Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Summary

Microbial consortia within bumble bee hives enrich and preserve pollen for bee larvae. Using next generation sequencing, along with laboratory and field-based experiments, this manuscript describes protocols used to test the hypothesis that fungicide residues alter the pollen microbiome, and colony demographics, ultimately leading to colony loss.

Abstract

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered "bee-safe," there is mounting evidence that fungicide residues in pollen are associated with bee declines (for both honey and bumble bee species). While the mechanisms remain relatively unknown, researchers have speculated that bee-microbe symbioses are involved. Microbes play a pivotal role in the preservation and/or processing of pollen, which serves as nutrition for larval bees. By altering the microbial community, it is likely that fungicides disrupt these microbe-mediated services, and thereby compromise bee health. This manuscript describes the protocols used to investigate the indirect mechanism(s) by which fungicides may be causing colony decline. Cage experiments exposing bees to fungicide-treated flowers have already provided the first evidence that fungicides cause profound colony losses in a native bumble bee (Bombus impatiens). Using field-relevant doses of fungicides, a series of experiments have been developed to provide a finer description of microbial community dynamics of fungicide-exposed pollen. Shifts in the structural composition of fungal and bacterial assemblages within the pollen microbiome are investigated by next-generation sequencing and metagenomic analysis. Experiments developed herein have been designed to provide a mechanistic understanding of how fungicides affect the microbiome of pollen-provisions. Ultimately, these findings should shed light on the indirect pathway through which fungicides may be causing colony declines.

Introduction

Managed and wild bee species are experiencing widespread declines, with major implications for both natural and agricultural systems¹. Despite concerted efforts to understand the causes of this problem, the factors driving honey bee declines are still not well understood^2,3,4. For certain species of wild, native bees, the situation has become dire^5,6. If bee populations cannot be sustained when they intersect with industrial agriculture, their populations will continue to fall, and the crops requiring pollinators (35% of worldwide production⁷) will endure reduced harvests.

While many potential factors such as pesticide exposure, disease, and habitat loss^1,4,8,9,10 have been implicated in honey bee decline, relatively little is known about the interactive effect of these stressors on native bees health, within or near agricultural systems. Many current research efforts continue to focus on insecticides, (e.g., neonicotinoids^11,12), although past research indicates that fungicides may also play a role in bee decline by impairing memory formation, olfactory reception¹³, nest recognition¹⁴, enzyme activity and metabolic functions^15,16,17. Globally, fungicides continue to be applied to flowering crops during bloom. Recent studies have documented that bees commonly bring fungicide residues back to the hive¹⁸, indeed, studies have shown a large proportion of tested hives contained fungicide residues^19,20. Further work has revealed that fungicide residue is associated with high rates of honey bee larval mortality^21,22,23 and the presence of "entombed pollen" within colonies, which although non-toxic, is devoid of microbial activity and is nutritionally compromised²⁴. Despite the fact that fungicides have long been considered "bee-safe," there is now evidence that exposure to fungicide alone can cause severe colony losses in a native bumble bee species, Bombus impatiens²⁵.

To establish causality between fungicide exposure and colony mortality, the modus operandi of these chemicals need to be determined. As evidenced in soils²⁶, sediments²⁷, and aquatic environments²⁸, by targeting fungi, fungicides most likely alter fungal abundance and diversity within pollen-provisions, thereby invoking a major community shift that may strongly favor bacteria. Without fungal competitors or antagonists, certain pathogenic bacteria can proliferate relatively unchecked, facilitating the spoilage of pollen-provisions. Past research has demonstrated that microorganisms, particularly yeasts and filamentous fungi, serve as nutritional symbionts for bees^29,30,31, protect against parasites and pathogens^32,33, and provide long-term preservation of pollen stores. Fungicides, therefore, may indirectly harm immature bees by disrupting the microbial community that is needed to provide these services and/or by increasing susceptibility to opportunistic pathogens and parasites¹². With increasing demands on food production, crops worldwide are being sprayed each year with fungicides during bloom, underscoring the need to understand the magnitude of such fungicide-induced effects.

