Name: collection and identification of pollen from honey bee colonies
Uploaded: 2021-01-19T14:00:00.000+00:00
Duration: 8 min 11 s
Description: Watch this Scientific Journal Video about Collection and Identification of Pollen from Honey Bee Colonies at JoVE.com

We thank Dr. Gretchen Jones (USDA-ARS, APMRU, College Station, TX) for assisting with color sorting and acetolysis analysis. This research was supported by research funds provided to R.R.S. by the Oregon State Beekeepers Association.

1. Collecting corbicular pollen from honey bee colonies using pollen traps
<ol>
	<li>Determine when to trap pollen from the desired apiary location. 
	&#8203;NOTE: Ideal climatic conditions include full sun exposure, low wind speeds, low humidity, and no forecasted precipitation during the desired period for pollen collection.</li>
	<li>Select optimal honey bee colonies for trapping pollen within the apiary location.
	<ol>
		<li>Assess colony strength by counting foragers returning to the colony entrance for 2 min. Select colonies with the highest total number of returning foragers.</li>
		<li>Select hives with woodenware that is in good condition, preferably without extra entrances and warped lids. Use colonies with fewer alternative entrances as they have increased likelihood of returning foragers reorienting to the trap entrance.</li>
		<li>Select south-facing hive entrances whenever possible. If hives are palletized, install pollen traps on every colony facing the same direction on a given pallet to avoid the drift of foragers into neighboring hive entrances.</li>
		<li>If desired, assess the brood nest of the colony by inspecting frames for the presence of larvae. Select colonies with relatively large amounts of larvae.</li>
	</ol>
	</li>
	<li>Install pollen traps on the selected honey bee colonies. 
	&#8203;NOTE: Installation will differ based on the type of pollen trap. Types include a) front-mount, b) bottom-mount, c) top-mount, or d) auger-hole entrance mount. See the discussion section for details.
	<ol>
		<li>For front-mount traps, attach the trap in front of the entrance with staples, screws, and tape, or connect the trap to bungee cords wrapped around the hive. For bottom-mount traps, place the trap underneath the lowest hive box, and fix the trap entrance near the original entrance. For auger-hole mount traps, attach the trap directly in front of an auger hole of a hive box using staples, screws, and tape. For top-mount traps, place the trap above the uppermost hive box and below the lid.</li>
	</ol>
	</li>
	<li>Seal all other possible entrances into the colony by using non-adhesive and moldable material, such as latex or polyurethane foam, or #8 hardware cloth (2.7 mm aperture) for auger holes. Use adhesive tape for small cracks.</li>
	<li>If using front-mount traps, place a barrier, such as a rubber mat, between the collection basket and grass to avoid moisture damage from dew.</li>
	<li>Engage the trapping mechanism of the pollen trap 24 h after installation and before the day&#39;s foraging flight commences (late evening/early morning). 
	&#8203;NOTE: This step is ideal, but not necessary. Engage pollen traps every other week if trapping pollen on the same colonies throughout a given period.</li>
	<li>Collect corbicular pollen from the collection tray, place it into plastic bags or centrifuge tubes, and store in a cooler with ice.
	<ol>
		<li>To assess the diversity and abundance of pollen species, e.g., landscape-level nutrition studies, collect pollen in two or three 72 h intervals40.</li>
		<li>For pesticide residue analysis, collect pollen in 24 h to 96 h intervals with a minimum of 3 g for processing41.</li>
	</ol>
	</li>
	<li>Clean the pollen by removing bee parts and other hive debris. 
	NOTE: Use disposable gloves when handling pollen samples, and change the disposable gloves between samples. Use separate tools to remove debris from the pollen collected in each trap. Rinse and dry before using the tools for another batch of trapped pollen.</li>
	<li>Store pollen at -20 &#176;C or below to maintain its compositional integrity if pollen is intended for pollen source identification, quantity assessment, or pesticide residue analyses41,42.</li>
	<li>After removing the traps from the hives, sterilize all equipment in a 5% bleach solution, rinse, and dry the equipment before the next use.</li>
</ol>
2. Pollen pellet color sorting for downstream pollen source identification and quantity assessment
<ol>
	<li>Ensure that there are at least 20 g of pollen sample to work with. Thoroughly mix the pollen sample in its bag or another suitably sized container to obtain a homogeneous mix of all pellets contained therein. To avoid unintentional bias in the next step, obscure the sample&#39;s color composition from view before removing a subsample from the sample bag.</li>
	<li>Using a scooper or large spoon, scoop out 10 g of pollen as a representative subsample of the whole. Slowly pour pellets out of the scooper onto the balance until the display reads 10 g. If the first scoop was not big enough, retrieve another scoop from the sample in the same way. 
	NOTE: These specified weight requirements (20 g and 10 g) serve as examples only. Researchers should adjust the quantity of pollen used in each step as appropriate for specific needs.</li>
	<li>Remove all bee parts and other debris from the 10 g subsample. Then, if necessary, add a little more pollen from the original sample to achieve a total weight of 10 g of the subsample.</li>
	<li>Sort each pollen pellet from the 10 g subsample into a color group. Use both pollen color and texture to differentiate between color groups. 
	NOTE: Some variation within a group is expected, but using the Pantone color guide during sorting can increase consistency.</li>
	<li>To ensure at least 0.25 g of each color group for downstream steps, place any pellets that are not plentiful enough to form a color group of at least 0.25 g into a miscellaneous group. Name each individual color group using the Pantone color guide. Label the miscellaneous group misc.</li>
	<li>Weigh each color group on a separate weigh paper, and/or count the number of pellets in each color group. Record the color group names and weights or counts on a datasheet. 
	NOTE: Choosing whether to weigh or count the number of pellets in each color group depends on the researcher&#39;s metric of interest and project goals.</li>
	<li>Create a microcentrifuge tube label for each color group using a solvent-resistant pen and adhesive paper tube labels. Include the current date, sample identifier, sample collection date, and color group number in the label. Apply the labels to clean, dry 2 mL microcentrifuge tubes.</li>
	<li>Weigh out 0.25 g (&#177; 0.05 g) of pollen pellets from each color group, and place this amount in the appropriately labeled microcentrifuge tube. 
	NOTE: If there is slight variation in color or texture in the pollen of a given color group, ensure that there is a representative sample of pellets within each tube. The reagent volumes and incubation and centrifugation times that follow are appropriate for 0.25 g of pollen. Therefore, use this quantity of pollen in the microcentrifuge tubes to be used in acetolysis. This protocol should provide ample, stained pollen for downstream plant source identification by light microscopy. If using a different quantity of pollen in acetolysis, the specifics of reagent volume and processing times should be adjusted accordingly.</li>
	<li>Place the remaining pollen from each color group into individual plastic bags (one bag per color) labeled with the color group name. Store these bags with the other parts of the appropriate original sample in -20 &#176;C storage.</li>
	<li>Thoroughly mix the pollen in the tube with a clean wooden toothpick for 10 to 15 s.</li>
</ol>
3. Preparation for acetolysis
<ol>
	<li>Before beginning any part of acetolysis for the first time, contact the designated institution&#39;s environmental health and safety (EHS) department for instructions on how acetolysis-related reagents and waste should be handled.</li>
	<li>Obtain stock solutions of the following reagents, and place them in the fume hood in accordance with EHS guidelines for chemical storage: 95% ethanol; distilled water; glacial acetic acid, anhydrous; concentrated sulfuric acid; glycerin; and clear nail polish.</li>
	<li>Prepare stock solutions of the following reagents, and place them in the fume hood in accordance with EHS guidelines for chemical storage: saturated sodium bicarbonate (8% w/v solution in distilled water); and safranin O (1% w/v solution in 50% ethanol).</li>
</ol>
4. Acetolysis
<ol>
	<li>Perform the pre-acetolysis procedure of glacial acetic acid wash. Carry out the following steps within the fume hood with lab coat, eye protection, and nitrile gloves.
	<ol>
		<li>Turn a heat block on to 80 &#176;C. 
		NOTE: Ensure that a squeeze bottle of saturated sodium bicarbonate is easily accessible. This can be used to neutralize acid spills in the fume hood if they occur.</li>
		<li>Label one glass beaker for acid waste, one for ethanol waste, and one for acetolysis mixture.</li>
		<li>Using previously prepared stock solutions, create working aliquots of the following reagents in labeled, appropriately sized glass beakers: ~23.0 mL of glacial acetic acid; ~33.0 mL of distilled water; ~23.0 mL of 95% ethanol; ~25.0 mL of sodium bicarbonate (for acid-contaminated solid waste). 
		&#8203;NOTE: These are the volumes required to complete the following acetolysis procedures on a total of 10 color group samples (10 microcentrifuge tubes).</li>
		<li>Slowly add 500 &#181;L of glacial acetic acid to each microcentrifuge tube containing 0.25 g of color group pollen. While visually inspecting the tube, stir the pollen with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. Place the used toothpick in the sodium bicarbonate waste beaker after use; repeat this process for each tube. 
		&#8203;NOTE: Use a clean, new toothpick for each tube. Ensure that each tube&#39;s lid is tightly closed.</li>
		<li>Centrifuge the samples for 3 min at 1,100 &#215; g. Decant the supernatant from the tubes into the acid waste beaker. Then, softly and briefly touch the open mouth of the tube with a clean paper towel to remove residual glacial acetic acid around the rim of the tube. 
		NOTE: Be careful not to lose the pollen pellet when decanting supernatants.</li>
	</ol>
	</li>
	<li>Perform the acetolysis procedure. 
	NOTE: Carry out the following steps within the fume hood with lab coat, eye protection, and butyl vinyl gloves.
	<ol>
		<li>Prepare the acetolysis mixture (9:1 glacial acetic acid:sulfuric acid) by first adding 10.8 mL of glacial acetic acid (from the working aliquot) to the beaker labeled acetolysis mixture. Then, using a 1000 &#181;L pipette fitted with 1250 &#181;L filtered pipette tips, slowly add 1200 &#181;L (1.2 mL) of concentrated sulfuric acid from the stock solution to the acetolysis mixture beaker containing glacial acetic acid. Discard the used pipette tip into the beaker of sodium bicarbonate. 
		NOTE: The acetolysis mixture beaker may become warm to the touch, and the mixture may turn yellow. There are two possibilities that cause the mixture to turn a dark color: (a) the reagents may be past their expiration dates, or (b) too much sulfuric acid may have been added. In any case, if the mixture turns dark, discard it in the acid waste beaker, and prepare a fresh acetolysis mixture.</li>
