than the model we present for testing the protocol (△AIC = 1.66). Analyses were conducted using SPSS software (IBM SPSS Statistics for WINDOWS v. 24.0. IBM Corp.). The significance threshold was set at α < 0.05. Data presented are means ± SE.

Success in faeces collection

We captured and released a total of 938 bumble bees, belonging to 14 species plus cuckoo bumble bees (subgenus Psithyrus) that were not identified to species (Table S1). Overall, faeces were collected for 737 of 932 bumble bees (79.1% success rate; information was missing for 6 bees).

Results of the GLMM revealed a significant effect of the protocol: the cooler protocol significantly increased the probability of collecting faeces compared to the standard protocol (F_1;938 = 4.41, t = 2.10, p = 0.036, Fig. 3). After accounting for potential species and site variations between each study, faeces collection success probability was 76% (0.76 ± 0.11, N = 496) with the standard protocol, and 86% (0.86 ± 0.07, N = 436) with the updated cooler protocol (as estimated by the GLMM; see Fig. 3). We found no effect of the Julian date on the probability of collecting faeces (F_1;938 = 0.04, t = 0.19, p = 0.85).

Identification of gut parasites

We were able to identify cells of Crithidia (Fig. 4) and spores of Vairimorpha (Fig. 5) in bumble bee faecal samples. The use of the phase contrast was especially important in observing Vairimorpha spores. The number of Crithidia cells ranged from 0 to 159 per sample, while the number of Vairimorpha spores ranged from 0 to 45 per sample.

We validated a non-destructive method commonly used in laboratory conditions to collect bumble bee faeces in the field, improved its efficiency in collecting faeces by 10% using a previously refrigerated cooler and identified cells of Crithidia and spores of Vairimorpha in faeces by microscopy.

Optimal protocol and comparison with other studies

Using petri dishes to collect bee faeces is a commonly used approach in laboratory settings (Chen et al., 2006; Gomez-Moracho et al., 2021). We adapted the protocol to field conditions and optimized faeces collection success by placing bumble bees in dark, cool, and low-sensory environment using a previously refrigerated cooler with no ice in it (Fig. 2). Faeces collection success was estimated at 76 ± 11% with the standard protocol by our model, and at 86 ± 7% with the cooler protocol. The updated cooler protocol thus increased our collection success by 10% and reduced variation compared to the standard protocol. Additionally, the low-cost and widespread availability of the equipment should promote its use in field studies and improve willingness to sample bees using non-destructive methods. As there are calls to reduce destructive sampling methods in bee research where possible (Miller et al., 2022; Montero-Castaño et al., 2022; Tronstad et al., 2022), using the updated cooler protocol will contribute to achieving this goal.

The number of Crithidia cells ranged from 0 to 159 in 0.02 µL of faeces extract diluted by 100 µL of water. Studies on wild bumble bees report ranges between 0 to 245 cells per 0.02 µL of macerated gut extract diluted by 300 µL (Giacomini et al., 2018), above what we recorded but of similar order of magnitude. Vairimorpha spore counts ranged between 0 and 45 spores per 0.02 µL diluted faecal sample, thus around 225,000 spores/µL of undiluted faecal sample, similar to what is usually recorded in wild bumble bees (Blaker et al., 2014). Using faeces to assess prevalence of Vairimorpha spores (i.e., the number of bee individuals infected) has been shown to be reliable in laboratory conditions, although faecal counts yielded more variation than PCR in infection intensity, i.e. number of spores recorded (Gomez-Moracho et al., 2021). However, this should not influence the use of the non-destructive faecal collection method if standardized within each study. If infection intensity is assessed by counting the cells and spores under the microscope, this should be performed using a Neubauer hemacytometer (Giacomini et al., 2018; Gomez-Moracho et al., 2021), to standardize data and allow between-studies comparisons.

Limitations and perspectives

Our studies did not test for species differences in defecation probability or parasite loads, which could be the focus of future work. However, we accounted for species variability in our models, and even though our samples were dominated by common species (e.g., B. impatiens and B. ternarius), we also tested the updated cooler protocol on few specimens of less common or even endangered species (e.g., B. fervidus and B. terricola, with success rates of 67% and 83.3%, respectively).