To-date, the primary knowledge gaps relating to native bee microbial ecology can be represented by the following questions: To what extent does fungicide change the microbial community within bee pollen-provisions? What are the downstream impacts of consuming pollen with a profoundly altered microbial community? In keeping with these ecologically germane questions, experiments were developed with the primary goals of revealing 1) that fungicide residue alone can cause severe colony decline in a native bee species; 2) the degree to which microbial communities in pollen-provisions are altered by fungicides, and 3) how bee health is affected by a severely altered microbial community. The experimental objectives were defined to address the above questions using a combination of laboratory- and field-based experiments. Using state-of-the-art metagenomic and molecular techniques alongside traditional methods of field observation, this research aims to piece together the potential effects of fungicides on bee health.

The first objective of this study is to demonstrate that fungicide exposure alone can cause significant colony losses among native bee species. A study involving large field cages was used to investigate the effects of fungicide exposure on the colony growth of Bombus impatiens, a ubiquitous, abundant native bee in the US (Figure 1, Figure 2, Figure 3). It was hypothesized that fungicide-treated hives would present lower fitness, and atypical demography compared to non-exposed hives. Data obtained from this experiment supported this hypothesis, demonstrating that fungicide residues within pollen can be the sole cause of profound colony losses in a native bumble bee species²⁵. The second objective of this study is to investigate the response of the pollen microbiome to fungicide exposure. It is hypothesized that the community composition of microbes within pollen-provisions exposed to fungicides will be different from that of untreated pollen. While fungal abundance and diversity are expected to decline significantly, bacteria and/or a single dominant fungal species will likely grow unchecked in the absence of other competing fungi. Through a series of in-vivo trials, these shifts in microbial community composition will be analyzed using metagenomics.

Protocol

1. Examine the Effect of Fungicide Exposure on Bumble Bee Colony Success Using Field Cage Experiments

1. Set up ten mesh cages in a field planted with oats. Dig a trench around each cage, and dig all four edges of the mesh cage into the ground to ensure that bees cannot escape. Stock the cages with potted, flowering plants that are known to be attractive to bees (e.g. buckwheat, borage, alyssum, cosmos, and sunflowers) (Figure 2).
2. Supplement the cages with a single tray (36 cm x 42 cm) of in-bloom clover. Cluster floral resources within one corner of the cage, occupying a space approximately 2.5 m x 1 m. Vegetate the remaining cage area by the oats.
3. Randomly assign the bumble bee colonies, each containing workers and a single queen, a treatment (fungicide present/absent, N = 5 colonies per treatment, 10 colonies total), then place them within a field cage (N = 1/cage) for 29 days (23 June - 21 July 2014).
4. Orient the colony boxes such that the colony's openings point to the south to provide the bees with optimal navigational conditions. Subsidize the colonies with sugar water bladders, placed inside the hive boxes to supplement nectar availability.
5. Apply chlorothalonil-based fungicide at a field-relevant level (20 g/L) to flowering plants in the five fungicide treatment cages, using a hand held pesticide sprayer, twice during the study (day 0 and 13). Coat the flowers uniformly such that no further liquid adheres to floral surfaces (Figure 3).
6. At the conclusion of the field cage study, remove the B. impatiens colonies from the cages by hand, cool the hives by placing in -20 °C freezer for 20 min.
7. Remove bees using sterile forceps and record the number of larvae, pupae, adult females (i.e. foragers), and adult males. Using an analytical balance record the dry-weight of the mother queen, larvae, pupae, adult females (i.e. foragers), and adult males.

2. Examine the Effects of Fungicide Exposure on Microbial Communities in the Pollen-provisions of Bumble Bee Nests Using Laboratory Based In Vivo Trials