		<li>Gently stir the acetolysis mixture with a glass rod or wooden stir stick to ensure it is homogenized. Place the used rod/stick in the beaker of sodium bicarbonate.</li>
		<li>Using a 1000 &#181;L pipette with 1250 &#181;L filtered pipette tips, slowly add 1000 &#181;L of acetolysis mixture from the beaker to each tube. While visually inspecting the tube, stir with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. Place the used toothpick in the sodium bicarbonate waste beaker after use. 
		&#8203;NOTE: Use a clean, new toothpick for each sample.</li>
		<li>Place the samples on the preheated (80 &#176;C) heat block. Incubate the tubes for 5 min, stirring each tube thoroughly with a clean toothpick halfway through incubation. Place each used toothpick in the beaker of sodium bicarbonate after use. 
		NOTE: Do not leave toothpicks in the samples; the acid will dissolve them.</li>
	</ol>
	</li>
	<li>Perform the post-acetolysis glacial acetic acid wash procedure. 
	&#8203;NOTE: Carry out the following steps in the fume hood with lab coat, eye protection, and butyl vinyl gloves.
	<ol>
		<li>Slowly add 500 &#181;L of glacial acetic acid to each tube. While visually inspecting the tube, stir with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. Place the used toothpick in the beaker of sodium bicarbonate after use. 
		&#8203;NOTE: Use a clean, new toothpick for each sample. Ensure that each tube&#39;s lid is tightly closed.</li>
		<li>Centrifuge the samples for 3 min at 1,100 &#215; g. Decant the supernatant from each tube into the acid waste beaker. Then, softly and briefly touch the open mouth of the tube with a clean paper towel to remove residual acid around the rim of the tube.</li>
		<li>Thoroughly rinse the butyl vinyl gloves under running water for at least 30 s, remove them, and set them to dry. 
		NOTE: Follow the manufacturer&#39;s guidelines on reuse of butyl vinyl gloves.</li>
	</ol>
	</li>
	<li>Perform three water rinses for each sample. Carry out the following steps within the fume hood with lab coat, eye protection, and nitrile gloves.
	<ol>
		<li>Add 1000 &#181;L of distilled water from the distilled water beaker to each tube. While visually inspecting the tube, stir with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. Place the toothpick in the sodium bicarbonate waste beaker after use. 
		&#8203;NOTE: Use a clean, new toothpick for each sample. Ensure that each tube&#39;s lid is tightly closed.</li>
		<li>Centrifuge the samples for 3 min at 1,100 &#215; g. Decant the supernatant from the tubes into the beaker of sodium bicarbonate. Then, softly touch the open mouth of the tube with a clean paper towel to remove residual water around the rim of the tube.</li>
		<li>Repeat steps 4.4.1-4.4.2 two additional times for a total of three water rinses.</li>
	</ol>
	</li>
	<li>Perform the ethanol rinse for each sample. 
	&#8203;NOTE: Carry out the following steps within the fume hood with lab coat, eye protection, and nitrile gloves.
	<ol>
		<li>Using a 1000 &#181;L pipette with 1250 &#181;L filtered pipette tips, add 1000 &#181;L of 95% ethanol from the ethanol beaker to each tube. Discard the pipette tip into non-hazardous waste. While visually inspecting the tube, stir with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. 
		&#8203;NOTE: Place the toothpick in the sodium bicarbonate waste beaker after use. Use a clean, new toothpick for each sample. Ensure that each tube&#39;s lid is tightly closed.</li>
		<li>Centrifuge the samples for 3 min at 1,100 &#215; g. Decant the supernatant from the tubes into the ethanol waste beaker, and softly touch the open mouth of the tube with a clean paper towel to remove residual ethanol from the tube.</li>
	</ol>
	</li>
	<li>Wear lab coat, eye protection, and nitrile gloves to stain samples. Mix the Safranin O stain stock solution using gentle inversion.
	<ol>
		<li>Using a disposable, plastic transfer pipette, add 5-10 drops of Safranin O stain to each tube. While visually inspecting the tube, stir with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. Leave the toothpick in the tube.</li>
		<li>Using a 1000 &#181;L pipette with 1250 &#181;L filtered pipette tips, add 1000 &#181;L of 95% ethanol from the ethanol beaker to each tube. Discard the pipette tip into non-hazardous waste. While visually inspecting the tube, stir with the toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. Place the used toothpick in the non-hazardous waste after use.</li>
		<li>Ensure that each tube&#39;s lid is tightly closed. Centrifuge for 3 min at 1,100 &#215; g. Decant the supernatant into the ethanol waste beaker. 
		&#8203;NOTE: Do not touch the mouth of the tube with a paper towel this time.</li>
		<li>Add 10-15 drops of glycerin to each tube using a plastic disposable transfer pipette. While visually inspecting the tube, stir the contents of the tube with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. 
		NOTE: Place the used toothpick in the non-hazardous waste after use. Use a clean, new toothpick for each sample. Ensure that all tube labels are legible.</li>
	</ol>
	</li>
	<li>Leave the tubes open in the fume hood to evaporate the ethanol for at least 2 h at ambient room temperature. Check the samples for ethanol odor: if it is detectable, the samples are not ready and should be left to dry until the ethanol odor dissipates.</li>
	<li>Clean all materials, and dispose of waste. Switch off both the centrifuge and heat block. Dispose of all solid and liquid waste in accordance with the designated institution&#39;s environmental health and safety guidelines.</li>
	<li>Prepare microscope slides for pollen identification; label them legibly. Label a clean glass microscope slide appropriately for each color group/sample that will be mounted. While visually inspecting the tube, stir the sample with a clean toothpick for 10-15 s, and ensure that the contents of the tube are thoroughly mixed. 
	&#8203;NOTE: Slide preparation can be done at the lab bench. Discard the toothpick in nonhazardous waste. Use a clean, new toothpick for each sample.
	<ol>
		<li>Using a clean disposable plastic transfer pipette, remove 1 drop of pollen residue from a tube, and place it in the center of its labeled microscope slide. Allow the drop to spread slightly. Place a clean coverslip over the drop on the slide.</li>
		<li>After the slide has dried, seal the coverslip to the slide with clear nail polish. Place a small drop of polish on each corner of the coverslip, and paint a border of polish around the perimeter of the coverslip where it meets the slide. Allow the nail polish to dry completely, and paint a second coat of polish around the perimeter of the coverslip.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Different pollen trap styles have their own advantages and consequences. The benefits and limitations of four commonly used trap styles, (1) front-mount, (2) bottom-mount, (3) auger-hole, and (4) top-mount pollen traps are discussed below. Front-mount traps are the most versatile style (Figure 1B). Installation is fast and easy; it can be done without lifting hive boxes, and these traps can fit on any Langstroth style of hive equipment. As the collection tray sits in front of the colony, it collects minimal debris from the colony. However, the collection tray is also more exposed to external elements-moisture from field irrigation, rainy or humid weather, or dew can come into contact with the pollen through the collection tray, potentially rendering the pollen unusable if the pellets become too saturated to separate. The risk of pollen saturation can be reduced by avoiding trapping during forecasted events of rain or high humidity. Placing a rubber mat under the trap and extra covering material (e.g., roofing felt) on top of the pollen trap can also shield the collection tray from weather.
Bottom-mount traps were used to collect pollen for the data in this paper (Figure 1A). They are not as convenient to install because they must be placed underneath the brood nest of the colony. Installation is time-consuming and results in a high volume of debris falling into the trap from the colony, such as bee parts and small bits of wax. The floor of the collection tray for most manufactured bottom-mount traps is made of fine mesh, which allows proper ventilation to protect the collected pollen from moisture. Auger-hole pollen traps help minimize disorientation of foragers if they primarily use auger holes as hive entrances instead of the entrance made by the bottom board of the hive (Figure 1C). As the collection tray for auger-hole pollen traps is very small, it must be emptied frequently to avoid overflowing of the collection tray. Given its upper placement on a hive, the top-mount pollen trap is the easiest trap style to install and remove, and the collected pollen sample is free of hive debris. However, this trap style is extra-sensitive to damaged hive equipment as the collection tray would be exposed to moisture if the lid, inner cover, and upper hive box are not properly sealed together.
The protocols described herein call for selecting colonies with large adult and larval populations (step 1.2). This selection method is intended to produce very large amounts of trapped pollen from these colonies. Colonies with substantial foraging populations may experience heavy congestion at the entrance upon trap installation. Selecting a large hive entrance will alleviate congestion. Large foraging populations may also collect very large quantities of pollen that can exceed the bounds of the collection tray. Use voluminous collection trays, as seen with most bottom- or top-mount trap styles, and empty trays frequently to accommodate large quantities of trapped pollen. If the desired research objective is to assess pollen quantities collected by colonies in an apiary, select representative colonies instead of optimizing the adult and larval populations for selection. All styles of pollen traps block the hive entrance and create a new entrance that differs spatially from the original entrance16. Pollen traps commonly fail to collect pollen when foragers are unable to reorient to the new entrance of the pollen trap upon installation. These foragers readily drift to neighboring hives, potentially cross-contaminating other pollen collection samples if they enter another hive with a pollen trap. Hence, foragers should be given at least 24 h to acclimate to the new entrance by keeping the trapping mechanism disengaged after installation. Selecting colonies with few or no additional hive entrances also reduces confusion when orienting to the new pollen trap entrance.
Additional hive entrances (e.g., holes and warped lids) should be sealed, but the risk of foragers drifting to neighboring hives will increase with these entrances present at the start of trap installation. Foragers will also readily drift into other hive entrances if a pollen trap is installed only on a single hive in a cluster of palletized hives. Foragers are less likely to drift if all the hives that are facing the same direction on the pallet have traps installed. Top-mount pollen traps may pose a higher risk of bee drift due to the substantial distance between the pollen trap entrance and the hive&#39;s original entrance. For this study, pollen traps were installed on multiple honey bee colonies in each experimental location to account for variation in the pollen quantity and taxa composition between each honey bee colony. Thus, pollen traps should be installed on multiple colonies to achieve robust pollen collections from the landscape because pollen collection can vary widely between colonies based on plant species type and total collected quantity12,13. Each pollen sample had a 7-day collection period. In future studies, collecting pollen in two or three consecutive 72 h intervals will increase the accuracy of pollen forage estimation40.