In addition, since this non-destructive method screens for two gut parasites commonly found in bumble bees, Crithidia and Vairimorpha (Graystock et al., 2020; Grupe & Quandt, 2020), this protocol cannot be used to look at other pathogens that infect different parts of the body (e.g. body fat, brain or the reproductive system) (Figueroa et al., 2023). The protocol solely allows users to assess pathogens that are visible under the microscope, which excludes non-sporulating Vairimorpha and viruses (Blaker et al., 2014; McNeil et al., 2020), that are also transmitted through an oral-faecal route in Hymenoptera (Yañez et al., 2020). However, these last two issues could be overcome by conducting molecular analyses (e.g., RT-PCR or qPCR; see (Babin et al., 2022)) on faecal samples. This has already been done in laboratory settings to assess the prevalence of important viruses found in bees (Chen et al., 2006). In their study, Chen et al. (2006) highlighted that faecal samples collected using petri dishes were amongst the best tissues to assess the prevalence of Black Queen Cell Virus (BQCV) and Deformed Wing Virus (DWV) by RT-PCR in the honey bee, but were less effective for Chronic Bee Paralysis Virus (CBPV) and Sacbrood Bee Virus (SBV) (Chen et al., 2006). Both BQCV and DWV are common and important threats to bumble bees (Burnham et al., 2021; McNeil et al., 2020; Tsvetkov et al., 2021); thus, assessing their prevalence using a non-destructive faecal screening approach could be of particular importance, especially for endangered species such as B. terricola or B. affinis. If the non-lethal faecal sampling approach were used in molecular analyses, the protocol would have to be adjusted to ensure the proper sample collection for these tools. This includes using double-distilled nuclease free water, keeping samples on ice and freezing them (at − 20°C for analyses on DNA and at − 80°C for analyses on RNA) within hours of collection and until being processed (Gomez-Moracho et al., 2021; McNeil et al., 2020; Tsvetkov et al., 2021).

The protocol described in this study was developed for wild bumble bees, and it remains to be confirmed whether it can be adapted to other species of wild bees, especially smaller solitary bees. Nonetheless, a recent study on Osmia bicornis used a similar faecal screening approach in laboratory conditions (Tian et al., 2018), suggesting that the non-destructive approach presented here could be applied to some smaller bee species, which have been identified as hosts of Crithidia (Figueroa et al., 2021). Considering the growing interest and need to assess the health social and solitary wild bees, especially endangered or declining species that are more exposed and impacted by diseases than common and non-endangered species (Averill et al., 2021; Cameron et al., 2011; Gillespie, 2010), the development of non-destructive sampling methods is crucial. The protocol presented here is a step that should help achieving sustainable sampling in bee research, by reducing potential damages of repeated sampling on wild bees when assessing their health.

Acknowledgements

We thank Aurélie Demers and Frédéric McCune for their help in collecting part of the data. Thanks to Pierre-Olivier Montiglio for providing access to his laboratory in 2020 during COVID lockdown, which ensured analyses of collected faeces in study 1. Many thanks to agricultural producers who allowed us to capture bumble bees on their site and to M.P. Beaudoin for her help in implementing study 2.

Author Contributions: M.L.T and V.F. obtained funding, with contribution from L.A. and G.L. for study 1, from S.M., L.A., P.B., C.C. and S.C. for study 3 and from JS.M. and S.M. for study 4. M.L.T. conceived the theoretical framework and design of these studies, with feedback from other authors. Data collection and curation was conducted by M.L.T. and C.B., with help from S.M. Finally, M.L.T conducted statistical analyses and wrote the first draft of the manuscript with edits from all authors.

Funding: This work was funded by the “Ministère de l'Agriculture, des Pêcheries et de l'Alimentation” (MAPAQ, project Prime-Vert #6938216) and the Liber Ero program. In 2020, M.L.T. salary was covered by P.B. NSERC grant (#2016-04193) and the MAPAQ (Prime-Vert, #6667567), while here was covered by the Liber Ero program in 2021 and 2022. Funding for C.B. was provided by a Mitacs BSI (IT30033) awarded to JS.M and S.MK.

Competing interests: Authors declare no competing interests;

Data and materials availability: All data will be made available on OSF.

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No competing interests reported.

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A non-destructive method to detect pathogens in bumble bees for conservation research

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Abstract

Figures

Introduction

Materials And Methods

Experimental design: bee captures and faeces collection

Studies and sites

a) Standard protocol

Study 1

Study 2

Study 3

Study 4

Identification of Crithidia and Vairimorpha parasites

Statistical analyses

Results

Success in faeces collection

Identification of gut parasites

Discussion

Optimal protocol and comparison with other studies

Limitations and perspectives

Declarations

References

Additional Declarations

Supplementary Files

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