1. Pulverize commercially purchased pollen to a fine powder using a standard laboratory ball-mill. Sterilize powdered pollen by soaking in 70% ethanol, allowing it to evaporate overnight under UV light. Verify sterility of pollen by plating ~0.5 mg on general-purpose agar media.
   1. To the dry, sterilized pollen, using sterilized pipettes, add field relevant dosage of fungicide: propiconazole at 14.3%; azoxystrobin at 22.9% for treatments (0.74 µL and 0.65 µL respectively/ day/ hive). Mix well using sterilized wooden sticks.
2. Place 6 experimental hives (n = 3 each for control and treatment) in a clean, hygienic laboratory benchtop maintained at room temperature. Each day weigh 4.27 g of pollen^34,35 mixed with fungicides (for treatments) or sterile water (for control) inside a hood using standard aseptic technique.
   1. Using the trap doors provided by the side of the cardboard box enclosing the hives, introduce the pollen inside the hives. Supplement the hives with sterilized sugar solution each week. Continue feeding regime for four weeks.
3. At the conclusion of the lab-based study, cool the hives by placing in -20 °C freezer for 20 min. Scrape out the pollen-provisions contained within brood chambers using sterilized forceps and spatulas and place in sterile storage tubes. Store at -80 °C. Count and record the weight of the workers and queen mother at the start and end of the experiment (step 1.7).
4. Isolate DNA from pollen-provision sample using commercially available DNA isolation kits (see Materials Table for details).
   1. Add 0.25 g of pollen-provision to extraction tubes, briefly vortex to mix.
   2. Make a 200 mg/mL lysozyme solution in deionized distilled water, enough for 50 µL/sample. Shake vigorously to completely form solution.
   3. Add 50 µL of the lysozyme solution to the extraction tubes with sample in it, and mix well by inverting several times. Incubate the tube for 10 min at 37 °C in a water bath. If precipitate has formed, heat solution to 60 °C until dissolved before use.
   4. Add 70 µL of aqueous lysis solution to extraction tubes, secure horizontally on a flatbed vortex pad with tape, and vortex for 10 min. Centrifuge tubes at 10,000 x g for 30 s at room temperature.
   5. Transfer the supernatant to a clean 2 mL collection tube. Add 250 µL of protein precipitation solution, and vortex for 5 s. Incubate at 4 °C for 5 min in an ice bath.
      NOTE: Expect between 400 µL to 500 µL of supernatant. Supernatant may still contain some particles.
   6. Centrifuge the tubes at room temperature for 1 min at 10,000 x g, and transfer up to 600 µL of supernatant to a clean 2 mL collection tube. Add 200 µL of aqueous inhibitor removal solution, vortex briefly, and incubate at 4 °C for 5 min. Centrifuge the tubes at room temperature for 1 min at 10,000 x g. Transfer up to 750 µL of supernatant into a clean 2 mL collection tube.
   7. Add 1200 µL of aqueous bind solution to the supernatant, and vortex for 5 s. Load approximately 675 µL of the supernatant onto a spin filter, and centrifuge at 10,000 x g for 1 min at room temperature. Discard the flow through.
   8. Repeat 2.4.7. twice.
      NOTE: A total of three loads for each sample processed are required.
   9. Add 500 µL ethanol, and centrifuge at room temperature for 30 s at 10,000 x g. Discard the flow through, and centrifuge again at room temperature for 1 min at 10,000 x g. Place spin filter in a clean 2 mL collection tube.
   10. Add 100 µL of elution buffer to the center of the filter membrane. Centrifuge at room temperature for 30 s at 10,000 x g. Discard the spin filter. Store collected DNA between -20 °C and -80 °C
5. Use isolated DNA for sequencing.
   1. Quantify, and normalize isolated DNA to 2 ng/µL by fluorometric analysis.
      1. Prepare reactions in triplicate for each extracted sample to compare the relative amounts of 28S (plant), analyze ITS (fungal), and 16S (bacterial) components of each pollen-provision sample³⁶. Ensure that each reaction contains 10 ng of total DNA, 2x of asymmetrical cyanine dye based master mix, and 2.5 µL each of forward and reverse primers pairs 28KJ/28B³⁷ for plant and ITS1/ITS5.8R for fungal DNA³⁸.
   2. Amplify DNA using the following parameters: 2 min pre-denaturation at 50 °C, 2 min initial denaturation at 95 °C, 40 cycles of (15 s at 98 °C, 15 s at 58 °C, 60 s at 72 °C), followed by a melt curve.
   3. Prepare a 2-step, nested PCR protocol using next-generation sequencing libraries targeting the 16S rRNA V3/V4 variable region and ITS/5.8s rRNA spacer region.
      1. Add 12.5 µL of DNA to 5 pmol of each primer and 2x master mix. Make separate reactions for each region (16S or ITS; see Table 1). Modify region specific primers as previously described^39,40 to add sequencer-specific adapter overhang nucleotide sequences to the gene-specific sequences.
      2. Perform initial amplification using the following parameters: 3 min initial denaturation at 95 °C, 25 cycles of (30 s at 95 °C, 30 s at 55 °C, 30 s at 72 °C), 5 min final extension at 72 °C.
      3. Following initial amplification, verify library size, and quantity by electrophoretic mobility, and clean using a 1x volume solid phase reversible immobilization beads to remove residual primers and reaction reagents. Pool 16S and ITS amplicons quantitatively to create a single amplicon pool for each sample.
      4. Add sequencer specific adapters and sample specific dual indexes using the following primers (see Table 1). Add 2.5 µL of amplified DNA to 5 pmol of each primer and 2x master mix. Perform library amplification using the following parameters: 3 min initial denaturation at 95 °C, 8 cycles of (30 s at 95 °C, 30 s at 55 °C, 30 s at 72 °C), 5 min final extension at 72 °C.
   4. Following PCR, clean finished libraries using a 1 x volume of solid phase reversible immobilization beads. Assess quality and quantity of the finished libraries using electrophoretic mobility, and fluorometry, respectively. Standardize libraries to 2 µM and pool prior to sequencing.
   5. Perform next-generation sequencing on a platform analogous to the adapters added in the secondary PCR, with an appropriate length to cover the entire amplicon⁴¹.
6. Sequence annotation and microbial composition analysis.
   1. Combine pair-end sequencing data (R1 and R2) into single contigs for all sequenced libraries. After merging both R1 and R2 files, a single fasta file for each of the libraries is produced.
      NOTE: This step and the processes described below (unless otherwise indicated) are performed in Mothur version 1.38.0³⁸.
   2. Screen each fasta file to remove ambiguous bases, unexpected long sequences, and long homopolymers.
      NOTE: The parameters for screening, and sequence removal are maxambig = 0, maxlength = 600, and maxhomop = 8 for both genes. Remove identical sequences, but keep a contingence table separately for all the libraries (a.k.a. count table in Mothur).
   3. Align unique sequences of the gene 16S library against the SILVA database version 123, and the gene ITS library against the UNITE ITS database⁴².
      NOTE: Sequences must be annotated from the kingdom level to the genus level.
      1. Subsequently, cluster aligned sequences with the function pre.cluster using diff=5, and remove chimeras.
   4. Classify sequences using the method Wang⁴³ based on the SILVA (for gene 16S) and UNITE ITS (for gene ITS) taxonomy files (cutoff value of 80%).
   5. Load into R⁴⁴ the contingence tables (one per gene) generated during the classification process in Mothur. At each taxonomic level, obtain relative abundance per library for further analysis of microbial community changes.
      NOTE: At each taxonomic level, merge together taxonomic groups when relative abundance is less than 2% for all the experimental repetitions.