As there is a high degree of temporal fluctuation in pollen collection, pollen estimation accuracy could be increased by repeating the pollen collection process in early, peak, and late blooming periods of the targeted cropping systems24,27,39. Pollen should be collected from multiple locations, albeit the same cropping system or landscape type, because of anticipated variation in quantity and plant species type between apiary locations14,27,33,43. Long-term pollen trapping can be detrimental to honey bee colonies. Potential impacts include reduced brood rearing, shortening of larval growth period, and cannibalism of eggs and young larvae in the hives19,44,45,46. Longer periods of pollen trapping, such as the entire growing season, can worsen the harmful effects on brood rearing in colonies. Pollen trapping can also cause a reduction in honey production and an increase in the moisture level of stored honey13. Rotating pollen traps between colonies in an apiary when continuously monitoring a landscape or cropping system could mitigate harm to colonies used for pollen trapping. Engaging pollen traps every other week will reduce detrimental impacts, particularly loss in honey production, if trapping pollen on the same colonies throughout a period of time13.
In addition, the pollen traps are preferably placed on strong colonies. Occasionally, the pollen traps may engage unintentionally. This could be avoided by locking the pollen trap mechanism when pollen trap collection is not desired. Pollen traps do not remove all corbicular pollen from honey bee foragers. Trapping efficiency depends on trap type, pollen pellet size, bee body size, the time of day, and weather conditions. Hence, corbicular pollen collection is not consistent when using pollen traps for different plant species and collection periods25,26. Smaller pollen pellets from plants such as Eucalyptus spp. and Tamarix spp. are less likely to be captured by pollen traps27. Notably, no highbush blueberry (Vaccinium corymbosum L.) pollen was found from the highbush blueberry collection sites in this study, which supports previous evidence that highbush blueberry pollen pellets are too small for pollen trap collection47. In contrast, pollen sourced from dandelion (Taraxacum officinale F.H. Wigg) was found in every cropping system in this study. Pollen pellets of some plant species can also be much larger than others, such as Taraxacum spp., and could possibly be over-represented in the analysis of pollen collections from pollen traps27. Capturing individual pollen foragers and manually removing their corbicular pollen will increase the accuracy of a pollen source assessment, but it is very time- and resource-intensive compared to using pollen traps (Table 1). Sorting pollen pellets into color groups is relatively straight forward, although it is time-consuming. Unless there is a specific research goal or objective, the quantity of pollen pellets should be limited to 10 g or less (for any given sample) for sorting into color groups. Sorting entire samples that contain greater quantities than this amount will drastically increase the time required to complete the analysis. It is, however, crucial that a pollen sample is very well-mixed before a subsample for color sorting is taken from it. Failing to mix the original sample may result in a subsample that is not representative of the whole, which should be avoided.
If the original sample container does not contain enough free space to allow for thorough mixing of pollen pellets, placing the whole sample in a large plastic bag or a small paper bag should suffice, even for large samples. Hard plastic, lidded containers will also work. Mixing the sample should be done gently, so that pollen pellets are not squashed or otherwise destroyed. Unintentional bias could subconsciously persuade one to scoop out &#34;the pretty purple pellets&#34;, for example, when removing a subsample from the whole. Therefore, the sample&#39;s color composition should be obscured from view while scooping out a subsample. In this way, getting a subsample that is truly representative of the whole is more likely. However, this subsampling method could fail to select pollen pellets that are in low abundance in the sample. Therefore, if identifying every individual plant taxon represented in the sample is a research goal, collecting a subsample will not be appropriate; the whole sample must be analyzed. Hence, pellets should be sorted in a glass Petri dish. Once sorting is completed, appropriate pages of the Pantone color guide can be placed underneath the dish to make color-matching between the guide and the sorted pollen easier. An example of this is illustrated in Figure 5.
When trapping pollen from honey bee colonies placed in crops for pollination, no more than ten total color groups should be used: nine individual colors and one &#34;miscellaneous&#34; color group composed of the minority colors in the sample. Placing a reasonable limit on the maximum number of color groups that a sample can be divided into prevents the researcher from getting bogged down by endlessly separating pellets into ever-increasing numbers of extremely specific groups, which, when sorting is complete, may not individually contain sufficient quantities for acetolysis. If trapping from colonies that are likely foraging from a very diverse assortment of plant species, more color groups may be necessary, and the protocols should be optimized to reflect that requirement. The present study focused on the pollen samples collected from honey bee colonies pollinating crops, and multiple taxa were commonly found in a color group, similar to previous studies29,30,31.
Acetolysis dissolves the lipids, proteins, and organic debris from the surface of pollen grains, revealing the distinguishing characters of the exine, so that the grains can be stained and identified more easily. It is an old and common methodology used in many kinds of pollen research37. The general steps are standardized; they vary little from protocol to protocol. However, the specifics of centrifugation speeds and times, incubation temperature and duration, pollen-quantity-driven reagent volumes and even supernatant removal method (decanting vs pipetting) may need to be experimentally optimized according to research goals and, to some extent, the types of pollen likely to be encountered48. Indeed, acetolysis can remove important diagnostic characters of the pollen from some taxa such as Malvaceae and Orchidaceae38. Therefore, not all pollen is amenable to standard methods of acetolysis. As stated above, these methods were optimized in this study for the purpose of identifying dominant plant taxon sources of pollen collected by crop-pollinating honey bees. Details to be considered if precise quantification of pollen grains is part of the study, have not been addressed in this paper.
The use of solvents and acids requires careful planning, proper personal protective equipment (PPE), and responsible waste disposal (Figure 6). It is critical that researchers determine the correct way to store reagents and dispose of waste before beginning any part of acetolysis. In this laboratory, butyl gloves are used during any part of the process that involves sulfuric acid and even glacial acetic acid as they have far better degradation and permeation ratings for both acids than nitrile gloves, while not compromising dexterity49. It would be prudent to consult the respective institution&#39;s safety guidelines for recommendations on appropriate gloves and other PPE49. Glacial acetic acid addition before the acetolysis step helps remove any residual moisture in the sample and prepares it for the important acetolysis reaction. The glacial acetic acid-sulfuric acid mixture in the acetolysis step may react violently with water, which is why it is important that all glassware and supplies be completely dry, and that all moisture is removed from the sample before acetolysis. The post-acetolysis addition of glacial acetic acid dilutes and neutralizes the acetolysis mixture.
Ethanol and glacial acetic acid, in particular, can dissolve the ink of microcentrifuge tube labels, if these reagents drip on the outside of the tube, even with solvent-resistant pens. Check the tube labels frequently throughout the process to be sure they are still legible. If logistically feasible, consider using LaserJet-printed labels as a safeguard against this possibility. The manner in which supernatants are decanted will influence whether reagents dribble down the outside of the microcentrifuge tubes. It is important to decant the supernatant with a confident, smooth hand, which comes with practice. Care should be taken to avoid loss of pollen samples from the centrifuge tube during decantation. Decanting too fast risks losing some or all of the pollen residue; decanting too slowly may result in the supernatant running down the tube. Although an incubation temperature of 100 &#176;C is commonly recommended, pollen could easily become &#34;overcooked&#34; at that temperature in the quantities used in this study (0.25 g), particularly if incubated for slightly longer durations29. In fact, even at 80 &#176;C, pollen grains can burst or otherwise be damaged if left in the acetolysis mixture too long. Incubation temperature and duration must be carefully determined to avoid destroying the pollen grains in the sample.
Staining pollen increases the definition and contrast of the exine features, making it easier to photograph and identify (Figure 7). Five drops (from a plastic transfer pipette) of 1% Safranin O effectively stained 0.25 g of pollen. However, different pollens stain differently. If pollen grains are stained too lightly or too heavily, identification may be difficult. When possible, the volume of the stain solution needed to appropriately stain the pollen species expected to be found in the study should be validated before beginning the processing of the experimental samples. Nonetheless, if one of the experimental samples is not properly stained, it can be corrected. To lighten a pollen sample that is stained too heavily, rinse the sample with water and then ethanol. If the pollen is not stained well enough to see distinguishing features, a few additional drops of stain can be added. The stain of these samples should be checked before adding glycerin. Likewise, some trial and error may be needed to determine the ideal volume of glycerin for the pollen residues. Fifteen drops of glycerin suitably protected the samples in this study from drying out, while also diluting the pollen residue to a concentration ideal for downstream identification via light microscopy. Other quantities of pollen residue may require more or less glycerin to prevent desiccation and facilitate mounting.

The western honey bee (Apis mellifera L.) is an important pollinator of many agricultural crops that depend on bee pollination1. For more than a decade, significant honey bee colony losses have been reported2,3,4,5,6,7,8,9. Several factors-including parasites and diseases, poor nutrition and pesticides-have been implicated in these colony declines10. Poor nutrition can be attributed to agricultural intensification and loss of foraging habitat11. It is imperative to understand the floral resources utilized by bees in different landscapes to improve bee nutrition and aid in bee conservation efforts. Pollen is the primary source of protein, lipids, vitamins, and minerals for bees and has been used in many agricultural and ecological studies to understand colony-level foraging preferences of honey bees, evaluate the impact of pollen trapping on honey bee colonies, and determine pesticide exposure to bees12,13,14.
Honey bees gather pollen from flowers, pack pollen into pellets on their corbicula - a tibial pollen basket on their hindleg - and return to the colony for storage. Corbicular pollen can be removed from foragers by capturing them at the hive entrance or on flowers, chilling them briefly to immobilize them, and then removing the pollen pellets from their hindlegs with forceps. The laborious process of hand-collecting corbicular pollen from individually captured foragers is slow and inefficient if one requires a considerable quantity of pollen. A simpler and more efficient method of collecting large volumes of pollen is by trapping corbicular pollen pellets from honey bees at hive entrances. Pollen traps are designed to dislodge corbicular pollen from the legs of the returning pollen foragers as they enter the hive15. The foragers must squeeze through mesh holes that are sized to narrowly allow the passage of a honey bee worker body.
As the honey bee passes through one of these holes, the larger pollen pellets are scraped off her legs and fall into a collection tray16. Studies have shown that pollen trapping stimulates foragers to collect more pollen, thus increasing the pollination efficiency of the surrounding crops and vegetation17,18,19,20. Pollen collection methodologies can also be used to understand the forage utilized by honey bees in the landscape as the first step to determining the quantity, quality, and taxa of flowering plant species. Effective pollen-trapping methodologies thus facilitate both pollination and honey bee nutrition research. A comparison of these pollen collection methods is illustrated in Table 1. Pollen foraging behavior will change based on the colony&#39;s need for stored pollen relative to its egg and larvae population levels21,22. As these changes include varying collection intensity, high variation in pollen quantity is often expected among colonies in the same location and among different locations of the same cropping system or landscape type23,24. Increasing the number of colonies and locations to trap pollen will help accommodate this variation.
Pollen traps vary in efficiency25,26. The size of pollen pellets collected by honey bees varies among plant species and can change based on the levels of pollen stores in the colony27,28. This poses the potential for smaller pollen pellets to be underrepresented and larger pellets to be overrepresented in samples collected via pollen traps. Adult bees vary in body size, which can also affect the representation of pollen collected in traps. There are also plant species that predominantly produce nectar that will be undetected if only assessing collected pollen in some landscapes. Trapping efficiency is also impacted by forager drift and disorientation, which is influenced by pollen trap type and the condition of the hive equipment. This problem can be mitigated by employing techniques specified in this paper. Investigators may consider additional research techniques, such as counting flower visitation by foragers, to supplement the results of colony-level foraging preferences. A useful method for assessing pollen diversity is sorting corbicular pollen by color. Although honey bees are generalist foragers, they also exhibit flower fidelity, where they collect pollen from the same plant species in the same location during any given collection trip. Based on this foraging behavior, it is assumed that any given corbicular pollen pellet is predominantly represented by a single plant species 27,29,30,31. Hence, scientists can describe pollen diversity by sorting corbicular pollen by pellet color and reporting the total number of colors detected or the proportion of the total represented by each color group12,32,33,34. This can be accomplished by measuring the mass or pellet count of each color group. Measuring the pellet count of each color group is suggested if there are known or suspected systematic differences in the weight of pellets from different taxa. Systematic differences could be caused by pellet size or the quantity of nectar that foragers add to pollen when forming a pellet.
Color sorting is a time-efficient and simple process, but may not have acceptable accuracy for some pollination research studies because different plant taxa may have similar pollen pellet colors35,36. Additionally, there is a logistical limit to the number of distinct color groups pollen pellets can be separated into. Thus, the separation of each individual plant taxon pollen into its own distinct pellet color group may not always be possible in pollination studies. Morphological characterization of pollen grains via light microscopy often complements color separation of pellets by distinguishing the pollen of two or more taxa in pellets of the same color group. Although, it is common to find pollen grains of multiple taxa in a given pollen pellet color group, individual pollen pellets collected by a honey bee generally comprise one predominant taxon, possibly with other taxa in minor quantities. Thus, it is common to assume taxonomic fidelity in corbicular pollen pellets of honey bees. Pollen pellets from other pollinators that do not exhibit flower fidelity behavior, such as bumble bees, will often contain many plant species and may not possess a predominant taxon. In instances where quantitative estimates of taxa proportions in polyfloral pollen pellets are desired, microscopic methods that include acetolysis, are additionally required for proper analysis.
Assessing morphological characteristics of acetolyzed pollen grains is the most common method for taxonomic identification16. The acetolysis procedure removes the pollen grain&#39;s protoplasm to expose diagnostic characteristics that can be observed under light microscopy37,38. Using this method, researchers can report different taxa, frequency of taxa found in specific cropping systems, and predominant taxa of pellet colors33,36. Acetolysis is the best analytical technique for revealing pollen morphology28. However, some acetolyzed pollen grains, such as many Rosaceae types, cannot be identified to genus or species level through acetolysis and light microscopy alone. Researchers consider scanning electron microscopy or metabarcoding as alternative methods to achieve genus- or species-level identification. These alternative methods, however, only provide qualitative taxon identification and fail to estimate the proportions of different pollen grain taxa in polyfloral pollen pellets36,39. In addition, the expense and necessary expertise is considerably higher for these methods. A comparison of these identification methods is illustrated in Table 1.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Methods</td>
			<td>Time</td>
			<td>Expense</td>
			<td>Resolution</td>
			<td>Expertise</td>
		</tr>
		<tr>
			<td>Pollen Collection</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pollen trapping</td>
			<td>Low</td>
			<td>Moderate</td>
			<td>Variable</td>
			<td>Moderate</td>
		</tr>
		<tr>
			<td>Pollen forager collection</td>
			<td>High</td>
			<td>Moderate</td>
			<td>High</td>
			<td>Low</td>
		</tr>
		<tr>
			<td>Pollen Identification</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Visual (color sorting only)</td>
			<td>Moderate</td>
			<td>Low</td>
			<td>Low</td>
			<td>Low</td>
		</tr>
		<tr>
			<td>Acetolysis</td>
			<td>Moderate</td>
			<td>Moderate</td>
			<td>Moderate</td>
			<td>Moderate</td>
		</tr>
		<tr>
			<td>Scanning electron microscopy</td>
			<td>High</td>
			<td>High</td>
			<td>High</td>
			<td>High</td>
		</tr>
		<tr>
			<td>Metabarcoding</td>
			<td>Variable</td>
			<td>High</td>
			<td>High</td>
			<td>High</td>
		</tr>
	</tbody>
</table>