Results

Field cage study:

Data obtained from the cage experiments showed that the bumble bee colonies had a significant response to fungicide exposure. The fungicide-treated hives produced significantly fewer workers (12.2 ± 3.8, mean ± SE) than the control hives (43.2 ± 11.2, F_1,9= 6.8, p = 0.03) (Figure 4). Additionally, the bee biomass of the fungicide-tr...

Discussion

Investigations into the effects of fungicides on bee health have remained an understudied aspect of pest management strategies. Our study aims to bridge this knowledge gap by using a suite of complementary techniques that explicitly isolate the potential factors driving bee declines. The planning, rationale, and rendering of these experiments are detailed below.

It is important to ensure that no bees are allowed to escape the mesh of the cage experiments, since this would compromise demography...

Materials

Name	Company	Catalog Number	Comments
Natupol Beehive	Koppert	USRESM1	16 hives
Propiconazole 14.3	Quali-Ppro	60207-90-1	Propiconazole 14.3%
Abound	Syngenta	4033540	Azoxystrobin 22.9%
Chlorothalonil	Syngenta	3452	Fungicide used for trials
Pollen granules	Bee rescued	B004D5650C	3X 16oz bottles, pollen for trials
Bacterial strains for inoculation	Currie Lab
Yeast strains for inoculation	Hittinger lab
Primer pairs	UW Biotech Center
DNA Isolation Kit	Mo Bio	12830-50	Commercial DNA isolation kit
Qubit dsDNA HS Assay Kit	Thermo Fisher	Q32851	DNA quantification tool
Select Master Mix for CFX	Thermo Fisher	4472952	Used to perform real-time PCR using SYBR GreenER dye.
Real-Time PCR Detection System	Bio Rad	1855196	Instrument used for PCR amplification
PCR Clean-Up Kit,	Axygen	10159-696	Used for efficient removal of unincorporated dNTPs, salts and enzymes
DNA 1000 Kit	Agilent	5067-1504	Used for sizing and analysis of DNA fragments
MiSeq Sequencer	Illumina		Used for next-generation sequencing
Assorted glassware (beaker, flasks, pipettes, test tubes, repietters)	VWR