Table 1: Comparison of different methods of pollen collection and identification based on time, expense, resolution, and expertise.&#160;Visual methods (color sorting only) report the total number of colors detected or the proportion of the total represented by each color group as a metric for determining pollen sources, but do not provide taxon identification.
The information available on trapping and sorting pollen and acetolyzing pollen grains is diverse and often spread across multiple sources, varying for researchers in different fields. This paper offers detailed insights into different types of pollen traps that can be used by both researchers and beekeepers to effectively collect large volumes of corbicular pollen. Also provided are protocols for preparing pollen samples - by acetolysis, staining, and slide mounting - for plant taxa identification. The methodologies detailed here are comprehensive and serve as a unique resource for identifying predominant plant species on which honey bees forage in a given landscape, particularly in cropping systems. Findings based on these methods from a previous study have been presented and document the diversity of pollen pellet colors and plant taxa from corbicular pollen collected by honey bees in five cropping systems14.

<table><tbody><tr><td>#8 hardware cloth</td><td></td><td></td><td>2.7 mm aperture</td></tr><tr><td>10 mL graduated cylinder</td><td></td><td></td><td></td></tr><tr><td>1000 uL micropipette tips</td><td></td><td></td><td></td></tr><tr><td>1250 mL filter micropipette tips</td><td></td><td></td><td></td></tr><tr><td>15 x 75 mm glass slides</td><td></td><td></td><td>Thickness: 0.93 mm - 1.05 mm</td></tr><tr><td>2 mL microcentrifuge tubes</td><td></td><td></td><td></td></tr><tr><td>250 mL graduated Borosilicate glass beakers (x3)</td><td>VWR</td><td>10754-952</td><td></td></tr><tr><td>50 mL graduated Borosilicate glass beakers (x6)</td><td>VWR</td><td>10754-946</td><td></td></tr><tr><td>95% EtOH</td><td>Pharmco AAPER</td><td>111000200DM55</td><td>ACS/USP/Kosher grade</td></tr><tr><td>Butyl vinyl gloves</td><td></td><td></td><td></td></tr><tr><td>Centrifuge</td><td></td><td></td><td>1060 x g maximum speed; horizontal swing preferred</td></tr><tr><td>Chemical safety goggles</td><td></td><td></td><td></td></tr><tr><td>Color guide</td><td>Pantone</td><td>SKU: GP1601A</td><td>Solid coated</td></tr><tr><td>Coverslip of 1 or 1.5</td><td></td><td></td><td>Thickness: 0.13 mm - 0.19 mm</td></tr><tr><td>Distilled water</td><td></td><td></td><td></td></tr><tr><td>Forceps</td><td></td><td></td><td></td></tr><tr><td>Fume hood</td><td></td><td></td><td></td></tr><tr><td>Glacial acetic acid</td><td>BDH Chemicals</td><td>BDH3092</td><td>ACS grade</td></tr><tr><td>Glass funnel</td><td></td><td></td><td></td></tr><tr><td>Glycerin</td><td>Humco</td><td>103196001_1</td><td>USP grade, 99.5%, anhydrous</td></tr><tr><td>Hazardous waste containers</td><td></td><td></td><td></td></tr><tr><td>Hive tool</td><td></td><td></td><td></td></tr><tr><td>Hot block</td><td></td><td></td><td>Must reach 80 degree C</td></tr><tr><td>Lab coat with long sleeves</td><td></td><td></td><td></td></tr><tr><td>Latex or polyurethane foam</td><td></td><td></td><td></td></tr><tr><td>Microscope</td><td></td><td></td><td></td></tr><tr><td>Nailpolish, clear</td><td></td><td></td><td></td></tr><tr><td>Nitrile gloves</td><td></td><td></td><td></td></tr><tr><td>P1000 pipette</td><td>VWR</td><td></td><td></td></tr><tr><td>Petri dish</td><td></td><td></td><td></td></tr><tr><td>Plastic spoon</td><td></td><td></td><td></td></tr><tr><td>Safranin</td><td>Ward&#039;s Science</td><td>470302-322</td><td>Lab grade</td></tr><tr><td>Smoker</td><td></td><td></td><td>For pollen trap installation</td></tr><tr><td>Sodium bicarbonate</td><td>EMD Millipore</td><td>SX0320</td><td>ACS grade, powder</td></tr><tr><td>Squirt bottles (x2)</td><td></td><td></td><td></td></tr><tr><td>Sulfuric acid</td><td>EMD Millipore</td><td>SX1244</td><td>ACS grade</td></tr><tr><td>Sundance Bottom Mount Pollen Trap</td><td>Betterbee Beekeeping Supply</td><td>PTRAPB</td><td></td></tr><tr><td>Tape</td><td></td><td></td><td></td></tr><tr><td>Wooden stir sticks</td><td></td><td></td><td></td></tr><tr><td>Wooden tooth picks</td><td></td><td></td><td></td></tr></tbody></table>

collection and identification of pollen from honey bee colonies

Researchers often collect and analyze corbicular pollen from honey bees to identify the plant sources on which they forage for pollen or to estimate pesticide exposure of bees via pollen. Described herein is an effective pollen-trapping method for collecting corbicular pollen from honey bees returning to their hives. This collection method results in large quantities of corbicular pollen that can be used for research purposes. Honey bees collect pollen from many plant species, but typically visit one species during each collection trip. Therefore, each corbicular pollen pellet predominantly represents one plant species, and each pollen pellet can be described by color. This allows the sorting of samples of corbicular pollen by color to segregate plant sources. Researchers can further classify corbicular pollen by analyzing the morphology of acetolyzed pollen grains for taxonomic identification. These methods are commonly used in studies related to pollinators such as pollination efficiency, pollinator foraging dynamics, diet quality, and diversity. Detailed methodologies are presented for collecting corbicular pollen using pollen traps, sorting pollen by color, and acetolyzing pollen grains. Also presented are results pertaining to the frequency of pellet colors and taxa of corbicular pollen collected from honey bees in five different cropping systems.

A previous study reported the assessment of the diversity of pollen collected by honey bees in the following agricultural crops: almond, cherry, highbush blueberry, hybrid carrot, and meadowfoam14. Using the described methods, corbicular pollen was collected, sorted by color, and the plant sources of each pellet color group identified to assess pollen diversity. Bottom-mount pollen traps were installed on colonies in multiple sites for each crop (Figure 1A). The quantity of pollen collected from each site was enough to fulfill sample weight requirements of the color sorting and acetolysis analysis methods. Every pollen collection sample had multiple distinguishable color groups (Figure 2 and Figure 3). In some samples, pollen color groups contained as few as 4-5 pellets; however, most groups had significantly more than that and thus served as their own labeled color group for acetolysis (Figure 4 and Figure 5). After acetolysis (Figure 6), bright-field light microscopy was used to effectively identify each color group to its lowest possible taxonomic rank by confirming the morphological characteristics with those of voucher specimens collected from the area surrounding each study site (Figure 7). 
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62064/62064fig01.jpg" /> 
Figure 1: Pollen traps installed on a honey bee colony to collect corbicular pollen. (A)&#160;Bottom-mount traps placed above the hive bottom board and directly above the lowest hive box. Other pollen trap styles include (B) front-mount and (C) auger-hole entrance mount traps. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62064/62064fig02.jpg" /> 
Figure 2: Trapping mechanism and collection tray of pollen trap. Returning pollen foragers must squeeze through the mesh trapping mechanism before reaching their hive. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62064/62064fig03.jpg" /> 
Figure 3: Collection tray of pollen trap. Corbicular pollen is scraped off the legs of returning pollen foragers by the pollen trap and falls into the collection tray. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62064/62064fig04.jpg" /> 
Figure 4: Sorting a sample of corbicular pollen into color groups. Corbicular pollen can be dried and weighed after it is sorted into color groups to report proportions of different color pellets collected. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62064/62064fig05.jpg" /> 
Figure 5: Four groups of pollen pellets sorted by color using the Pantone color guide. The color groups are labeled as (A)&#160;gray, Pantone 5855C, (B)&#160;brown, Pantone 7557C, (C) yellow, Pantone 458C, and (D) light brown, Pantone 3547C. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62064/62064fig06.jpg" /> 
Figure 6: Acetolysis equipment setup inside the fume hood. The heat block, reagents, solvent waste and acid waste containers, labeled beakers, pipette, pipette tips, stir sticks, and microcentrifuge tubes situated inside the fume hood. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62064/62064fig07.jpg" /> 
Figure 7: Micrograph of stained, acetolyzed pollen grains. Many facets of acetolyzed mustard (Brassicaceae) pollen grains at 40x magnification. Scale bar = 50.398 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/62064/62064fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The pollen collected from almond crop locations had relatively lower pollen diversity than pollen collected from other crops, with an average of 3.0 &#177; 0.5 pellet colors and 3.2 &#177; 1.2 plant taxa per site (Table 2)14. The remaining four cropping systems had higher pollen diversity levels with an average of 6.0 &#177; 2.0 pellet colors and 8.0 &#177; 1.5 plant taxa per site in cherry, 8.8 &#177; 1.4 pellet colors and 13.5 &#177; 2.0 plant taxa per site in highbush blueberry, 7.0 &#177; 1.0 pellet colors and 11.0 &#177; 0.0 plant taxa per site in hybrid carrot, and 10.0 &#177; 0.0 pellet colors and 13.0 &#177; 1.5 plant taxa per site in meadowfoam14.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Crop</td>
			<td>Mean number of pollen pellet colors/site (SE)</td>
			<td>Mean number of plant taxa/site (SE)</td>
			<td colspan="3">Total taxa identified</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Family</td>
			<td>Genus</td>
			<td>Species</td>
		</tr>
		<tr>
			<td>Almond</td>
			<td>3.0 (0.5)</td>
			<td>3.2 (1.2)</td>
			<td>4</td>
			<td>3</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Blueberry</td>
			<td>8.8 (1.4)</td>
			<td>13.5 (2.0)</td>
			<td>5</td>
			<td>10</td>
			<td>13</td>
		</tr>
		<tr>
			<td>Carrot</td>
			<td>7.0 (1.0)</td>
			<td>11.0 (0.0)</td>
			<td>3</td>
			<td>5</td>
			<td>6</td>
		</tr>
		<tr>
			<td>Cherry</td>
			<td>6.0 (2.0)</td>
			<td>8.0 (1.5)</td>
			<td>4</td>
			<td>7</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Meadowfoam</td>
			<td>10.0 (0.0)</td>
			<td>13.0 (1.5)</td>
			<td>5</td>
			<td>4</td>
			<td>14</td>
		</tr>
	</tbody>
</table>

Table 2: Diversity of corbicular pollen collected from honey bees in five cropping systems. Diversity metrics include mean number of pellet colors (&#177; SE), mean number of plant taxa (&#177; SE), and total taxa identified.&#160;This table has been modified from14. Abbreviation: SE = standard error.

Watch this Scientific Journal Video about Collection and Identification of Pollen from Honey Bee Colonies at JoVE.com

Collection and Identification of Pollen from Honey Bee Colonies

We describe methods for collecting corbicular pollen from honey bees as well as protocols for color sorting, acetolysis, and microscope slide preparation of pollen for taxonomic identification. In addition, we present pellet color and taxonomic diversity of corbicular pollen collected from five cropping systems using pollen traps.

Researchers often collect and analyze corbicular pollen from honey bees to identify the plant sources on which they forage for pollen or to estimate ...

collection-and-identification-of-pollen-from-honey-bee-colonies

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

7.0K Views.  Oregon State University. We describe methods for collecting corbicular pollen from honey bees as well as protocols for color sorting, acetolysis, and microscope slide preparation of pollen for taxonomic identification. In addition, we present pellet color and taxonomic diversity of corbicular pollen collected from five cropping systems using pollen traps.

Video: Collection and Identification of Pollen from Honey Bee Colonies

Behavioural Pharmacology in Classical Conditioning of the Proboscis Extension Response in Honeybees (Apis mellifera)

Honeybees (Apis mellifera) are well known for their communication and orientation skills and for their impressive learning capability1,2. Because the survival of a honeybee colony depends on the exploitation of food sources, forager bees learn and memorize variable flower sites as well as their profitability. Forager bees can be easily trained in natural settings where they forage at a feeding site and learn the related signals such as odor or color. Appetitive associative learning can also be studied under controlled conditions in the laboratory by conditioning the proboscis extension response (PER) of individually harnessed honeybees3,4. This learning paradigm enables the study of the neuronal and molecular mechanisms that underlie learning and memory formation in a simple and highly reliable way5-12. A behavioral pharmacology approach is used to study molecular mechanisms. Drugs are injected systemically to interfere with the function of specific molecules during or after learning and memory formation13-16.
Here we demonstrate how to train harnessed honeybees in PER conditioning and how to apply drugs systemically by injection into the bee flight muscle.

Behavioural Pharmacology in Classical Conditioning of the Proboscis Extension Response in Honeybees Apis mellifera

We demonstrate how to implement a behavioral pharmacology method in an appetitive olfactory conditioning paradigm in honeybees (Apis mellifera) by systemic application of drugs. This method allows investigation of the mechanisms underlying learning and memory formation in a simple and reliable way.

Honeybees (Apis mellifera) are well known for their communication and orientation skills and for their impressive learning capability1,2. Because the ...

Neuroscience

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

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered &#34;bee-safe,&#34; 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.

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.

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered ...

Environment

Evaluating the Effect of Environmental Chemicals on Honey Bee Development from the Individual to Colony Level

The presence of pesticides in the beekeeping environment is one of the most serious problems that impacts the life of a honey bee. Pesticides can be brought back to the beehive after the bees have foraged on flowers that have been sprayed with pesticides. Pesticide contaminated food can be exchanged between workers which then feed larvae and therefore can potentially affect the development of honey bees. Thus, residual pesticides in the environment can become a chronic damaging factor to honey bee populations and gradually lead to colony collapse. In the presented protocol, honey bee feeding methods are described and applied to either an individual honey bee or to a colony. Here, the insect growth regulator (IGR) pyriproxyfen (PPN), which is widely used to control pest insects and is harmful to the development of honey bee larvae and pupae, is used as the pesticide. The presenting procedure can be applied to other potentially harmful chemicals or honeybee pathogens for further studies.

Herein we present a method to feed pesticide contaminated food to both an individual honey bee and a beehive colony. The procedure evaluates the pesticide effect on individual honey bees by in vivo feeding of basic larval diet and also on the natural condition of beehive colony.

The presence of pesticides in the beekeeping environment is one of the most serious problems that impacts the life of a honey bee. Pesticides can be ...

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Although solitary bees provide crucial pollination services for wild and managed crops, this species-rich group has been largely overlooked in pesticide regulation studies. The risk of exposure to fungicide residues is likely to be especially high if the spray occurs on, or near host plants while the bees are collecting pollen to provision their nests. For species of Osmia that consume pollen from a select group of plants (oligolecty), the inability to use pollen from non-host plants can increase their risk factor for fungicide-related toxicity. This manuscript describes protocols used to successfully rear oligolectic mason bees, Osmia ribifloris sensu lato, from egg to prepupal stage within cell culture plates under standardized laboratory conditions. The in vitro-reared bees are subsequently used to investigate the effects of fungicide exposure and pollen source on bee fitness. Based on a 2 &#215; 2 fully crossed factorial design, the experiment examines the main and interactive effects of fungicide exposure and pollen source on larval fitness, quantified by prepupal biomass, larval developmental time, and survivorship. A major advantage of this technique is that using in vitro-reared bees reduces natural background variability and allows the simultaneous manipulation of multiple experimental parameters. The described protocol presents a versatile tool for hypotheses testing involving the suite of factors affecting bee health. For conservation efforts to be met with significant, lasting success, such insights into the complex interplay of physiological and environmental factors driving bee declines will prove to be critical.

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Fungicide sprays on flowering plants may expose solitary bees to high concentrations of pollen-borne fungicide residues. Using laboratory-based experiments involving in vitro-reared bee larvae, this study investigates the interactive effects of consuming fungicide-treated pollen derived from host and non-host plants.

Although solitary bees provide crucial pollination services for wild and managed crops, this species-rich group has been largely overlooked in pesticide ...

Tactile Conditioning And Movement Analysis Of Antennal Sampling Strategies In Honey Bees (Apis mellifera L.)

Honey bees (Apis mellifera L.) are eusocial insects and well known for their complex division of labor and associative learning capability1, 2. The worker bees spend the first half of their life inside the dark hive, where they are nursing the larvae or building the regular hexagonal combs for food (e.g. pollen or nectar) and brood3. The antennae are extraordinary multisensory feelers and play a pivotal role in various tactile mediated tasks4, including hive building5 and pattern recognition6. Later in life, each single bee leaves the hive to forage for food. Then a bee has to learn to discriminate profitable food sources, memorize their location, and communicate it to its nest mates7. Bees use different floral signals like colors or odors7, 8, but also tactile cues from the petal surface9 to form multisensory memories of the food source. Under laboratory conditions, bees can be trained in an appetitive learning paradigm to discriminate tactile object features, such as edges or grooves with their antennae10, 11, 12, 13. This learning paradigm is closely related to the classical olfactory conditioning of the proboscis extension response (PER) in harnessed bees14. The advantage of the tactile learning paradigm in the laboratory is the possibility of combining behavioral experiments on learning with various physiological measurements, including the analysis of the antennal movement pattern.

Tactile Conditioning And Movement Analysis Of Antennal Sampling Strategies In Honey Bees Apis mellifera L.

In this protocol we show how to condition harnessed honey bees to tactile stimuli and introduce a 2D motion capture technique for analyzing the kinematics of fine-scale antennal sampling pattern.

Honey bees (Apis mellifera L.) are eusocial insects and well known for their complex division of labor and associative learning capability1, 2. The worker ...

Preparation of Single-cohort Colonies and Hormone Treatment of Worker Honeybees to Analyze Physiology Associated with Role and/or Endocrine System

Honeybee workers are engaged in various tasks related to maintaining colony activity. The tasks of the workers change according to their age (age-related division of labor). Young workers are engaged in nursing the brood (nurse bees), while older workers are engaged in foraging for nectar and pollen (foragers). The physiology of the workers changes in association with this role shift. For example, the main function of the hypopharyngeal glands (HPGs) changes from the secretion of major royal jelly proteins (MRJPs) to the secretion of carbohydrate-metabolizing enzymes. Because worker tasks change as the workers age in typical colonies, it is difficult to discriminate the physiological changes that occur with aging from those that occur with the role shift. To study the physiological changes in worker tissues, including the HPGs, in association with the role shift, it would be useful to manipulate the honeybee colony population by preparing single-cohort colonies in which workers of almost the same age perform different tasks. Here we describe a detailed protocol for preparing single-cohort colonies for this analysis. Six to eight days after single-cohort colony preparation, precocious foragers that perform foraging tasks earlier than usual appear in the colony. Representative results indicated role-associated changes in HPG gene expression, suggesting role-associated HPG function. In addition to manipulating the colony population, analysis of the endocrine system is important for investigating role-associated physiology. Here, we also describe a detailed protocol for treating workers with 20-hydroxyecdysone (20E), an active form of ecdysone, and methoprene, a juvenile hormone analogue. The survival rate of treated bees was sufficient to examine gene expression in the HPGs. Gene expression changes were observed in response to 20E- and/or methoprene-treatment, suggesting that hormone treatments induce physiological changes of the HPGs. The protocol for hormone treatment described here is appropriate for examining hormonal effects on worker physiology.

Here we describe our detailed protocol for the preparation of single-cohort honeybee colonies&#160;&#8211; a useful tool for analyzing the role-associated worker physiology. We also describe detailed protocols for treating workers with juvenile hormone and ecdysone to evaluate the involvement of these hormones in the regulation of worker behavior and/or physiology.

Honeybee workers are engaged in various tasks related to maintaining colony activity. The tasks of the workers change according to their age (age-related ...

Monitoring Colony-level Effects of Sublethal Pesticide Exposure on Honey Bees

The effects of sublethal pesticide exposure to honey bee colonies may be significant but difficult to detect in the field using standard visual assessment methods. Here we describe methods to measure the quantities of adult bees, brood, and food resources by weighing hives and hive parts, by photographing frames, and by installing hives on scales and with internal sensors. Data from these periodic evaluations are then combined with running average and daily detrended data on hive weight and internal hive temperature. The resulting datasets have been used to detect colony-level effects of imidacloprid applied in a sugar syrup as low as 5 parts per billion. The methods are objective, require little training, and provide permanent records in the form of sensor output and photographs.

Sublethal doses of pesticides may affect colonies in ways that are difficult to detect using only periodic or visual methods. Methods to measure total adult bee mass, brood, and food resources by weighing hives and hive parts, photographing frames, and installing sensors, are provided. Data analysis is also addressed.

The effects of sublethal pesticide exposure to honey bee colonies may be significant but difficult to detect in the field using standard visual assessment ...

A 3D Printed Pollen Trap for Bumble Bee (Bombus) Hive Entrances

To verify the plant sources from which bumble bees forage for pollen, individuals must be collected to remove their corbicular pollen loads for analysis. This has traditionally been done by netting foragers at nest entrances or on flowers, chilling the bees on ice, and then removing the pollen loads from the corbiculae with forceps or a brush. This method is time and labor intensive, may alter normal foraging behavior, and can result in stinging incidents for the worker performing the task. Pollen traps, such as those used on honey bee hives, collect pollen by dislodging corbicular pollen loads from the legs of workers as they pass through screens at the nest entrance. Traps can remove a large quantity of pollen from returning forager bees with minimal labor, yet to date no such trap is available for use with bumble bee colonies. Workers within a bumble bee colony can vary in size making size selection of entrances difficult to adapt this mechanism to commercially reared bumble bee hives. Using 3D printing design programs, we created a pollen trap that successfully removes the corbicular pollen loads from the legs of returning bumble bee foragers. This method significantly reduces the amount of time required by researchers to collect pollen from bumble bee foragers returning to the colony. We present the design, results of pollen removal efficiency tests, and suggest areas of modifications for investigators to adapt traps to a variety of bumble bee species or nest box designs.

A 3D Printed Pollen Trap for Bumble Bee Bombus Hive Entrances

We present a non-lethal and automated mechanism to collect pollen from bumble bee (Bombus) workers returning to a hive. Instructions for producing, preparing, installing and using the devices are included. By using 3D-printed objects, modification to the design was timely, efficient and allowed for quick turnaround for testing.

To verify the plant sources from which bumble bees forage for pollen, individuals must be collected to remove their corbicular pollen loads for analysis.  ...

Preparation of Virus-Enriched Inoculum for Oral Infection of Honey Bees (Apis mellifera)

Honey bees are of great ecological and agricultural importance around the world but are also subject to a variety of pressures that negatively affect bee health, including exposure to viral pathogens. Such viruses can cause a wide variety of devastating effects and can often be challenging to study due to multiple factors that make it difficult to separate the effects of experimental treatments from preexisting background infection. Here we present a method to mass produce large quantities of virus particles along with a high throughput bioassay to test viral infection and effects. Necessitated by the current lack of a continuous, virus-free honey bee cell line, viral particles are amplified in vivo using honey bee pupae, which are extracted from the hive in large volumes using minimally stressful methodology. These virus particles can then be used in honey bee cage bioassays to test inocula viability, as well as various other virus infection dynamics, including interactions with nutrition, pesticides, and other pathogens. A major advantage of using such particles is that it greatly reduces the chances of introducing unknown variables in subsequent experimentation when compared to current alternatives, such as infection via infected bee hemolymph or homogenate, though care should still be taken when sourcing the bees, to minimize background virus contamination. The cage assays are not a substitute for large-scale, field-realistic experiments testing virus infection effects at a colony level, but instead function as a method to establish baseline viral responses that, in combination with the semi-pure virus particles, can serve as important tools to examine various dimensions of honey bee-virus physiological interactions.

Preparation of Virus-Enriched Inoculum for Oral Infection of Honey Bees Apis mellifera

Here we describe two protocols: first to propagate, extract, purify, and quantify large quantities of honey bee non-enveloped virus particles, including a method for removing honey bee pupae and second to test the effects of viral infection using a highly repeatable, high-throughput cage bioassay.

Honey bees are of great ecological and agricultural importance around the world but are also subject to a variety of pressures that negatively affect bee ...

An Improved Method to Measure Pollen Hydration Profiles in Arabidopsis thaliana

A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana

Sexual reproduction in flowering plants requires initial interaction between the pollen grain and the stigmatic surface, where a molecular dialog is established between the interacting partners. Studies across a range of species have revealed that a series of molecular checkpoints regulate the pollen-stigma interaction to ensure that only compatible, generally intraspecific pollen is successful in effecting fertilization. In species that possess a 'dry stigma', such as the model plant Arabidopsis thaliana, the first post-pollination, prezygotic compatibility checkpoint is the establishment of pollen hydration. 
This phase of pollination is tightly regulated, whereby signals from the pollen grain elicit the release of water from the stigma, thus permitting pollen hydration. The ability to accurately measure and track pollen hydration over time is key to the design of experiments directed at understanding the regulation of this critical step in reproduction. Published protocols frequently utilize flowers that have been excised from the parent plant, maintained on liquid or solid media, and bulk pollinated. 
This paper describes a noninvasive, in vivo pollination bioassay that permits minute-by-minute hydration tracking of individual A. thaliana pollen grains at high resolution. The assay is highly reproducible, able to detect very subtle variations of pollen hydration profiles, and thus is suitable for the analysis of mutants that affect pathways regulating pollination. Although the protocol is lengthier than those described for bulk pollinations, the precision and reproducibility it provides, along with its in vivo nature, make it ideal for the detailed dissection of pollination phenotypes.

Author Spotlight: A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana  

An improved method to measure pollen hydration profiles in Arabidopsis thaliana is described here. The new method offers higher resolution, is noninvasive, and is highly reproducible. The protocol represents a new tool for a finer dissection of the processes that regulate the early stages of pollination.

Sexual reproduction in flowering plants requires initial interaction between the pollen grain and the stigmatic surface, where a molecular dialog is ...

Collection and Identification of Pollen from Honey Bee Colonies

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Behavioural Pharmacology in Classical Conditioning of the Proboscis Extension Response in Honeybees (Apis mellifera)

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

Evaluating the Effect of Environmental Chemicals on Honey Bee Development from the Individual to Colony Level

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Tactile Conditioning And Movement Analysis Of Antennal Sampling Strategies In Honey Bees (Apis mellifera L.)

Preparation of Single-cohort Colonies and Hormone Treatment of Worker Honeybees to Analyze Physiology Associated with Role and/or Endocrine System

Monitoring Colony-level Effects of Sublethal Pesticide Exposure on Honey Bees

A 3D Printed Pollen Trap for Bumble Bee (Bombus) Hive Entrances

Preparation of Virus-Enriched Inoculum for Oral Infection of Honey Bees (Apis mellifera)

A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana